Germline Stem Cells [2 ed.] 1493940155, 9781493940158

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Methods in Molecular Biology 1463

Michael Buszczak Editor

Germline Stem Cells Second Edition

METHODS

IN

MOLECULAR BIOLOGY

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

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

Germline Stem Cells Second Edition

Edited by

Michael Buszczak Department of Molecular Biology University of Texas Southwestern Medical Center Dallas, TX, USA

Editor Michael Buszczak Department of Molecular Biology University of Texas Southwestern Medical Center Dallas, TX, USA

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-4939-4015-8 ISBN 978-1-4939-4017-2 (eBook) DOI 10.1007/978-1-4939-4017-2 Library of Congress Control Number: 2016945156 © Springer Science+Business Media New York 2008, 2017 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, express or implied, with respect to the material contained herein or for any errors or omissions that may have been made. Printed on acid-free paper This Humana Press imprint is published by Springer Nature The registered company is Springer Science+Business Media LLC New York

Preface Adult stem cells maintain tissue homeostasis by producing progeny that replace cells lost through the course of normal cellular turnover or because of injury. Germline stem cells are unique amongst stem cells because they produce daughter cells that develop into gametes, which ultimately serve to give rise to the next generation. Thus, germline stem cells are essential for the continued propagation of many sexually reproducing organisms. Germ cells are imbued with several unique properties. They are the only cells to undergo meiosis. Germ cells across species express many of the same markers and experience extensive epigenetic reprogramming. These cells have specialized mechanisms that help maintain the integrity of the genome and prevent the integration of selfish DNA elements. Several somatic tumors express genes normally specific to germ cells and germ cells themselves can give rise to specific types of cancer. Therefore, a better understanding of germ cells will have a broad impact across many fields of biology. Germline stem cell biology has witnessed an explosion of new findings and techniques since that last edition of Germline Stem Cells in the Methods in Molecular Biology series. The optimization of live-cell imaging techniques, improved cell purification protocols and the identification of germ cells in a number of genetically tractable organisms now allows for the characterization of germ cells at a greater depth and higher resolution than previously attainable. This new edition of Germline Stem Cells is intended to provide researchers with selected genetic, molecular, biochemical, and cell biological techniques used in germ cell research. While the focus here is on primordial germ cells and germline stem cells, many of the techniques and principles presented in the chapters of this issue may be applicable to many different types of adult stem cells. I would like to thank Prof. John M. Walker and the staff at Springer for their invitation, assistance, and patience during the preparation of this book. I would like to thank Nevine Shalaby for her help throughout this entire process. I would also like to express my sincere appreciation and gratitude to the various contributors for sharing their insights and expertise with the germline stem cell research community. Dallas, TX, USA

Michael Buszczak

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

v ix

1 Analysis of the C. elegans Germline Stem Cell Pool. . . . . . . . . . . . . . . . . . . . . . . . . . Sarah L. Crittenden, Hannah S. Seidel, and Judith Kimble 2 Methods for Studying the Germline of the Human Parasite Schistosoma mansoni . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Julie N.R. Collins and James J. Collins III 3 Evaluation of the Asymmetric Division of Drosophila Male Germline Stem Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mayu Inaba and Yukiko M. Yamashita 4 Live Imaging of the Drosophila Testis Stem Cell Niche . . . . . . . . . . . . . . . . . . . . . . Leah J. Greenspan and Erika L. Matunis 5 RNA Isolation from Early Drosophila Larval Ovaries . . . . . . . . . . . . . . . . . . . . . . . . Dana Gancz and Lilach Gilboa 6 Live-Cell Imaging of the Adult Drosophila Ovary Using Confocal Microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Nevine A. Shalaby and Michael Buszczak 7 Measurement of mRNA Poly(A) Tail Lengths in Drosophila Female Germ Cells and Germ-Line Stem Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Aymeric Chartier, Willy Joly, and Martine Simonelig 8 Identification of Germ-Line Stem Cells in Zebrafish. . . . . . . . . . . . . . . . . . . . . . . . . Bruce W. Draper 9 Primordial Germ Cell Isolation from Xenopus laevis Embryos . . . . . . . . . . . . . . . . Amanda M. Butler, Tristan Aguero, Karen M. Newman, and Mary Lou King 10 Single-Cell Lineage Analysis of Oogenesis in Mice . . . . . . . . . . . . . . . . . . . . . . . . . . Lei Lei and Allan C. Spradling 11 Visualization and Lineage Tracing of Pax7+ Spermatogonial Stem Cells in the Mouse . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ˜ a, Gina M. Aloisio, Ileana Cuevas, Yuji Nakada, Christopher G. Pen and Diego H. Castrillon 12 Transplantation as a Quantitative Assay to Study Mammalian Male Germline Stem Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Aileen R. Helsel and Jon M. Oatley 13 Mouse Fetal Germ Cell Isolation and Culture Techniques . . . . . . . . . . . . . . . . . . . Cassy M. Spiller, Guillaume Burnet, and Josephine Bowles

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49 63 75

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Contents

Rattus norvegicus Spermatogenesis Colony-Forming Assays. . . . . . . . . . . . . . . . . . 185 F. Kent Hamra Identification of Mouse piRNA Pathway Components Using Anti-MIWI2 Antibodies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 205 Takamasa Hirano, Hidetoshi Hasuwa, and Haruhiko Siomi Efficient Induction and Isolation of Human Primordial Germ Cell-Like Cells from Competent Human Pluripotent Stem Cells. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 217 Naoko Irie and M. Azim Surani

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

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Contributors TRISTAN AGUERO  Department of Cell Biology, University of Miami Miller School of Medicine, Miami, FL, USA GINA M. ALOISIO  Department of Pathology and Cecil H. and Ida Green Center for Reproductive Biology Sciences, University of Texas Southwestern Medical Center, Dallas, TX, USA JOSEPHINE BOWLES  School of Biomedical Sciences, The University of Queensland, Brisbane, QLD, Australia GUILLAUME BURNET  School of Biomedical Sciences, The University of Queensland, Brisbane, QLD, Australia MICHAEL BUSZCZAK  Department of Molecular Biology, University of Texas Southwestern Medical Center, Dallas, TX, USA AMANDA M. BUTLER  Department of Cell Biology, University of Miami Miller School of Medicine, Miami, FL, USA DIEGO H. CASTRILLON  Department of Pathology and Cecil H. and Ida Green Center for Reproductive Biology Sciences, University of Texas Southwestern Medical Center, Dallas, TX, USA AYMERIC CHARTIER  mRNA Regulation and Development, Institut de Ge´ne´tique Humaine, CNRS UPR1142 and University of Montpellier, Montpellier, Cedex 5, France JULIE N.R. COLLINS  Department of Pharmacology, University of Texas Southwestern Medical Centerr, Dallas, TX, USA JAMES J. COLLINS III  Department of Pharmacology, University of Texas Southwestern Medical Center, Dallas, TX, USA SARAH L. CRITTENDEN  Department of Biochemistry, Howard Hughes Medical Institute, University of Wisconsin-Madison, Madison, WI, USA ILEANA CUEVAS  Department of Pathology and Cecil H. and Ida Green Center for Reproductive Biology Sciences, University of Texas Southwestern Medical Center, Dallas, TX, USA BRUCE W. DRAPER  Department of Molecular and Cellular Biology, University of California, Davis, Davis, CA, USA DANA GANCZ  Department of Biological Regulation, Weizmann Institute of Science, Rehovot, Israel LILACH GILBOA  Department of Biological Regulation, Weizmann Institute of Science, Rehovot, Israel LEAH J. GREENSPAN  Department of Cell Biology, Johns Hopkins University School of Medicine, Baltimore, MD, USA F. KENT HAMRA  Department of Pharmacology, Cecil H. & Ida Green Center for Reproductive Biology Sciences, University of Texas Southwestern Medical Center, Dallas, TX, USA HIDETOSHI HASUWA  Department of Molecular Biology, Keio University School of Medicine, Tokyo, Japan AILEEN R. HELSEL  Center for Reproductive Biology, School of Molecular Biosciences, College of Veterinary Medicine, Washington State University, Pullman, WA, USA

ix

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Contributors

TAKAMASA HIRANO  Department of Molecular Biology, Keio University School of Medicine, Tokyo, Japan; Division for Mammalian Development, National Institute of Genetics, Mishima, Shizuoka, Japan MAYU INABA  Department of Cell and Developmental Biology, Life Sciences Institute, Howard Hughes Medical Institute, University of Michigan Medical School, Ann Arbor, MI, USA; Department of Molecular Biology, University of Texas Southwestern Medical Center, Dallas, TX, USA NAOKO IRIE  Wellcome Trust Cancer Research UK Gurdon Institute, University of Cambridge, Cambridge, UK; Department of Physiology, Development and Neuroscience, University of Cambridge, Cambridge, UK WILLY JOLY  mRNA Regulation and Development, Institut de Ge´ne´tique Humaine, CNRS UPR1142 and University of Montpellier, Montpellier, Cedex 5, France JUDITH KIMBLE  Department of Biochemistry, Howard Hughes Medical Institute, University of Wisconsin-Madison, Madison, WI, USA MARY LOU KING  Department of Cell Biology, University of Miami Miller School of Medicine, Miami, FL, USA LEI LEI  Department of Cell and Developmental Biology, University of Michigan Medical School, Ann Arbor, MI, USA ERIKA L. MATUNIS  Department of Cell Biology, Johns Hopkins University School of Medicine, Baltimore, MD, USA YUJI NAKADA  Department of Pathology and Cecil H. and Ida Green Center for Reproductive Biology Sciences, University of Texas Southwestern Medical Center, Dallas, TX, USA KAREN M. NEWMAN  Department of Cell Biology, University of Miami Miller School of Medicine, Miami, FL, USA JON M. OATLEY  Center for Reproductive Biology, School of Molecular Biosciences, College of Veterinary Medicine, Washington State University, Pullman, WA, USA; Center for Reproductive Biology, College of Veterinary Medicine, Washington State University, Pullman, WA, USA CHRISTOPHER G. PEN˜A  Department of Pathology and Cecil H. and Ida Green Center for Reproductive Biology Sciences, University of Texas Southwestern Medical Center, Dallas, TX, USA HANNAH S. SEIDEL  Department of Biochemistry, Howard Hughes Medical Institute, University of Wisconsin-Madison, Madison, WI, USA NEVINE A. SHALABY  Department of Molecular Biology, University of Texas Southwestern Medical Center, Dallas, TX, USA; Institute for Biology, Freie Universit€ at, Berlin, Germany MARTINE SIMONELIG  mRNA Regulation and Development, Institut de Ge´ne´tique Humaine, CNRS UPR1142 and University of Montpellier, Montpellier, Cedex 5, France HARUHIKO SIOMI  Department of Molecular Biology, Keio University School of Medicine, Tokyo, Japan CASSY M. SPILLER  School of Biomedical Sciences, The University of Queensland, Brisbane, QLD, Australia ALLAN C. SPRADLING  Department of Embryology, Howard Hughes Medical Institute, Carnegie Institution for Science, Baltimore, MD, USA

Contributors

xi

M. AZIM SURANI  Wellcome Trust Cancer Research UK Gurdon Institute, University of Cambridge, Cambridge, UK; Department of Physiology, Development and Neuroscience, University of Cambridge, Cambridge, UK YUKIKO M. YAMASHITA  Department of Cell and Developmental Biology, Life Sciences Institute, Howard Hughes Medical Institute, University of Michigan Medical School, Ann Arbor, MI, USA

Chapter 1 Analysis of the C. elegans Germline Stem Cell Pool Sarah L. Crittenden, Hannah S. Seidel, and Judith Kimble Abstract The Caenorhabditis elegans germline is an excellent model for studying the regulation of a pool of stem cells and progression of cells from a stem cell state to a differentiated state. At the tissue level, the germline is organized in an assembly line with the germline stem cell (GSC) pool at one end and differentiated cells at the other. A simple mesenchymal niche caps the GSC region of the germline and maintains GSCs in an undifferentiated state by signaling through the conserved Notch pathway. Downstream of Notch signaling, key regulators include novel LST-1 and SYGL-1 proteins and a network of RNA regulatory proteins. In this chapter we present methods for characterizing the C. elegans GSC pool and early germ cell differentiation. The methods include examination of the germline in living and fixed worms, cell cycle analysis, and analysis of markers. We also discuss assays to separate mutants that affect the stem cell vs. differentiation decision from those that affect germ cell processes more generally. Key words Stem cells, Progenitor cells, C. elegans, Germline, Proliferation, Meiosis, Mitosis, EdU, Cell cycle

1

Introduction Identification of stem cells and the pathways that regulate them are important for both clinical research and more basic biomedical science. The Caenorhabditis elegans germline is a simple and wellstudied model for understanding the genetic and molecular regulation of stem cells [1–13]. Several qualities distinguish the C. elegans gonad as a model for GSC regulation. First, in contrast to other GSC models with asymmetrically dividing stem cells, C. elegans GSCs are maintained as a pool (Fig. 1a) [14, 15]. Second, all stages of germ cell development, from stem cell to differentiated gamete, are present in the adult gonad at one time (Fig. 1a). Third, establishment and maintenance of C. elegans germline stem cells is controlled by a simple, single-celled

Electronic supplementary material: The online version of this chapter (doi: 10.1007/978-1-4939-4017-2_1) contains supplementary material, which is available to authorized users. Michael Buszczak (ed.), Germline Stem Cells, Methods in Molecular Biology, vol. 1463, DOI 10.1007/978-1-4939-4017-2_1, © Springer Science+Business Media New York 2017

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A

Progenitor Zone Distal pool

Early meiotic prophase

Proximal pool

DTC niche Stem cell-like Transition state between states

B

C Proliferation phase

Differentiated state

LST-1 SYGL-1

GLP-1/Notch signaling ?

Maintenance phase

2

FBF-1 FBF-2

Germline stem cell self-renewal

Germline development L1

#GC 2

L2 16 L3 ~60 L4 ~400 Adult (24 hour)

Adult (96 hour)

~2000 cell death embryos

~2000

Fig. 1 Introduction to C. elegans germline development. (a) Diagram of adult hermaphrodite. Gonad arms are color coded. Yellow, progenitor zone. Green, meiosis. Pink, oocytes. Blue, sperm. Red, the somatic distal tip cell (DTC), which is located at the distal end and provides a niche for proliferating germline stem cells. Germ cells differentiate as they move away from the DTC, toward the proximal end of the germline. (b) Pathway controlling the progenitor/ differentiation switch in the C. elegans germline. (c) Diagram showing the stages of germline development from the larval proliferative phases through the adult maintenance phase. Approximate germ cell numbers for each stage are given on the right

C. elegans Germline Stem Cells

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mesenchymal niche, the distal tip cell (DTC) (Fig. 1a) [16]. Finally, regulators of stem cell self-renewal have been identified and analyzed in depth (Fig. 1b). Both the Notch signaling pathway [1] and the PUF family of RNA regulators maintain stem cells in C. elegans [1, 7, 17, 18]. These regulators also control stem cells in other systems. For example, Notch signaling has been suggested to play roles in stem cell regulation in vertebrates ([4] and refs. therein). PUF proteins are required for germline stem cell self-renewal in flies ([1] and refs. therein) and have been found in human spermatogonia and embryonic stem cells [19]. The C. elegans GSCs generate the germline during larval development, expanding it from 2 to 2000 cells (Fig. 1c); they maintain the germline during adulthood, replenishing it as mature gametes are lost to cell death and fertilization (Fig. 1c); and they regenerate the germline after starvation [14, 20–22]. GSCs are pluripotent, giving rise to both sperm and oocytes, which, after fertilization, produce a totipotent embryo. GSC behavior is influenced by sexual identity [23], food abundance [21, 22, 24, 25], food quality [26], age [27, 28], and rate of gamete production [23, 29]. In addition, a number of screens have identified genes that modify the activity of the core pathway controlling GSCs. Modifiers include splicing factors, components of the proteasome, RNA binding proteins, and genes of unknown function (see Refs. in [1, 2, 30–32]). In this chapter we describe methods for studying C. elegans germline stem and progenitor cells. Criteria to identify stem cells can vary; thus it is crucial to define what is known in the system being worked on and to define the criteria for identifying cell types. In the C. elegans germline, immature stem and progenitor cells are at the distal end, adjacent to the distal tip cell (Fig. 1a). The region of the germline containing these cells has been referred to by various names, including mitotic region, mitotic zone, and proliferation zone. It has become clear that the undifferentiated state is maintained independent of the cell cycle state, so a term such as progenitor zone [27, 28] is more appropriate. The progenitor zone contains the germline stem cells (GSCs) as well as their progeny in various early states of differentiation. Lineage tracing is not yet possible in the C. elegans germline, so GSCs cannot be unequivocally identified. However, several experiments indicate that GSCs reside within the distal part of the progenitor zone. There is a pool of 35–70 undifferentiated cells found within the distal 5–8 rows of the germline [15] that have similar cell cycle properties [14, 33–35], can regenerate the entire germline after starvation [21, 22], and express high levels of the mitotic activators GLP-1, lst-1, sygl-1, FBF-1 and FBF-2 and low levels of the meiotic activators GLD-1 and GLD-2 (Fig. 1b and Fig. 8) [1, 9, 15, 31, 36–38]. This group of cells comprises the GSC pool.

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Farther from the DTC, the proximal pool includes germ cells in the early phases of differentiation. Two separate decisions, both of which are required for gametogenesis, occur in this pool: the decision to enter the meiotic cell cycle [15] and the decision to become sperm or oocyte [39]. The proximal pool cells express markers of both meiotic and sexual differentiation, such as GLD1, HIM-3, and FOG-1 (Fig. 8) [15, 38, 40–43]. Proximal pool germ cells are similar to transit-amplifying cells in other systems in that they have been triggered to differentiate but are still dividing mitotically. Their amplification is not robust; they are estimated to divide 1–2 times before entering meiosis [38]. In addition to transit-amplifying cells, the proximal pool includes germ cells that are premeiotic, defined as cells that will enter meiotic prophase without passing through mitosis [14, 34, 41] (Fig. 1a and Fig. 8). Since, at present, there are no good markers for premeiotic S phase, we cannot reliably distinguish premeiotic S-phase cells from mitotically dividing cells. An outstanding question is whether all cells in the progenitor zone have stem cell potential. It is technically difficult to test this directly in the C. elegans germline; however, it is known that distal and proximal pools have shared characteristics. They all express the GLP-1/Notch receptor and depend on its activity to inhibit entry into meiosis. They all cycle and do so at similar rates and they also divide with a random orientation [8, 14, 23, 33, 34, 44]. They all have sexual identity based on sex-specific differences in morphology, gene expression, and cell cycle rate [23, 33, 43, 45–47]. Whether these shared characteristics indicate shared potential is unclear. We will present methods for identifying and characterizing undifferentiated and proliferating germ cells, including identification of the progenitor zone, putative stem cells, and the premeiotic region. We will then discuss how we characterize new mutants. There are excellent chapters about the germline and useful techniques available on the WormBook website (http://www.wormbook. org), the WormBook Methods website (http://www.wormbook. org/toc_wormmethods.html), and in several books [48–51].

2 2.1

Materials Reagents

1. 4 % agarose in dH2O for microscopy of live C. elegans. 2. M9: 3 g/L KH2PO4, 6 g/L Na2HPO4, 5 g/L NaCl, 1 mM MgSO4. 3. E. coli M9 minimal media: 3 g/L KH2PO4, 6 g/L Na2HPO4, 0.5 g/L NaCl, 1 g/L NH4Cl, 2 mM MgSO4, 0.1 mM CaCl2, 0.4 % glucose, 1.25 μg/ml thiamin. 4. M9 plus 0.25 mM levamisole.

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5. Slides and coverslips. 6. Subbing solution: (a) Bring 200 ml distilled water to 60  C. (b) Add 0.4 g gelatin. (c) Cool to 40  C. (d) Add 0.04 g chrome alum. (e) Add 1 mM sodium azide. (f) Add poly-L-lysine (Sigma) to 1 mg/ml. Store subbing solution at 4  C. To sub slides, put subbing solution on slide for 10 min at room temperature. Wick off excess liquid. Dry in 65  C oven for 30 min. Slides can be stored in the oven or at room temperature. 7. Paraformaldehyde (16 % stock, Electron Microscopy Sciences). 8. PBSB: PBS (for 1 L: 8 g NaCl, 0.2 g KCl, 1.44 g Na2HPO4, 0.24 g KH2PO4, pH to 7.2 with NaOH) containing 0.5 % BSA. 9. PBSBTw: PBSB containing 0.1 % Tween 20 to prevent sticking. 10. DAPI (40 ,6-diamidino-2-phenylindole) (Molecular Probes, Invitrogen). 11. Vectashield (Vector Labs). 12. Hoechst 33342 (Molecular Probes, Invitrogen). 13. SYTO-12 (Molecular Probes, Invitrogen). 14. Rabbit anti-PH3 polyclonal antibody (Upstate Biotechnology). 15. Mouse anti-PH3 monoclonal antibody (Cell Signaling Technology). 16. M9-agar plates: 1.2 % agar, 0.6 % agarose in M9 salts containing 0.1 mg/ml carbenicillin [52]. 17. Click-iT® EdU Alexa Fluor® 488 Imaging Kit (ThermoFisher). 18. TO-PRO-3 (Molecular Probes, Invitrogen). 19. Thymidine-deficient E. coli MG1693 (E. coli Genetic Stock http://cgsc.biology.yale.edu/top.html, Center (CGSC), CGSC #6411). 20. Psygl-1::H2B::GFP::sygl-1; Poma-1::OMA-1::GFP (Caenorhabditis Genetics Center (CGC), http://cbs.umn.edu/cgc, C. elegans strain #JK5018). 21. Plag-2::GFP (Caenorhabditis Genetics Center (CGC), http:// cbs.umn.edu/cgc, C. elegans strain #JK2868). 22. Plag-2::myr::GFP (Caenorhabditis Genetics Center (CGC), http://cbs.umn.edu/cgc, C. elegans strain #JK4475).

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23. Plim-7::GFP (Caenorhabditis Genetics Center (CGC), http:// cbs.umn.edu/cgc, C. elegans strain #DG1575). 24. VALAP (equal parts petrolatum, lanolin, and paraffin wax, melted to combine). 2.2

Web Resources

1. Reinke microarray data: http://cmgm.stanford.edu/kimlab/ germline/. 2. In situ RNA expression database: http://nematode.lab.nig.ac. jp/. 3. WormBase: http://www.wormbase.org/. 4. modENCODE: http://www.modencode.org/. 5. C. elegans strain collection (CGC): http://www.cbs.umn.edu/ CGC/. 6. WormBook: http://www.wormbook.org/. 7. WormMethods: http://www.wormbook.org/toc_wormmethods. html. 8. WormAtlas: http://www.wormatlas.org.

3

Methods

3.1 Identification of the Progenitor Zone in Wild-Type C. elegans Hermaphrodites

The C. elegans germline is composed of two U-shaped tubes containing approximately 2000 germ cells in different states of differentiation, which can be observed in living animals (see Fig. 2a). The germline is syncytial, although cells in the progenitor zone are nearly completely enclosed by membrane and are referred to as germ cells. The undifferentiated germ cells, including GSCs, reside at one end, adjacent to the somatic distal tip cell. The organization of progenitor and meiotic cells is similar in hermaphrodites and males. C. elegans hermaphrodites are self-fertile XX animals which first make sperm, then oocytes, whereas males are XO animals that make only sperm. In wild-type young adult hermaphrodites (see Note 1), the progenitor zone is approximately 20 cell diameters in length and contains approximately 225–250 germ cells. The length of the progenitor zone is defined as the number of cell diameters between the distal tip cell and the transition zone (TZ) (Subheading 3.5, Fig. 8, and refs [14, 41]). The TZ contains cells in early meiotic prophase; when stained with DAPI, the DNA in these nuclei has a distinctive crescent shape [53, 54] (crescents, Fig. 3, Subheadings 3.3 and 3.5). Most cells between the DTC and the transition zone are proliferating; however, approximately 50–100 germ cells in the most proximal rows are premeiotic [14, 34]. In wild-type germlines, apoptosis occurs in some oogenic cells in the late pachytene stage of meiotic prophase (Subheading 3.4, ref. [55]). Apoptosis does not occur in the male germline [55].

C. elegans Germline Stem Cells

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undifferentiated germ cells

pachytene germ cells

sp vulva

7

oocytes

uterus/spermatheca

wild-type

B sp

intestine sp vulva

fbf-1 fbf-2 undifferentiated germ cells throughout

C

gld-2 gld-1; fbf-1 fbf-2

intestine

vulva

Fig. 2 Microscopy of wild-type and mutant germlines. (a) Differential interference contrast (DIC) micrograph of a wild-type adult hermaphrodite. (b) DIC micrograph of an fbf-1 fbf-2 double mutant germline. Mature sperm are seen at the distal end. (c) DIC micrograph of a gld-2 gld-1; fbf-1 fbf-2 mutant germline containing only undifferentiated germ cells. Arrowheads indicate distal end of germline 3.2 Mounting Live Animals for Light Microscopy (see Notes 2 and 3)

1. Make an agarose pad on slide: drop melted 4 % agar on glass slide, quickly put another slide on top at a 90 angle and press to create a thin layer of agarose (about 0.4 mm thick) (see ref [56], p. 596). Slides with lab tape can be used as spacers for the top slide [57, 58]. 2. Pick animals onto agarose pad next to a drop of M9 containing 0.25 mM levamisole (see Note 4). Levamisole will prevent animals from moving. Tricaine and tetramisole are also used [59]. Animals can be rescued from agarose pads by carefully sliding coverslip to the side and then picking the animals from the pad to a plate. 3. Cover with coverslip, avoiding bubbles. For short-term observation, it is not necessary to seal the edges of the coverslip. For longer term imaging, coverslips can be sealed with VALAP [50]. 4. Observe germline using differential interference contrast (DIC) and/or fluorescence microscopy.

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Progenitor zone

Transition zone (TZ)

Pachytene region

Wild-type

Progenitor nuclei

TZ nuclei

Pachytene nuclei

Fig. 3 Identification of progenitor zone cells and meiotic germ cells. DAPI-stained distal arm of a wild-type adult hermaphrodite. Progenitor zone, transition zone (TZ), and pachytene region nuclei are enlarged below. Dashed and dotted lines indicate the boundaries between the zones. Note the crescents of DNA in the TZ nuclei and the condensed strands of chromatin in the pachytene nuclei. Arrowhead points to DTC nucleus. Modified from Eckmann et al., 2014 [63] 3.3 Immunohistochemistry of Extruded Gonads

We use two methods for extruding and processing gonads (somatic tissues remain associated with the germline) for immunohistochemistry: extrusion on slides and extrusion in solution ([41, 60]; Schedl lab website—protocols http://genetics.wustl.edu/tslab/). One benefit of extruding in solution is the increased efficiency for preparing large numbers of gonads, although there are anecdotal reports of improved staining using the slide method. There are several detailed protocols for staining C. elegans gonads that include alternate fixation protocols [60, 61] (see Notes 5 and 6). If only DAPI staining is required, skip the primary antibody step and incubate the fixed germlines with DAPI. Continue on from there (see Note 7). 1. Extrude gonads. To extrude on a slide, pick 10–20 animals into 7 μl PBS containing 0.25 mM levamisole on subbed slide. Cut the animals with a syringe needle or scalpel behind the pharynx or at the tail. The germline and intestine will extrude from the cuticle (see Note 8). Make a sample area with a PAP pen to keep the liquid from spreading across the slide (see Note 9). To extrude in solution, pick or wash 10–100s of animals off plate into a glass or plastic dish containing PBS, 0.25 mM levamisole plus 0.1 % Tween 20 to prevent sticking. The total volume can be anywhere from 200 μl to a couple milliliters depending on your personal preference and size of the dish. Use a micropipettor to transfer samples into a microfuge tube. For the following steps, the volumes should be matched to the technique

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you use to extrude gonads. For extrusion on slides, generally 25–50 μl is sufficient to cover the sample area. For the solution method, volumes for fixation, permeabilization, and washes are generally 200–1000 μl, and volumes for antibody incubation are 50–100 μl; we usually place tubes on a rotator or rocker during all steps. 2. Fix. The fixation method will depend on the antibody antigen combination you are using (see Note 5) [60]. A good place to start is 1–3 % paraformaldehyde for 10–20 min. If you extruded on a slide, keep the slides in a humidified chamber (e.g., a box with a wet paper towel taped to the lid) throughout the procedure. 3. Permeabilize. Remove paraformaldehyde and replace with 0.1 % Triton X100 in PBSB for 5 min at room temperature. 4. Block. In PBSBTw for 30 min at room temperature. 5. Incubate with primary antibody. Dilution, incubation time, and temperature should be determined for each antibody. 6. Wash three times with PBSBTw for 15 min each. 7. Incubate with secondary antibody. Dilution, incubation time, and temperature should be determined for each antibody in PBSBTw (see Note 10). Include 0.1 μg/ml DAPI to stain DNA (see Note 11). 8. Wash three times with PBSBTw for 15 min each. 9. Add 7–10 μl Vectashield or other mounting medium; if samples are in a tube, pipet onto a slide, place coverslip over gonads, and seal with nail polish. The volume of mounting medium will depend on the size of the coverslip. The volume should be large enough that gonads are not squashed, but small enough that they are not floating around. Excess liquid can be carefully wicked away using a kimwipe. 10. Observe using fluorescence microscopy. 3.4 Visualization of DNA and Germline Apoptosis in Live Animals [55, 62]

1. Incubate animals in 33 μM SYTO-12 (taken up by dying cells) and/or 50 μg/ml Hoechst 33342 (DNA dye) in M9 with some E. coli in a microfuge tube for 4–5 h (see Notes 12 and 13). 2. Transfer to seeded plate for 30–60 min. 3. Mount on agarose pad for DIC microscopy. 4. Observe live animals on agarose pads (Subheading 3.2) using fluorescence microscopy and DIC. Hoechst can be observed using DAPI filter sets. SYTO-12 emits green fluorescence and can be observed using FITC filter sets.

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3.5 Scoring the Length and Cell Number in the Progenitor Zone

The progenitor zone extends from the distal-most germ cell to the start of the transition zone. 1. Using DAPI-stained germlines (Subheading 3.3), identify the distal end by locating the DTC nucleus (oval with diffuse DAPI staining, Figs. 3 and 4) (see Note 14). 2. The start of the transition zone has been defined as either the position of the first crescent [41] or the position at which there are multiple crescents (to reduce the chance of scoring occasional anaphase chromosomes that are curved similar to crescent-shaped meiotic nuclei [8, 14, 63]). These two positions are usually within 1–3 rows of each other (see Note 15) (Fig. 3). 3. Count germ cell rows and total number of cells (see Note 16) starting with the row immediately adjacent to the DTC nucleus and continuing to the transition zone (Fig. 4). Counting can be done visually either through microscope eyepieces or on a computer monitor. For training yourself to count with reasonable accuracy, count the germline multiple times until your numbers are within about 5 % of each other. A software tool for counting germ cells (IRISES) is available and facilitates the counting process with minimal manual input [64]. 4. Progenitor zones in young adult wild-type hermaphrodite germlines average 19–20 cell diameters from the distal end (range 15–24) and contain 225–250 germ cells (range 200 to  300) (see Note 1). These numbers change with age (e.g., [27, 28]), so comparing staged animals is important. In addition, the number of germ cells in the progenitor zone does not always correlate with the length (e.g., male progenitor zones are longer than hermaphrodite, but contain the same number of cells [23]).

Hoechst DTC 1

2 3456

7 8 9 10 12

14

16

18 20

22

24

Fig. 4 Counting cell diameters from distal end of germline. Distal arm of a wildtype adult hermaphrodite gonad stained with Hoechst. DTC nucleus is circled in red, germ cell nuclei are circled in white. Numbers begin with 1 for the nucleus closest to the DTC and increase moving proximally. Dashed line indicates TZ boundary. Modified from Byrd et al., 2014 [62]

C. elegans Germline Stem Cells

3.6 Characterization of the GSC Pool and GSC Number

11

1. sygl-1 and lst-1 expression. Two GLP-1/Notch targets, sygl-1 and lst-1, are expressed in a subset of germ cells immediately adjacent to the DTC in the progenitor zone. The expression of these genes is dependent on GLP-1/Notch signaling, making their expression a useful marker for GLP-1/Notch responsive germ cells [65] within the distal pool. Expression patterns for endogenous sygl-1 and lst-1 can be examined by in situ hybridization; detailed methods can be found in Kershner et al., 2014 [65] (see Note 17). sygl-1 and lst-1 are also expressed in maturing oocytes, so mutants that cause distal germ cells to differentiate may express sygl-1 and lst-1 distally due to differentiation rather than maintenance of an undifferentiated state. sygl-1 expressing cells can also be visualized using animals carrying a single copy chromosomal insertion of a transgene containing the sygl-1 promoter driving GFP::H2B [65]. Note that some germlines expressing this transgene are not completely normal and that expression is partially silenced. Germline silencing is common in C. elegans and can be alleviated by maintaining transgenic animals in a licensing background [66–68] and/or by growing them at 25  C. See Kershner et al., 2014 (Ref. [65] Materials and methods) for further details. 2. emb-30 assay. In this assay, germ cell movement is blocked and germ cells are scored for differentiation with time [15]. The emb-30(ts) mutant at restrictive temperature blocks the metaphase to anaphase transition, arresting cell division and cell movement [15]. After shift to restrictive temperature, the distal-most 6–8 cell rows, containing about 50 germ cells, arrest in M phase and remain undifferentiated, indicating that this pool of cells is maintained in an immature state. More proximal germ cells enter the meiotic cell cycle, indicating that cells in that position in wild-type gonads have been triggered to differentiate and would normally continue into meiosis as they move proximally (Fig. 5, compare panels (a) and (b), Note 18). (a) Maintain emb-30(tn377ts) at 15  C. Animals are fertile; however, progenitor zones are slightly smaller than in wild type. Maintain strains on plates that are equilibrated to 15  C. If you are determining distal pool size in mutants, compare mutant; emb-30 to þ; emb-30 in parallel. (b) Stage animals. For animals comparable to hermaphrodites grown for 24 h after L4 at 20  C (see Note 1), pick L4s and let them grow at 15  C for 36 h. (c) Shift staged animals to restrictive temperature (25  C) by picking adults to plates that have been equilibrated at 25  C.

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A

0 hours

B

15 hours

anti-PH3

anti-GLD-1

DAPI

Fig. 5 emb-30 assay. Gonads were shifted from permissive (15  C) to restrictive (25  C) temperatures for times shown in white lettering (Panel (a), 0 h at restrictive temperature; Panel (b), 15 h at restrictive temperature). Gonads were extruded and stained with anti-PH3 (green), anti-GLD-1 (red), and DAPI (cyan). Distal end is marked with white triangle, GLD-1 boundaries with red carats, most proximal PH3 with green arrow. GLD-1 often has two stepwise increases in intensity—each is marked with a caret in the 0 h panels (a). These steps can be absent or look graded in some gonads, making it hard to score. Also note the variable nuclear morphology in distal end of the 15 h DAPI-stained gonad (b). Dashed line indicates TZ boundary. Modified from Cinquin et al., 2010 [15]

(d) Every hour, or at your chosen time points, fix gonads (Subheading 3.3) and stain with DAPI, anti-PH3 and anti-GLD-1 or anti-HIM-3 (Fig. 5a, b). Score the position of the boundary between metaphase-arrested nuclei (visualized with DAPI and anti-PH3) and transition zone nuclei (visualized with DAPI). Also score the position of the boundary between low and high levels of meiotic proteins such as GLD-1 and/or HIM-3. Positions can be scored in germ cell diameters and/or microns (see [15, 28]; Note 18). 3.7 Markers for Progenitor Zone and Early Stages of Differentiation

A number of markers are helpful for marking the progenitor zone, the distal pool of progenitor cells, differentiating progenitors, germ cells in the meiotic cell cycle, and somatic gonad cells that affect proliferation (Table 1, Fig. 8). Progenitor zone markers include the Notch receptor, GLP-1, which is abundant in membranes of progenitor zone cells [69]; endogenous sygl-1 and lst-1 transcripts ([65]; see Subheading 3.6) and a transgene containing the sygl-1 promoter driving H2B::GFP ([65]; see Subheading 3.6), which are abundant in the distal-most germ cells; the PUF family RNA regulators, FBF-1 and FBF-2, which are abundant in the cytoplasm of progenitor zone cells [70–72]; the p53 homolog, CEP-1, which is present in progenitor zone nuclei [73]; and the meiotic cohesin, REC-8, which is enriched in the nuclei of progenitor zone cells under certain fixation conditions [8, 41]. All of these markers are also expressed either at lower levels in early meiotic prophase or in other parts of the germline (Table 1, Fig. 8). Meiotic markers include the RNA regulator, GLD-1, which increases in the cytoplasm of germ cells in

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Table 1 Useful markers Marker

Molecular identity

Pattern

lst-1

Novel GLP-1/Notch target

Transcript is present in a distal subset of germ cells [65]

sygl-1

Novel GLP-1/Notch target

Transcript is present in a distal subset of germ cells [65]

Psygl-1::H2B:: GFP::sygl-1

sygl-1 promoter driving histone fused to GFP

GFP is present in a distal subset of germ cell nuclei [65].

GLP-1

Notch receptor

Membrane-associated, strong in mitotic germ cells, lower levels in early meiotic region [69]

FBF-1 and FBF-2

PUF family RNA regulators

Cytoplasmic, strong in progenitor zone. Levels decrease as germ cells progress through meiosis. See Lamont et al., 2004 for details of expression patterns [70, 71]

REC-8

Cohesin

On chromosomes in meiotic prophase (except when used as mitotic marker) (see ref [41, 142])

CEP-1

C. elegans p53

Nuclear in progenitor zone and proximal cells [73]

PH3

Histone H3 phosphorylated on serine 10

Chromosomes in late prophase (mitotic and meiotic) and M phase [14, 41, 80]

GLD-1

maxi-KH/STAR domain RNA binding protein

Cytoplasmic, increasing in proximal mitotic cells, strong in meiotic germ cells [40, 47]

HIM-3

Component of central element of synaptonemal complex

Cytoplasmic in proximal progenitor zone, on chromosomes in meiotic prophase [42]

PGL-1

P-granule component

Only in germ cells, not in somatic cells [137]

SP56

Sperm-specific protein

Spermatocytes and mature sperm [138]

RME-2

C. elegans yolk receptor

Oocyte membranes [141]

Plag-2::GFP

lag-2 promoter driving GFP

Somatic DTC [14, 74]

Plim-7::GFP

lim-7 promoter driving GFP

Somatic sheath cells 1–4 [75, 121]

the proximal pool and peaks in the meiotic region [40, 47], and the meiotic protein HIM-3, which is present in proximal progenitor zone cells and becomes localized to chromosomes in early meiotic prophase [41, 42]. Hansen and colleagues [41] used a combination of REC-8 and HIM-3 staining to identify a region of overlap called the meiotic entry region. The meiotic entry region has substantial overlap with the premeiotic S region identified by cell cycle analysis [14, 34, 41] (Fig. 8).

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A

cap

L4

B

long external processes cap

plexus

Adult

Fig. 6 DTC morphology. Gonad from hermaphrodite expressing myristoylated GFP driven by the lag-2 promoter. Hermaphrodites have a single DTC/gonad arm. The DTC has a membranous cap that covers the distal-most germ cells during larval development (a) and adulthood (b). During adulthood, the DTC extends a complex network of processes (plexus) as well as long external processes. Male gonads have two DTCs per gonad arm. Their morphology is less complex, but similar to the hermaphrodite. Modified from Byrd et al., 2014 [62] 3.8 Markers for the Somatic Gonad

The somatic niche, the DTC, can be visualized using a transgene with the lag-2 promoter driving GFP [14, 74]. Finer details of membrane structure can be seen using a transgene with myristoylated GFP [62]. In hermaphrodites, the DTC has a membranous cap that surrounds the distal-most germ cells. The DTC produces extensive processes in early adulthood; these processes surround and intercalate into the membranes between the germ cells in the distal pool (plexus, Fig. 6). During adulthood, the DTC also extends long, external processes [14, 62, 75, 76]. There is a similar DTC architecture in males; however, the lag-2 promoter is much weaker in the male DTCs, making it harder to visualize processes [23]. Other ligands such as APX-1 are also expressed in the niche and play a role in stem cell maintenance [77] (see Note 19). The somatic sheath cells also play a role in germline patterning, especially during larval stages [78 and refs. therein]. Sheath cells can be visualized using a transgene with the lim-7 promoter driving GFP. The sheath cells also have extensive contact with the surface of the germ cells and extensive intercalation between germ cells [75, 78].

3.9 Cell Cycle Analysis

Cell cycle analysis allows estimation of cell cycle lengths, identification of quiescent or slow-cycling cells, and identification of cells entering meiotic S phase [79]. Anti-phosphohistone H3 (PH3) staining, which labels condensed chromosomes in late G2 and Mphase nuclei [80], is used to determine mitotic index (% of cells in M phase) (Subheading 3.10) [14, 34, 41]. Alternatively, mitotic figures can be scored by DAPI staining [44]. EdU labeling is used to determine the S-phase labeling index (% of cells in S phase) and to estimate total cell cycle length (Subheadings 3.11–3.18). Under fixation conditions that preserve morphology well, EdU labeling combined with nuclear size can be used to identify G1 vs. G2 cells

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(Subheading 3.14). The proximal region of the progenitor zone, where mitotic index is low and labeling index is the same as the rest of the progenitor zone, is likely composed largely of premeiotic germ cells (Subheading 3.18). Some premeiotic S-phase cells are also found in the transition zone. 3.10 Mitotic Index with Anti-PH3

1. Prepare animals for antibody staining (see Subheading 3.3). 2. Stain with anti-PH3. In our hands, mouse anti-PH3 (Cell Signaling) reliably stains prophase and metaphase nuclei. Anaphase and telophase nuclei stain weakly and more variably. 3. Score position and number of PH3-positive nuclei in progenitor zone (Fig. 5, see Note 16). 4. Count number of nuclei/row for normalizing. 5. Calculate mitotic index by dividing number of PH3-positive cells by the total number of cells in the region of interest.

3.11

EdU Labeling

3.12 Label with EdU by Feeding

EdU labeling can be performed by feeding or by soaking [23, 28, 34, 38, 52]. Both methods are appropriate for pulse labeling (15–30 min), but only feeding is appropriate for pulse-chase labeling or for labeling over longer time periods, because the stress of soaking can affect cell cycle dynamics (see Note 21). Germ cells can also be labeled with BrdU [14, 81] and Cy3dUTP [33, 82, 83]; however, BrdU is more difficult than EdU to visualize well and Cy3dUTP has to be injected. 1. Grow an overnight culture of E. coli MG1693 in LB or in M9 minimal media supplemented with 5 μg/ml thymine. 2. To 100 ml of E. coli M9 minimal media, add: 3 ml of MG1693 overnight culture. 100 μl 0.5 mM thymidine (0.5 μM). 200 μl 10 mM EdU (Invitrogen) (20 μM) (see Note 20). Grow at least 24 h at 37  C. Spin to pellet E. coli. Resuspend in about 1 ml M9. 3. Seed plates with 200–400 μl labeled MG1693. Try to cover most of the plate so that animals are continuously feeding on labeled bacteria. To prevent growth of bacteria after seeding (which should be avoided because plates do not contain EdU), use nematode growth media lacking peptone and containing 60 μg/ml carbenicillin. Alternatively, plates containing E. coli M9 agar þ 60 μg/ml carbenicillin can be used, although we have found that the ammonia in E. coli M9 minimal media can affect the cell cycle of C. elegans germ cells. Plates can be stored at 4  C for a couple of weeks. 4. Put animals on plates for desired time period.

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5. To chase, wash animals off plate with M9, wash once with M9, and place animals on plate containing unlabeled bacteria. 6. Fix with 3 % PFA 30 min at room temperature. Weaker fixation conditions can also be used (e.g., 1 % PFA for 10 min), but they do not preserve chromosome morphology well and therefore make it more difficult to distinguish mitotic figures and the nuclear size difference between G1 and G2 cells (see Subheading 3.14). 7. Permeabilize with 0.1 % Triton X-100 5 min at room temperature or with 20  C methanol for 5–15 min. 8. Do the Click reaction according to the instructions. We do 2  30 min half reactions (suggested by [84]). 3.13 Label with EdU by Soaking

1. Transfer animals by picking or washing into M9 þ 0.1 % Tween-20 þ 0.05–1 mM EdU. Incubate with rotation for 15 min. 2. Dissect gonads immediately (with no recovery period) and fix according to the protocol for labeling with EdU by feeding, above.

3.14 Calculating Labeling Index and Identifying G1 and G2 Cells

Cell cycle stage in mitotically dividing germ cells correlates with nuclear size: G1 and early S-phase nuclei are small; G2 and late Sphase nuclei are large; and mid S-phase nuclei are intermediate in size (Fig. 7, Movie 1). The size difference between G1 and G2 cells is large enough that after pulse labeling with EdU to mark S-phase cells, G1 and G2 cells can be distinguished by eye. This correspondence between cell cycle stage and nuclear size has been vetted in wild-type germlines of both sexes, but it may not hold true under conditions that alter nuclear size, such as after knockdown of certain cell cycle regulators [34, 85]. Keep in mind that the fraction of cells detected as EdU-positive will be affected by the sensitivity of the detection system. Early S-phase nuclei label weakly with EdU and may be detected as G1 cells by image acquisition systems that are less sensitive. 1. Pulse animals with EdU for a short time, e.g., 15–30 min (see Subheadings 3.11–3.13). 2. Fix and stain as described in Subheadings 3.11–3.13. Co-stain with DAPI. 3. Examine germline at high magnification (e.g., 63), and identify G1, S-phase, and G2 cells using the following criteria. We find it convenient to acquire z-stack images at 0.5–1 μm intervals and to identify cell cycle stages on a computer monitor. Cells can be easily marked and categorized using the Cell Counter plug-in for ImageJ (http://fiji.sc/Cell_Counter). (a) G1 cells are EdU-negative and have a nuclear size equal to or smaller than the smallest EdU-positive cells (Fig. 7c, Movie 1). Additional characteristics of G1 cells are that

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A DAPI

Progenitor zone

Transition zone

EdU (S-phase)

10 µm

B

C Cell-cycle phase G1

DAPI EdU (S-phase) G1, EdU- small nuclei 10 µm G2, EdU- large nuclei Paired occurence of G1 cells

Nuclear EdU size labeling? Small

Example

—

Early S

Weak

Mid S

Strong

Late S

Strong & punctate

G2

Large

—

Prophase

n/a

—

Metaphase

n/a

—

Anaphase

n/a

—

Telophase

n/a

—

Fig. 7 EdU labeling. (a) Image of an adult hermaphrodite germline stained with DAPI to visualize DNA (top) and labeled with a 15-min pulse of EdU to mark S-phase cells (bottom). Images are maximum-intensity zprojections. (b) Two-color images of adult hermaphrodite germlines treated as in (a). Magenta, DAPI staining. Green, EdU. Yellow circles, G1 cells. Blue circles, G2 cells. Caret, occurrence of G1 cells in pairs. (c) EdU labeling and nuclear size characteristics for cells in different stages of the cell cycle. Scale bar, 2 μm. n/a, nuclear size not applicable during M phase

they are infrequent (5–10 %) and nearly always occur in pairs (Fig. 7b, Movie 1), a pattern consistent with G1 being very short [28, 34]. Chromatin in G1 cells tends to be rather condensed (Fig. 7c), such that the 12 chromosomes are often distinguishable as 12 DAPI-stained puncta (not shown). (b) S-phase nuclei are EdU-positive. S-phase nuclei can be further categorized as early, mid, or late according to nuclear size and pattern of EdU labeling (Fig. 7c, Movie 1). Early S-phase nuclei are small and label weakly

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with EdU; mid S-phase nuclei are mid-sized and label strongly and uniformly with EdU; late S-phase nuclei are large and incorporate EdU in a punctate pattern, likely reflecting the late-replicating X-chromosome [33]. (c) G2 nuclei are EdU-negative and have a nuclear size equal to or larger than the largest EdU-positive nuclei (Fig. 7c, Movie 1). 4. To calculate the labeling index, count number of EdU-positive germ cells and total number of germ cells in region of interest (e.g., entire progenitor zone, one row, or a subregion) then divide number of EdU-positive germ cells by total germ cell number. Analogous calculations are performed to determine the G1 or G2 index. 3.15 Estimating Total Length of the Cell Cycle

3.16 Measure Cell Cycle Length by Measuring G2

There have been several estimates for the total length of the cell cycle in mitotically dividing germ cells [14, 33, 38]. There is variability in cell cycle length among germ cells within a single gonad as well as at different stages of development and under different environmental conditions [28]. Thus, it is important to compare experimental conditions to wild type in your own hands. Cell cycle length can be calculated by measuring the length of G2 or by measuring the length of G2 þ M þ G1. The first method involves labeling animals with EdU and determining the length of time until all M-phase cells become labeled with EdU. The second method determines the time until all cells become labeled with EdU. Measuring the length of G2 is more straightforward because there is a one-to-one correspondence between cells labeled at time zero and cells queried for their labeling status at later time points. 1. Label animals with EdU (on plates) by feeding for 15–30 min to 12 h. 2. Fix and stain some animals every 30 min to 2 h. 3. Determine the fraction of M-phase cells that are EdU-positive at each time point. 4. The time it takes for 50 % of M-phase cells to become EdUpositive is the median length of G2 (the time it takes for the cells in late S phase to traverse G2 and enter M phase). The time it takes for 100 % of M-phase cells to become EdU-positive is the maximum length of G2. 5. Median total length of cell cycle is estimated by dividing the median length of G2 by the fraction of cells in G2. The fraction of cells in G2 represents the proportion of cell cycle time spent in this phase. 6. Estimates of the maximum total length of the cell cycle depend on assumptions about covariance between the length of G2 and the length of M þ G1 þ S. If cells having a longer G2 are

C. elegans Germline Stem Cells

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assumed to have a proportionally longer M þ G1 þ S, then the maximum length of the cell cycle is estimated by dividing the maximum length of G2 by the fraction of cells in G2. If cells having a longer G2 are not assumed to have a proportionally longer M þ G1 þ S, then the maximum length of the cell cycle is given by the following equation: (median total length of cell cycle * (M þ G1 þ S index)) þ time at which all M-phase cells are EdU-positive. These two calculations can yield different estimates if the distribution of G2 lengths is long-tailed. 3.17 Measure Cell Cycle Length by Measuring G2 þ M þ G1

1. Label animals with EdU by feeding for 15–30 min to 12 h. 2. Fix and stain some animals every 30 min to 2 h. 3. Determine labeling index (see Subheading 3.14) for each time point. 4. Median length of G2 þ M þ G1 is given by twice the time it takes for the labeling index to reach mid-way between its starting value and 100 % (the time it takes for cells in early G2 to traverse through G2, M, and G1 and finally label in S phase [86]). The time it takes for 100 % of cells to become EdU-positive is the maximum length of G2 þ M þ G1. 5. Median total length of cell cycle is estimated by dividing the median length of G2 þ M þ G1 by the fraction of cells not in S-phase (i.e., 1–the labeling index) and represents the proportion of cell cycle time spent in these phases. 6. As described for measuring cell cycle length via G2, estimates of the maximum total length of the cell cycle depend on assumptions about covariance between the length of G2 þ M þ G1 and the length of S. If cells having a longer G2 þ M þ G1 are assumed to have a proportionally longer S, then the maximum length of the cell cycle is estimated by dividing the maximum length of G2 þ M þ G1 by the fraction of cells in G2 þ M þ G1. If cells having a longer G2 are not assumed to have a proportionally longer S, then the maximum length of the cell cycle is given by the following equation: (median total length of cell cycle * (S index)) þ time at which all cells are EdU-positive. These two calculations can yield very different estimates if the distribution of G2 þ M þ G1 lengths is long-tailed.

3.18 Identification of Premeiotic Cells in the Progenitor Zone

1. Calculate mitotic index at each position along the distalproximal axis (see Subheading 3.10). 2. Calculate labeling index (15 min pulse) at each position along the distal-proximal axis (see Subheading 3.14). 3. In wild type, the proximal 4–6 rows of the progenitor zone have a lower mitotic index, but the labeling index remains constant. Since these cells do not progress into mitosis (no

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A Progenitor zone

Transition zone

Pachytene

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34

Proposed sites: Distal pool

Proximal pool

DTC plexus

B Cell cycle characteristics S-phase index

~50-70%

~30%

~1%

M-phase index 2.5% G1+M+G2

3.5%

2.5% 10,000  g. Wash pellet once with 70 % EtOH and resuspend in formamide. Use spectrophotometer to determine RNA concentration and adjust to 50 mg/ml with formamide. Freeze 0.5 ml aliquots at 80  C. 5. Prehybridization solution: 50 % Deionized formamide, 5 SSC, 1 mg/ml yeast RNA, 1 % Tween-20. Store at 20  C. 6. Hybridization solution: 50 % Deionized formamide, 5 % dextran sulfate (sodium salt, average molecular weight >500 kD; Sigma), 5 SSC, 1 mg/ml yeast RNA, 1 % Tween-20. Store at 20  C. 7. Wash hybridization solution: 25 % Formamide (non-deionized works), 3.5 SSC, 0.5 % Tween-20, 0.05 % Triton-X100. Store at 20  C. 8. TNT: 0.1 M Tris pH 7.5, 150 mM NaCl, 0.1 % Tween-20. 9. Blocking solution: 7.5 % Heat-inactivated horse serum in TNT. Store at 20  C. 10. Antibody solution: Blocking solution with anti-digoxigeninAP, Fab fragments (Roche) diluted 1:2000 (see Note 5). 11. Alkaline phosphatase buffer: 100 mM Tris, pH 9.5, 100 mM NaCl, 50 mM MgCl2, 0.1 % Tween-20 brought up to volume with 10 % filter-sterilized polyvinyl alcohol (average molecular weight 30–70 kD; Sigma) solution. 12. In situ hybridization development solution: Alkaline phosphatase buffer plus 4.5 μl/ml NBT (4-nitro blue tetrazolium chloride) and 3.5 μl/ml BCIP (4-toluidine salt). 13. Glycerol. 14. Microscope slides and cover slips. 15. Stereomicroscope.

Examining Schistosome Germline Development

2.5 Components for dsRNA Synthesis

41

1. 10 High-yield transcription buffer: 0.4 M Tris pH 8.0, 0.1 M MgCl2, 20 mM spermidine, 0.1 M DTT. 2. 1 μg of PCR product with inverted T7 promoter sites (see Note 3). 3. 100 mM rNTPs (i.e., 25 mM rCTP, rGTP, rUTP, and rATP). 4. Inorganic pyrophosphatase (New England Biolabs). 5. T7 RNA polymerase (see Note 6).

3

Methods Important: Investigators considering schistosome work should contact the Schistosome Resource Center (http://www.niaid.nih.gov/labsandresources/res ources/dmid/schisto/pages/default.aspx). This center provides training for safely working with schistosomes and provides stages of the parasite’s life cycle to investigators. These services are provided free of charge and are supported by the National Institutes of Allergy and Infectious Disease (NIAID). For more information see Lewis et al. [22].

3.1 Fixation and Permeabilization of Parasites for In Situ Hybridization and EdU Detection

1. Euthanize schistosome-infected mice with pentobarbital plus heparin. Perfuse parasites from mice using 37  C DMEM + 5 % FBS. Collect parasites in 50 ml conical tubes and use DMEM + 5 % FBS to rinse bloody perfusate. Transfer ten to several hundred worms to a 15 ml conical tube in 9 ml media. 2. Separate male and female parasites by adding 1 ml anesthetic solution to 9 ml of parasites in DMEM + 5%FBS. Rock samples by hand gently for 1–2 min or until parasites are relaxed and separated. 3. Remove anesthetic solution and kill the parasites in 1 ml of 0.6 M MgCl2 for 1 min. 4. Replace MgCl2 with 4 % formaldehyde in PBSTx, and incubate for 4 h at room temperature (RT) on a rocker. 5. Rinse once with PBSTx. 6. Dehydrate in methanol and store at 20  C. Samples can be stored for weeks, if not months or years, at 20  C. 7. Rehydrate samples in 50 % methanol solution in PBSTx, for 5–10 min at RT. 8. Incubate in PBSTx, for 5–10 min at RT. 9. Add bleaching solution, incubate for 1 h at RT under bright light. 10. Rinse twice in PBSTx and then incubate in proteinase K solution for 45 min at RT (see Note 1). 11. Post-fix in 10 ml 4 % formaldehyde in PBSTx, for 10 min at RT. 12. Rinse in PBSTx.

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Julie N.R. Collins and James J. Collins III

3.2 In Vivo EdU Labeling in S. mansoni

1. Inject schistosome-infected mice intraperitoneally with 100–200 mg/kg EdU in PBS. To monitor cell proliferation in adult parasites, inject mice more than 6 weeks following infection. 2. To monitor the differentiation of germ cells, sacrifice mice and recover parasites at various times following the EdU pulse. At day 1 EdU is largely restricted to the germline stem cells; however at days 2–7 EdU is chased into various differentiated progeny in the testes and ovaries. 3. Fix and permeabilize parasites as described in Subheading 3.1. 4. Remove PBSTx and incubate 10–15 parasites in 25 μl of click chemistry development solution for 30 min at RT in the dark (see Note 7). 5. Wash parasites with constant agitation for 20 min in 1 ml PBSTx. Repeat wash step five times. 6. Stain parasites with 1 μg/ml DAPI in PBSTx overnight at 4  C. 7. Clear parasites in 80 % glycerol for 4 h at RT. 8. Mount parasites on slides, seal cover slips with nail polish, and visualize by fluorescence microscopy.

3.3 Riboprobe Synthesis

1. In an RNase-free 1.7 ml tube combine: l

2 μl 10 in vitro transcription buffer.

l

1 μl 10/6 rNTP solution.

l

0.4 μl 10 mM digoxigenin-11-UTP.

l

0.6 μl RNase inhibitor.

l

2.0 μl T3, T7, SP6 RNA polymerase.

l

1 μl of unpurified PCR product (100–500 ng) (see Note 3).

l

Bring to 20 μl with DEPC water.

2. Incubate at 37  C for 4 h or 27  C overnight. 3. Add 1 μl DNase, and incubate for 20 min at 37  C. 4. Add 5 μl 4 M LiCl and 50 μl EtOH prechilled to Incubate at 20  C for 10 min.

20  C.

5. Centrifuge at >10,000  g for 10 min at 4  C. 6. Decant EtOH and resuspend pellet in 20 μl DEPC water. 7. Quantify RNA concentration on a spectrophotometer using 1 μl riboprobe. 8. Run 2 μl on a 1 % agarose gel. 9. Add 17 μl of hybridization solution and store at

20  C.

Examining Schistosome Germline Development

3.4 Whole-Mount In Situ Hybridization

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1. Transfer parasites fixed and permeabilized as in Subheading 3.1 to a 1.7 ml tube or a 24-well tissue culture plate (see Note 8). 2. Wash in 1:1 (PBSTx:prehybridization solution) for 10 min at RT. 3. Replace with prehybridization solution and incubate for at least 2 h at 52  C. 4. Preheat hybridization solution + riboprobe (100–500 ng/ ml) at 72  C for 5 min and hold at 52  C. 5. Replace prehybridization solution with hybridization solution + riboprobe (100–500 ng/ml). Hybridize overnight (>16 h) at 52  C. 6. Preheat wash hybridization solution, 2 SSC + 0.1 % Triton-X, and 0.2 SSC + 0.1 % Triton-X at 52  C. 7. Remove hybridization solution and wash twice at 52  C for 30 min in wash hybridization solution. 8. Wash twice at 52  C for 30 min with 2 SSC + 0.1 % Triton-X. 9. Wash twice at 52  C for 30 min with 0.2 SSC + 0.1 % Triton-X. 10. Wash twice with TNT for 10 min at RT. 11. Incubate in blocking solution for 2 h at RT. 12. Incubate in antibody solution overnight at 4  C. 13. Wash six times for 20 min each with TNT. 14. Develop in in situ hybridization development solution until desired signal to noise is reached, usually 1–4 h at RT. 15. Development can be stopped by removing development solution and adding PBSTx. 16. Rinse with PBSTx several times. 17. Incubate in 100 % EtOH for 10–20 min at RT. 18. Remove EtOH and add a few drops of PBSTx. Allow parasite to rehydrate for about 5 min. 19. Remove PBSTx and clear parasites in 80 % glycerol in 1 PBS. Incubate at 4  C overnight. 20. Mount on slides and image by bright-field microscopy.

3.5 Generation of dsRNA for RNAi

1. In an RNase free 1.7 ml tube combine: l

10 μl 10 high-yield transcription buffer.

l

20 μl 100 mM rNTPs

l

5.0 μl T7 RNA polymerase.

l

1.0 μl Inorganic pyrophosphatase.

l

l

5 μl of unpurified PCR product with inverted T7 promoters (1 μg) (see Note 3). 59 μl DEPC water.

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2. Incubate at 37  C for 4 h to overnight (see Note 9). 3. Treat with 5 μl of DNase for 20 min at 37  C. 4. Precipitate at RT for 10 min by adding 50 μl of 7.5 M ammonium acetate and 750 μl of EtOH. 5. Centrifuge at >10,000  g for 10 min at 4  C. 6. Decant EtOH and resuspend the pellet in 300 μl DEPC water. 7. Anneal dsRNA by successive 3-min incubations at 95, 75, and 55  C. Allow to cool at RT for 5 min. 8. Dilute 1:10 and determine concentration on a spectrophotometer. Expect 0.5 mg of dsRNA per 100 μl prep. 9. Run remainder of sample dilution on 1 % agarose gel (see Note 10). 10. Dilute the dsRNA to 1 μg/μl using DEPC water. Store at 20  C. 3.6 RNAi in Adult S. mansoni

1. Euthanize mice with pentobarbital plus heparin. Perfuse parasites from mice using 37  C DMEM + 5 % FBS. Collect parasites in 15 cm sterile Petri dishes. 2. In biosafety cabinet, pool parasites in 50 ml conical tubes and allow them to settle. Replace bloody media with fresh DMEM + 5 % FBS until blood is removed. 3. Replace DMEM + 5 % FBS with warmed tissue culture media suitable for maintaining schistosomes (see Note 11). 4. Transfer 5–10 healthy male/female worm pairs in 5 ml of media in a 6 cm Petri dish. This is day 0. 5. Add dsRNA to a final concentration of 20–30 μg/ml. 6. Replace media and dsRNA on days 1 and 2. 7. Change media every other day and provide new dsRNA every 5–8 days until desired experimental endpoint (see Note 12).

4

Notes 1. Proteinase K activity can decrease over time and vary depending on the vendor or manufacture lot. In our experience proteinase K treatment is essential and can greatly enhance signal. However, overdigestion can degrade tissue integrity and signal intensity. Thus, optimal proteinase K concentration and incubation times should be empirically determined for the particular set of genes being examined. Furthermore, since schistosomes can vary greatly in size (e.g., male worms are larger than female worms) proteinase K treatment regimes should be tailored to the specimen of interest.

Examining Schistosome Germline Development

45

2. Components of the click chemistry development solution should be mixed together fresh for every experiment. The ascorbic acid solution should be made immediately before the experiment and added to the development solution last. 3. We have found that the most convenient way to generate templates for riboprobes and dsRNA is to clone PCR products into plasmid pJC53.2 [23]. This vector allows for convenient TA-based cloning and includes T3 and SP6 promoters flanked by inverted T7 promoters. Thus, a single PCR product generated with T7 primers can be used to produce sense, antisense, and double-stranded RNA depending on the RNA polymerase used. This plasmid is available from addgene (https://www. addgene.org/26536/). For generation of PCR products for in vitro transcription we use Platinum Taq (Invitrogen) to amplify DNA, confirm by agarose gel electrophoresis that a single band of the correct size is amplified, and add specified amount of the PCR to the in vitro transcription reaction. 4. At this concentration, yeast RNA often takes several days to go into solution. Thus, we will start the dissolving process a few days before we wish to begin the RNA purification (e.g., start on Friday if you want to begin on Monday). Gentle heat can also be applied; we typically use a setting of 2–3 on our stirring hot plate. For the purification, it is often convenient to purify enough yeast RNA for several months of experiments; thus we typically start with 6 g of yeast RNA in 600 ml of water. Since these volumes necessitate the use of a large-capacity centrifuge, and typical centrifuge bottles are not compatible with solvents such as chloroform, we use Nalgene FEP 250 ml centrifuge bottles (Fisher Scientific) for all steps with phenol and chloroform. Alternatively, steps with organic solvents can be performed using appropriate 50 ml conical tubes. 5. For fluorescence in situ hybridization, as an alternative to the anti-digoxigenin-AP antibody the anti-digoxigenin-POD antibody (1:1000) (Roche) can be used. Following antibody incubation and washes, signals can be detected by Tyramide Signal Amplification (TSA Plus, Perkin Elmer) following the manufacturer’s instructions. These samples can be labeled with DAPI and observed by fluorescence microscopy. 6. As a cost-saving measure, consider purifying your own HIStagged T7 RNA polymerase [24]. 7. Alternatively, EdU detection can be performed in combination with fluorescence in situ hybridization. For these approaches perform fluorescence in situ hybridization on EdU-labeled parasites using the modifications detailed in Note 5 and detect EdU after TSA. Make sure to use fluorophores that can be distinguished by fluorescence microscopy (e.g., FITC and Cy3).

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8. Due to the large number of wash steps involved in the in situ hybridization procedure a great deal of time can be saved by making or purchasing mesh baskets for washing. These baskets can be easily fabricated with a small plastic tube, nylon mesh, and a hotplate [25]. After pipetting the parasites in these baskets, washes can be performed by simply moving the baskets to separate wells on a multiwell plate. Alternatively, Intavis Bioanalytical Instruments sells an instrument that can automate the entire in situ hybridization process. 9. As the dsRNA is synthesized the reaction will often become viscous. 10. Under these non-denaturing conditions we find that dsRNA often migrates roughly similar to dsDNA. However, we occasionally note additional high-molecular-weight bands. 11. Many types of commercially available cell culture media have been used to culture S. mansoni including DMEM, RPMI, and M199 [18]. For adult S. mansoni we have had success with Basch Media 169 that was specifically developed for culturing larval schistosomes [26]. 12. We have seen phenotypic consequences of dsRNA treatment as early as day 5. But some phenotypes take 2–3 weeks to manifest. Thus, verification of mRNA and/or protein depletion is key to determining the length of the dsRNA treatment regime. References 1. Chitsulo L, Engels D, Montresor A, Savioli L (2000) The global status of schistosomiasis and its control. Acta Trop 77(1):41–51, doi: S0001-706X(00)00122-4 [pii] 2. van der Werf MJ, de Vlas SJ, Brooker S, Looman CW, Nagelkerke NJ, Habbema JD, Engels D (2003) Quantification of clinical morbidity associated with schistosome infection in subSaharan Africa. Acta Trop 86(2-3):125–139 3. Basch PF (1991) Schistosomes: development, reproduction, and host relations. Oxford University Press, New York 4. Pearce EJ, MacDonald AS (2002) The immunobiology of schistosomiasis. Nat Rev Immunol 2(7):499–511. doi:10.1038/nri843 5. Skinner DE, Rinaldi G, Koziol U, Brehm K, Brindley PJ (2014) How might flukes and tapeworms maintain genome integrity without a canonical piRNA pathway? Trends Parasitol 30(3):123–129. doi:10.1016/j.pt.2014.01. 001 6. Skinner DE, Rinaldi G, Suttiprapa S, Mann VH, Smircich P, Cogswell AA, Williams DL, Brindley PJ (2012) Vasa-like DEAD-Box RNA

helicases of Schistosoma mansoni. PLoS Negl Trop Dis 6(6):e1686. doi:10.1371/journal. pntd.0001686, PNTD-D-12-00056 [pii] 7. Tsai IJ, Zarowiecki M, Holroyd N, Garciarrubio A, Sanchez-Flores A, Brooks KL, Tracey A, Bobes RJ, Fragoso G, Sciutto E, Aslett M, Beasley H, Bennett HM, Cai J, Camicia F, Clark R, Cucher M, De Silva N, Day TA, Deplazes P, Estrada K, Fernandez C, Holland PW, Hou J, Hu S, Huckvale T, Hung SS, Kamenetzky L, Keane JA, Kiss F, Koziol U, Lambert O, Liu K, Luo X, Luo Y, Macchiaroli N, Nichol S, Paps J, Parkinson J, PouchkinaStantcheva N, Riddiford N, Rosenzvit M, Salinas G, Wasmuth JD, Zamanian M, Zheng Y, Taenia solium Genome C, Cai X, Soberon X, Olson PD, Laclette JP, Brehm K, Berriman M (2013) The genomes of four tapeworm species reveal adaptations to parasitism. Nature 496 (7443):57–63. Doi: 10.1038/nature12031 8. Wang Y, Stary JM, Wilhelm JE, Newmark PA (2010) A functional genomic screen in planarians identifies novel regulators of germ cell development. Genes Dev 24(18):2081–2092. doi:10.1101/gad.1951010, 24/18/2081 [pii]

Examining Schistosome Germline Development 9. Newmark PA, Wang Y, Chong T (2008) Germ cell specification and regeneration in planarians. Cold Spring Harb Symp Quant Biol 73:573–581. doi:10.1101/sqb.2008.73.022, sqb.2008.73.022 [pii] 10. Collins JJ III, King RS, Cogswell A, Williams DL, Newmark PA (2011) An atlas for Schistosoma mansoni organs and life-cycle stages using cell type-specific markers and confocal microscopy. PLoS Negl Trop Dis 5(3):e1009. doi:10. 1371/journal.pntd.0001009 11. Collins JJ III, Wang B, Lambrus BG, Tharp ME, Iyer H, Newmark PA (2013) Adult somatic stem cells in the human parasite Schistosoma mansoni. Nature 494(7438):476–479. doi:10.1038/nature11924 12. Beckmann S, Buro C, Dissous C, Hirzmann J, Grevelding CG (2010) The Syk kinase SmTK4 of Schistosoma mansoni is involved in the regulation of spermatogenesis and oogenesis. PLoS Pathog 6(2):e1000769. doi:10.1371/journal. ppat.1000769 13. Quack T, Knobloch J, Beckmann S, Vicogne J, Dissous C, Grevelding CG (2009) The forminhomology protein SmDia interacts with the Src kinase SmTK and the GTPase SmRho1 in the gonads of Schistosoma mansoni. PLoS One 4(9):e6998. doi:10.1371/journal.pone. 0006998 14. Hahnel S, Lu Z, Wilson RA, Grevelding CG, Quack T (2013) Whole-organ isolation approach as a basis for tissue-specific analyses in Schistosoma mansoni. PLoS Negl Trop Dis 7(7):e2336. doi:10.1371/journal.pntd. 0002336 15. Cogswell AA, Collins JJ III, Newmark PA, Williams DL (2011) Whole mount in situ hybridization methodology for Schistosoma mansoni. Mol Biochem Parasitol 178(12):46–50. doi:10.1016/j.molbiopara.2011. 03.001, S0166-6851(11)00091-0 [pii] 16. Pearson BJ, Eisenhoffer GT, Gurley KA, Rink JC, Miller DE, Sa´nchez Alvarado A (2009) Formaldehyde-based whole-mount in situ hybridization method for planarians. Dev Dyn 238(2):443–450. doi:10.1002/dvdy. 21849 17. King RS, Newmark PA (2013) In situ hybridization protocol for enhanced detection of gene expression in the planarian Schmidtea mediterranea. BMC Dev Biol 13:8. doi:10.1186/ 1471-213X-13-8

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18. Stefanic S, Dvorak J, Horn M, Braschi S, Sojka D, Ruelas DS, Suzuki B, Lim KC, Hopkins SD, McKerrow JH, Caffrey CR (2010) RNA Interference in Schistosoma mansoni schistosomula: selectivity, sensitivity and operation for largerscale screening. PLoS Negl Trop Dis 4(10): e850. doi:10.1371/journal.pntd.0000850 19. Skelly PJ, Da’dara A, Harn DA (2003) Suppression of cathepsin B expression in Schistosoma mansoni by RNA interference. Int J Parasitol 33(4):363–369, doi:S00207519030 00304 [pii] 20. Boyle JP, Wu XJ, Shoemaker CB, Yoshino TP (2003) Using RNA interference to manipulate endogenous gene expression in Schistosoma mansoni sporocysts. Mol Biochem Parasitol 128 (2):205–215, doi: S0166685103000781 [pii] 21. Rouhana L, Weiss JA, Forsthoefel DJ, Lee H, King RS, Inoue T, Shibata N, Agata K, Newmark PA (2013) RNA interference by feeding in vitro-synthesized double-stranded RNA to planarians: methodology and dynamics. Dev Dyn 242(6):718–730. doi:10.1002/dvdy. 23950 22. Lewis FA, Liang YS, Raghavan N, Knight M (2008) The NIH-NIAID schistosomiasis resource center. PLoS Negl Trop Dis 2(7): e267. doi:10.1371/journal.pntd.0000267 23. Collins JJ III, Hou X, Romanova EV, Lambrus BG, Miller CM, Saberi A, Sweedler JV, Newmark PA (2010) Genome-wide analyses reveal a role for peptide hormones in planarian germline development. PLoS Biol 8(10) 24. He B, Rong M, Lyakhov D, Gartenstein H, Diaz G, Castagna R, McAllister WT, Durbin RK (1997) Rapid mutagenesis and purification of phage RNA polymerases. Protein Expr Purif 9(1):142–151. doi:10.1006/prep.1996.0663, S1046-5928(96)90663-4 [pii] 25. Sive HL, Grainger RM, Harland RM (2007) Baskets for in situ hybridization and immunohistochemistry. CSH Protoc 2007: pdb prot4777. doi: 10.1101/pdb.prot4777 26. Basch PF (1981) Cultivation of Schistosoma mansoni in vitro. I Establishment of cultures from cercariae and development until pairing. J Parasitol 67(2):179–185 27. Collins JJ III, Newmark PA (2013) It’s no fluke: the planarian as a model for understanding schistosomes. PLoS Pathog 9(7):e1003396. doi:10.1371/journal.ppat.1003 396

Chapter 3 Evaluation of the Asymmetric Division of Drosophila Male Germline Stem Cells Mayu Inaba and Yukiko M. Yamashita Abstract Asymmetric cell division (ACD) is utilized in many stem cell systems to produce two daughter cells with different cell fates. Despite the fundamental importance of ACD during development and tissue homeostasis, the nature of ACD is far from being fully understood. Step-by-step observation of events during ACD allows us to understand processes that lead to ACD. Here we describe examples of how we evaluate ACD in vivo using the Drosophila male germline stem cell system. Key words Asymmetric cell division (ACD), Drosophila male germline stem cells (GSCs), Centrosome orientation, Spindle orientation, Centrosome orientation checkpoint (COC)

1

Introduction Drosophila male germline stem cells (GSCs) divide asymmetrically to produce one GSC and one differentiating cell, the gonialblast (GB) (Fig. 1a). Two daughter cells of a GSC division are stereotypically positioned with respect to the niche: only GSCs are attached to the niche component, the hub, but not GBs (Fig. 1a). The hub, a cluster of postmitotic somatic cells, is located at the tip of the testis and surrounded by GSCs. Hub cells secrete ligands that are required for GSC maintenance. Ectopic expression of a niche ligand, Unpaired (Upd), results in an expansion of GSC-like cells outside the niche, indicating that Upd plays a major role in GSC specification. In addition to the extrinsic fate determinants provided by the niche, a number of intrinsic factors and organelles have been reported to be deposited asymmetrically between GSCs and GBs, upon division (Fig. 1a), indicating that these intrinsic factors may also contribute to the asymmetric outcome of GSC divisions. For example, centrosomes, sister chromatids, midbody rings, and histone H3 are reported to segregate asymmetrically during GSC divisions [1–4].

Michael Buszczak (ed.), Germline Stem Cells, Methods in Molecular Biology, vol. 1463, DOI 10.1007/978-1-4939-4017-2_3, © Springer Science+Business Media New York 2017

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a

Hub GSC (Germline stem cell) Spindle orienation

Centrosome orientation

GB (Gonialblast)

Old histone H3 New histone H3 Sister chromatids of XY chromosomes

Stereotypical positioning of two daughter cells

Midbody ring Mother centrosome Daughter centrosome

b

DAPI Vasa

b’

DAPI

b”

Vasa

Fig. 1 Asymmetric division of the Drosophila male germline stem cells. (a) A schematic of asymmetric GSC division and asymmetrically segregating cellular factors. (b) A confocal microscope section of a Drosophila testis tip region. The hub can be identified as a tightly packed cell cluster as shown using DAPI (encircled by dotted cyan line in b0 ). Five germline stem cells (vasa positive, b00 ) directly attached to the hub are present in this z-section encircled by yellow dotted lines in (b). Scale bar: 10 μm

For example, the mother centrosome is stereotypically inherited by male GSCs by consistently staying near the hub-GSC junction. Similarly, mouse radial glial progenitor cells inherit the mother centrosome [5]. In contrast, Drosophila neuroblasts [6] and female GSCs [3] inherit the daughter centrosomes. These studies indicate the universality of the stereotypical centrosome inheritance during ACD. Asymmetric centrosome inheritance has been reported to correlate with asymmetric cytokinesis. The midbody ring (MR), the remnant of the cytokinetic contractile ring, is segregated to one of the two daughter cells in a stereotypical manner in several cell types that undergo asymmetric cell division. The role of the MR in cell fate specification remains elusive: however, MRs have been shown

Asymmetric Stem Cell Division in Drosophila Testis

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to accumulate preferentially in induced pluripotent stem (iPS) cells and cancer stem cells [7, 8], whereas another study showed that stem cells tend to release the MR in various cell lines [9]. In the Drosophila testes, the MR is inherited by the differentiating gonialblast, whereas in ovaries, the MR is inherited by the GSCs. In both cases, the cells that inherit the MR are the ones that inherit the daughter centrosomes [3], as opposed to the reported case of MR being inherited by the mother centrosome-containing cells in human embryonic stem cells and cancer cell lines [8]. Although the biological significance of asymmetric MR inheritance is not well understood beyond correlative relationships, it is tempting to speculate that the MR contributes to the asymmetric cell fate directly or indirectly. Recently, it was reported that the Drosophila GSCs selectively inherit preexisting histone 3 (H3), whereas newly synthesized H3 is enriched in the differentiating gonialblasts [4, 10], implying the possibility that distinct epigenetic information might be asymmetrically segregated during male GSC ACD to confer distinct cell fates. These studies suggest the presence of intricate mechanisms to asymmetrically segregate cellular components, possibly contributing to ACD. Yet, such asymmetries associated with ACD have only begun to be revealed and it remains unclear how critical these asymmetries are in determining asymmetric cell fates. Importantly, it is likely that these asymmetries collectively contribute to asymmetric fates/behaviors of daughter cells (stem cells vs. differentiating cells) with each asymmetry regulating a limited aspect of the cells’ fate/behavior. Thus, failure of establishing one particular asymmetry may not result in readily detectable defects such as a stem cell tumor or depletion, and may not be appreciable without specific markers or sensitized background. Accordingly, investigation of ACD involves careful observation to detect subtle changes to fully understand the significance of each asymmetry in ACD. Such subtle changes caused by defective ACD may underlie the complexity of human pathologies, making it difficult to pinpoint a single cause of a disease. Here, we describe how to assess asymmetries during male GSC ACD, which can serve as a platform to identify potentially many more novel asymmetries. These methods are only to showcase known asymmetries/polarities, and to provide basic methods to detect them. Discovery of new asymmetries and their functions would require the development of new ideas, methods, and reagents. First, we summarize the method of immunofluorescence staining and antibody combinations that can be used to assess distinct aspects of ACD. Later, we also provide a method to observe unfixed testes for live observation. The description of these methods will be accompanied by rationales behind them.

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Materials

2.1 Preparation of Sample

1. Fixative 4 % formaldehyde in PBS: made from 16 % formaldehyde. PBS: phosphate-buffered saline (NaCl 8 g; KCl 2 g; NaHPO4 1.46 g; KH2PO4 0.24 g/1 l water, pH 7.4). 2. PBST: PBS (Phosphate-buffered saline) supplemented with 0.1 % Triton X-100. 3. 3 % Bovine serum albumin heat-shock fraction, pH 7, 98 %, in PBST. 4. VECTASHIELD (Vector Laboratories #H-1200) with 40 ,6diamidino-2-phenylindole (DAPI). 5. Schneider’s insect medium with L-glutamine (Sigma-Aldrich #S0146) supplemented with 10 % heat-inactivated fetal bovine serum (FBS). 6. Microscope slides. 7. Cover slips. 8. Gold Seal™ Rite-On™ Micro Slides two etched rings (Live imaging).

2.2

Useful Antibodies

1. Centrosomal marker; mouse anti-γ-tubulin (GTU-88, 1:500 dilution; Sigma). 2. Germ cell marker; rabbit anti-Vasa (1:200 dilution; Santa Cruz Biotechnology Inc.) or rat anti-Vasa (1:20 dilution; Developmental Studies Hybridoma Bank, DSHB). 3. Hub cell marker; mouse anti-Fasciclin III (FasIII, 7G10) (1:40 dilution; DSHB) (see Note 1). 4. Mitosis marker phospho-histone H3 (PH3); rabbit anti-phospho-Histone H3 (Thr3) (1:200 dilution; Upstate, Millipore) or rabbit phospho-Histone H3 (Ser10) (1:500 dilution; Cell Signaling Technology) (see Note 2). 5. AlexaFluor-conjugated secondary antibodies (1:400 dilution; Molecular Probes).

3 3.1

Methods Fly Culture

Flies are raised on standard Bloomington medium at 25  C and assayed at day 0 to day 2 after eclosion. Low nutrient condition or aging has been shown to result in high frequency of centrosome misorientation [11, 12].

Asymmetric Stem Cell Division in Drosophila Testis

3.2 Whole-Mount Immunohistochemistry of Drosophila Testes

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1. Dissect testes in PBS (or Schneider’s insect medium if followed by in vitro treatments) and collect them in a tube with 1 ml of fixative at room temperature (RT), within 30 min of dissection. 2. Gently rock the tubes on a nutator for an additional 30 min at RT. 3. Briefly rinse fixed testes three times in 1 ml of PBST. 4. Gently rock testes in 1 ml of PBST for 30 min to 1 h at RT. 5. Remove PBST, add primary antibody solution (antibodies appropriately diluted in a total of 100 μl 3 % BSA in PBST), and gently rock at 4  C overnight. 6. Remove antibody solution and add 1 ml of PBST. Gently rock the tubes for 20 min. Repeat three times. 7. Incubate with diluted secondary antibodies in 3 % BSA in PBST and gently rock at RT for 2–4 h or at 4  C overnight. 8. Remove antibody solution and add 1 ml of PBST. Gently rock the tubes for 20 min. Repeat three times. 9. Mount testes with 1 drop of VECTASHIELD with DAPI. Remove excess amount of VECTASHEILD as much as possible by gentry applying paper towel from the top of the cover slip. An excess amount of mounting media will result in low-quality imaging. 10. Scoring has to be performed using a confocal microscope (see below for scoring criteria). We use a Zeiss LSM700 confocal microscope with a 40 oil-immersion objective (NA ¼ 1.4) or a Leica TCS SP5 or SP8 confocal microscope with a 63 oilimmersion objective (NA ¼ 1.4).

3.3 Scoring Centrosome Orientation

Immunofluorescence staining of testes using primary antibody combinations γ-tubulin, Vasa, and Fas III can be used to score the centrosome orientation in GSCs (see Note 3). Throughout the male GSC cell cycle, the mother centrosome remains close to the hub-GSC junction. Once the centrosome is duplicated, the newly formed daughter centrosome moves towards the distal side of the cell [13]. This stereotypical centrosome orientation (the mother being at the apical side) is normally observed in more than 90 % of GSCs during interphase in wild-type testes (Fig. 2a) [13]. We set the criterion for scoring of “centrosome orientation” based on the apical centrosome location relative to the hub-GSC junction as shown in Fig. 2. GSCs are scored as having “oriented centrosomes” when at least one of the two centrosomes is in the pink area (Fig. 2c). When neither of the two centrosomes is closely associated with hub-GSC junction, it is defined as “centrosome misorientation” (Fig. 2b) [11]. Because of this scoring criterion, even “completely random” centrosome orientation will result in only up to 56.25 % “misorientation” (calculated as the probability in which both the centrosomes are

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Fig. 2 Centrosome positioning of Drosophila male GSCs. (a) Interphase GSCs attached to the hub (white dotted circles) with oriented centrosomes (arrowheads). (b) An example of a GSC (circle) with misoriented centrosomes (arrowheads). (c) Criterion of orientation for scoring. Centrosomes are scored as “oriented” when one of the two centrosomes is in the pink area close to the hub-GSC junction (orange line). (d) An example of a mitotic GSC (circle) with an oriented spindle (arrowheads indicate centrosomes, arrows indicate pH3-positive chromosome). (e) Two mitotic GSCs (circles) with misoriented spindles in a testis. Hub is indicated by an asterisk (*). Scale bar: 10 μm

located outside of the pink area, 0.75  0.75 ¼ 0.5625 assuming that the pink area represents ~25 % of the entire GSC surface area). In our experience, the highest centrosome misorientation frequency was about 50 % of GSCs, which closely matches the highest possible centrosome misorientation frequency based on the model. The low frequency of centrosome misorientation in wild type (70–80 % of GSCs can be scored. 3.4 Scoring Spindle Orientation

Immunofluorescence staining of testes using the primary antibody combination γ-tubulin, Vasa, Fas III, and PH3 can be used to score spindle orientation in GSCs. Mitotic GSCs can be identified by anti-PH3 staining (Fig. 2d, e). Mitotic GSCs are found at a very low frequency under normal conditions (~2 GSCs per 10 testes). Importantly, CO2 anesthetization longer than 7 min dramatically decreases the mitotic index; therefore it is recommended to dissect testes within 7 min. In addition, it is necessary to let flies recover for at least 2 h before dissection if they must be first sorted based on genotypes. Spindle orientation can be judged by the same criterion as centrosome orientation according to apical centrosome/spindle pole position of mitotic GSCs (Fig. 2c).

3.5 Evaluating the Centrosome Orientation Checkpoint

Centrosome orientation checkpoint (COC) can be scored by immunofluorescenceCentrosome orientation checkpoint (COC) staining of testes using the primary antibody combination Vasa, PH3, and Fas III. Additional steps: 1. Dissect testes and transfer to Schneider’s insect medium. After sample collection, add colcemid (100 μM final concentration, Calbiochem) and incubate at 25  C for 4.5 h. 2. Briefly rinse with PBS three times remove PBS and add 1 ml of fixative. 3. Continue to step 2 in Subheading 3.2. 4. Score the frequency of mitotic GSCs, gonialblasts (GBs), and spermatognia (SG) (number of mitotic GSC, GB, or SG per testis). SGs are scored per stage (i.e., 2-cell, 4-cell, 8-cell SG). Although the mitotic index defined as “mitotic GSCs/total GSCs” (and mitotic 2-cell SG/total 2-cell SG, and so on) would be more accurate, this requires scoring total numbers of GSCs, GBs, and SGs for each testis, which is very time consuming. Thus, we typically count the number of GSCs/ GBs/SGs using fewer testes (than required for scoring mitotic indexes), and if the numbers are not dramatically changed, we use mitotic cells/testis as the mitotic index. We have shown that colcemid-induced depolymerization of microtubules leads to centrosome misorientation, which causes COC activation and thus G2 cell cycle arrest in GSCs [15].

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a

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Fig. 3 A technique to detect defective COC activity of GSC by colcemid treatment. (a) Colcemid treatment induces centrosome misorientation in GSC. (a0 ) GBs or SG does not possess COC. (b, c) A typical mitotic index pattern of colemid-treated testes. Number of mitotic cells per testes increases under the presence of colcemid when COC is inactive (wild-type GB ~ SG in b or mutant GSC defective of COC in c)

In contrast, GSCs defective in COC, as well as spermatogonia, which do not possess COC, enter mitosisGonialblasts (GBs), and are then arrested due to activation of the spindle assembly checkpoint (Fig. 3a, a0 ). Since colcemid treatment artificially increases centrosome misorientation, and arrests COC-proficient GSCs in G2 phase, and COC-defective GSCs in mitosis, respectively (Fig. 3a), it provides a sensitive and robust method to detect defective COC. For example, after 4.5 h of colcemid treatment of testes defective in COC (such as those from par-1 or cnn mutants) >1 GSC per testis was found to be arrested in mitosis (Fig. 3c). This is in stark contrast to wild-type GSCs, which shows about 0.2 mitotic GSCs per testis (Fig. 3b). The mitotic index of SG from the same testes serves as a positive control of spindle assembly checkpoint-dependent mitotic arrest. 3.6 Generation of Heat-ShockInduced Germline Clones

1. Cross hs-FLP, nos-FRT-stop-polyA-FRT-gal4, UAS-GFP [16] females with UAS-RNAi males for experimental or wild type males for control crosses. 2. Heat shock progeny at days 0–2 after eclosion.

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3. Dissect testes at various time points (for example, 3, 7, 14, 21 days) after heat shock. To identify the GSC clones, immunofluorescence staining of the testes using the primary antibody combination Vasa and Fas III can be used together with GFP. Although other genetic schemes to induce GSC clones can be used for the same purpose, the use of hs-FLP, nos-FRT-stoppolyA-FRT-gal4, UAS-GFP has two unique advantages: (1) clonal induction is limited to germ cells, and thus there will be no complication due to clonal induction in somatic gonadal cells (cyst cells and hub cells). (2) Any UAS transgenes can be introduced into this genetic scheme easily, and thus the effect of gene overexpression or RNAi-mediated knockdown on ACD can be easily tested (see below for more detail) [14]. Generation of mosaic clones in an otherwise wild-type background has served to investigate cell autonomy of gene function. Also, mosaic clones can be used to assess gene function in adult tissues, even when homozygous animals are lethal at earlier developmental stages. It has been shown that GSCs compete for niche occupancy, which can generate apparently inconsistent results, depending on whether a mutant exists as a mosaic clone or homogenous population of GSCs. For example, GSC clones mutant for JAK-STAT signaling components (such as Hop, Dome, STAT92E) are quickly lost from the niche, leading the authors to suggest that these genes are essential for GSC self-renewal [17, 18]. However, it was later shown that GSCs mutant for STAT can be well maintained if all of GSCs are mutant for STAT [19]. These results revealed the necessity of addressing gene functions using multiple experimental paradigms. In this context, an experimental caveat is the fact that the GSC clones are typically generated by FLP-mediated mitotic recombination of homologous chromosomes using mutant alleles, whereas nos-gal4-mediated RNAi is used to deplete the gene product from all GSCs. When inconsistent results are obtained between FRT clones of mutant alleles and nos-gal4 > UAS-RNAi, it is unclear whether it is due to the gene’s requirement in GSC competition (but not self-renewal) or different strengths of the allele vs. RNAi used. By using hs-FLP, nos-FRT-stop-polyA-FRT-gal4, UAS-GFP, the maintenance of RNAi clones can be directly compared to “global knockdown” of the gene (nos-gal4 > UASRNAi), and the gene’s requirement in GSC competition can be unambiguously tested. 3.7 Assessment of Asymmetric vs. Symmetric Stem Cell Division

Although spindle misorientation may lead to symmetric GSC divisions to generate two GSCs, spindle misorientation is not an accurate predictor of symmetric outcome. It has been reported that gonialblast can crawl back to the niche, acquiring hub attachment to yield two GSCs, even after GSCs have undergone mitosis with correctly oriented spindles [20]. Conversely, spindle misorientation

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does not necessarily lead to symmetric self-renewal, since one daughter of GSC division may eventually lose hub attachment and leave the niche. Therefore, it is important to assess asymmetric vs. symmetric GSC divisions, independently of spindle orientation. By following marked GSC clones, the frequency of symmetric division can be inferred. By using the transgenic line described above (hs-FLP, nos-FRT-stop-polyA-FRT-gal4, UAS-GFP [19]), the asymmetric vs. symmetric outcome of GSC division can be assessed. Upon heat-shock treatment, only a subset of GSCs undergo “FLP out” of the “FRT-stop-polyA-FRT” cassette, thereby expressing nos-gal4, UAS-GFP. Heat-shock treatments can be adjusted so that only one GSC per testis is a clone on average. Such a singlet clone can convert to a doublet if GSCs undergo symmetric self-renewal (Fig. 4). By using this method, we estimated that wild-type GSCs undergo symmetric self-renewal about 2 % of the time [16]. Although accurate frequency of symmetric self-renewal is obtained via mathematical modeling [16] by taking other parameters (such as the probability of two neighboring GSCs undergoing clonal induction independently) into account [16], high frequency of symmetric divisions can be easily detected by this method [14]. Prolonged chase time (>24 h) often leads to more variation of clone frequency possibly due to other parameters than symmetric division (e.g., cell adhesion, cell cycle difference). Thus, for detection of symmetric self-renewal, data collection at 24 h after heat-shock treatment is recommended. Immunofluorescence staining of testes with primary antibody Vasa and Fas III together with detecting GFP-positive clone can be used to score the symmetric outcome of GSC division. 3.8 Microscopy of Living Tissue

1. Dissect testes and transfer to Schneider’s insect medium. 2. Add vital dyes as necessary and incubate for desired time. If using the membrane dye, FM4-64FX Lipophilic Styryl Dye (5 μg/ml, Molecular Probes), add the dye 1 min prior to analysis and perform imaging within 15 min. (Sample can be rinsed with PBS after dye incorporation if longer imaging is necessary. FM4-64FX is also fixable.) If using the organelle dye Lysotracker or the DNA dye Hoechst 33342, then dissect the testes in PBS, incubate with Lysotracker (DND-99; Invitrogen) or Hoechst 33342 (2 μg/ml) for 30 min, and rinse with PBS twice prior to imaging. 3. Mount on the slide glasses with two etched rings (see Subheading 2) and cover with a cover slip. 4. Start imaging immediately. In most cases, we perform imaging within 30 min. See [21] for prolonged live imaging. Whole living Drosophila testis can be observed under the conventional confocal microscope without isolating or fixing the cells.

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a ~2% Symmetric self-renewal

Asymmetric division

96%~

Symmetric differentiation

~2%

Vasa (Germ cells), GFP

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singlet

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Fig. 4 Examining symmetric GSC divisions. (a) GFPþ clone is induced at low frequency ( UAS-GFP-αTubulin) (Fig. 5b–d) [22]. MTnanotubes are fragile structures and have escaped our detection in

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fixed samples. As revealed by this example, microscopic structures can be missed due to sample processing, and the imaging of unfixed tissue will help identify such structures (see Notes 4 and 5).

4

Notes 1. FasIII usage is optional. Once eyes are trained, the hub is easily distinguishable by using DAPI staining as a cluster of ~15–20 cells with tightly packed nuclei and small amount of cytoplasm at the testis apical tip (Fig. 1b). 2. Histone H3 is phosphorylated at Thr-3 by haspin during prophase and dephosphorylated during anaphase. Phosphorylation at Ser-10 is maintained until anaphase/telophase and thus more suitable as a marker to detect all stages of mitosis. 3. GSCs can be identified as Vasa-positive germ cells directly attached to the hub (Figs. 1b and 2a). Typically 8–10 GSCs are found in each testis. GSCs are only ~7 μm in diameter, and the study of ACD involves detection of subcellular structure within GSCs. Furthermore, the study of ACD in the tissue context inevitably involves imaging of multiple cells (with many cells being positioned along the z-axis). In particular, for scoring of centrosome orientation (see below), the signal from neighboring cells (below/above the cells of interest) can be confusing. Accordingly, a microscope with sufficient resolution is required, and it will take some training to be able to judge localization of subcellular structures within cells, distinguishing them from signals from neighboring cells. 4. Later, we developed a fixation technique to preserve MTnanotubes by adding a low concentration of taxol (1 μM) to the fixative (Fig. 5d). 5. 3D rendering data processing further helps understand geometric relationships among cells and subcellular structures. Figure 5c, d shows examples of MT-nanotubes extended from a GSC invaginate into hub cells. Images were processed with Imaris software (Bitplane) to reconstitute z-stacks.

Acknowledgement The research in the Yamashita laboratory is supported by the Howard Hughes Medical Institute and the National Institute of Health (R01 GM118308). YMY is supported by the John D. and Catherine T. MacArthur Foundation.

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References 1. Yamashita YM, Mahowald AP, Perlin JR et al (2007) Asymmetric inheritance of mother versus daughter centrosome in stem cell division. Science 315:518–521 2. Yadlapalli S, Yamashita YM (2013) Chromosome-specific nonrandom sister chromatid segregation during stem-cell division. Nature 498:251–254 3. Salzmann V, Chen C, Chiang C-YA et al (2014) Centrosome-dependent asymmetric inheritance of the midbody ring in Drosophila germline stem cell division. Mol Biol Cell 25:267–275 4. Tran V, Lim C, Xie J et al (2012) Asymmetric division of Drosophila male germline stem cell shows asymmetric histone distribution. Science 338:679–682 5. Wang XQ, Tsai JW, Imai JH et al (2009) Asymmetric centrosome inheritance maintains neural progenitors in the neocortex. Nature 461:947–955 6. Januschke J, Llamazares S, Reina J et al (2011) Drosophila neuroblasts retain the daughter centrosome. Nat Commun 2:243 7. Gromley A, Yeaman C, Rosa J et al (2005) Centriolin anchoring of exocyst and SNARE complexes at the midbody is required for secretory-vesicle-mediated abscission. Cell 123:75–87 8. Kuo T-C, Chen C-T, Baron D et al (2011) Midbody accumulation through evasion of autophagy contributes to cellular reprogramming and tumorigenicity. Nat Cell Biol 13:1214–1223 9. Ettinger AW, Wilsch-Br€auninger M, Marzesco A-M et al (2011) Proliferating versus differentiating stem and cancer cells exhibit distinct midbody-release behaviour. Nat Commun 2:503 10. Xie J, Wooten M, Tran V et al (2015) Histone H3 threonine phosphorylation regulates asymmetric histone inheritance in the drosophila male germline. Cell 163:920–933 11. Cheng J, Turkel N, Hemati N et al (2008) Centrosome misorientation reduces stem cell division during ageing. Nature 456:599–604

12. Roth TM, Chiang C-YYA, Inaba M et al (2012) Centrosome misorientation mediates slowing of the cell cycle under limited nutrient conditions in Drosophila male germline stem cells. Mol Biol Cell 23:1524–1532 13. Yamashita YM, Jones DL, Fuller MT (2003) Orientation of asymmetric stem cell division by the APC tumor suppressor and centrosome. Science 301:1547–1550 14. Inaba M, Venkei ZG, Yamashita YM (2015) The polarity protein Baz forms a platform for the centrosome orientation during asymmetric stem cell division in the Drosophila male germline. eLife 4:e04960 15. Venkei ZG, Yamashita YM (2015) The centrosome orientation checkpoint is germline stem cell specific and operates prior to the spindle assembly checkpoint in Drosophila testis. Development 142:62–69 16. Salzmann V, Inaba M, Cheng J et al (2013) Lineage tracing quantification reveals symmetric stem cell division in drosophila male germline stem cells. Cell Mol Bioeng 6:441–448 17. Kiger AA, Jones DL, Schulz C et al (2001) Stem cell self-renewal specified by JAK-STAT activation in response to a support cell cue. Science 294:2542–2545 18. Tulina N, Matunis E (2001) Control of stem cell self-renewal in Drosophila spermatogenesis by JAK-STAT signaling. Science 294:2546–2549 19. Leatherman JL, Dinardo S (2010) Germline self-renewal requires cyst stem cells and stat regulates niche adhesion in Drosophila testes. Nat Cell Biol 12:806–811 20. Sheng XR, Matunis E (2011) Live imaging of the Drosophila spermatogonial stem cell niche reveals novel mechanisms regulating germline stem cell output. Development 138:3367–3376 21. Cheng J, Hunt AJ (2009) Time-lapse live imaging of stem cells in Drosophila testis. Current Protoc Stem Cell Biol Chapter 2: Unit 2E.2 22. Inaba M, Buszczak M, Yamashita YM (2015) Nanotubes mediate niche-stem-cell signalling in the Drosophila testis. Nature 523:329–332

Chapter 4 Live Imaging of the Drosophila Testis Stem Cell Niche Leah J. Greenspan and Erika L. Matunis Abstract Live imaging of adult tissue stem cell niches provides key insights into the dynamic behavior of stem cells, their differentiating progeny, and their neighboring support cells, but few niches are amenable to this approach. Here we discuss a technique for long-term live imaging of the Drosophila testis stem cell niche. Culturing whole testes ex vivo for up to 12.5 h allows for tracking of cell-type specific behaviors under normal and various chemically or genetically modified conditions. Fixing and staining tissues after live imaging allows for the molecular confirmation of cell identity and behavior. Utilization of live imaging in intact niches will facilitate further understanding of the cellular and molecular mechanisms that regulate stem cell function in vivo. Key words Drosophila testis, Niche, Stem cell, Fluorescence, Time-lapse live imaging, Confocal microscopy

1

Introduction Adult stem cells both self-renew and differentiate in order to regulate proper tissue maintenance. While the analysis of fixed tissues provides important information regarding adult stem cell biology, live imaging of stem cells within an intact local microenvironment, or niche, reveals unique insights into stem cell behavior that cannot always be extrapolated from fixed images. For example, live imaging of the Drosophila testis niche has shown that although stem cells primarily undergo asymmetric divisions where one daughter cell remains a stem cell while the other daughter goes on to differentiate, germline stem cells can self-renew via symmetric renewal (in which two stem cells are made) and dedifferentiation (in which differentiated daughters migrate back to the niche and revert to stem cells) [1]. Spermatogonial fragmentation has since been shown to occur in mammalian testes using live imaging and lineage tracing [2]. The well-characterized Drosophila testis stem cell niche provides an ideal model system to study stem cell behavior and

Michael Buszczak (ed.), Germline Stem Cells, Methods in Molecular Biology, vol. 1463, DOI 10.1007/978-1-4939-4017-2_4, © Springer Science+Business Media New York 2017

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Cyst Cell CySC

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Fig. 1 The Drosophila testis stem cell niche consists of terminally differentiated quiescent hub cells (red) that signal to the adjacent germline (GSCs, green) and somatic cyst stem cells (CySCs, white). GSCs divide asymmetrically to produce daughter cells (gonialblasts) that transit amplify to form spermatogonial clusters (green). These clusters undergo meiosis and differentiate into sperm. CySCs also divide asymmetrically to produce daughter cyst cells (white). Two cyst cells encase each spermatogonial cluster and are required for the differentiation process

function [3] (Fig. 1). It supports two types of stem cells, germline stem cells (GSCs) and somatic cyst stem cells (CySCs), which attach to a group of terminally differentiated hub cells that secrete ligands promoting stem cell fate. GSCs divide asymmetrically to produce a differentiating daughter (or gonialblast) that undergoes transitamplifying divisions to form a cluster of interconnected spermatogonia. Spermatogonia subsequently enter meiosis and spermiogenesis, eventually forming sperm. CySCs also divide asymmetrically and produce daughters called cyst cells that encapsulate the germline and elongate with the growing spermatogonial clusters. The CySCs and their progeny are essential for the survival and proper differentiation of the germline [3]. Previous work in the Drosophila ovary provided the groundwork for live imaging in Drosophila gonadal tissues [4, 5] including extended live imaging (14 h) of the ovarian stem cell niche [6]. In addition, protocols for imaging testis tips [7] or whole mounted testes on slides [1] have been established. Here we describe a method for imaging the stem cell niche of whole Drosophila testes using glass-bottom imaging dishes. This allows for the manipulation of imaging media and also the capacity to fix and stain testes after long-term imaging. This protocol is updated from that published by Sheng and Matunis with new adaptations developed by our lab and the DiNardo lab [8].

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Materials Prepare all solutions using ultrapure water (sensitivity 16–18 MΩ cm dH2O) and molecular grade reagents unless specified otherwise.

2.1 Dissection of Testes from Adult Male Drosophila

1. Flies expressing fluorescent reporters or proteins in cells of interest (see Note 1). 2. Dumont #5 forceps, blunt and fine tipped pairs. 3. Dissecting dish: Beveled edge watch glass lined with silicone to protect forceps from damage (see Note 2). 4. 900 glass Pasteur pipettes. 5. Stereomicroscope (e.g., Zeiss Stemi SV 6 with a Schott-Fostec external light source). 6. CO2 pad for anesthetizing flies. 7. Round #2 camel hair paintbrush removed.

with most of the hair

8. 1 Becker Ringer’s solution [9]: 111 mM NaCl, 1.88 mM KCl, 64 μM NaH2PO4, 816 μM CaCl2, 2.38 mM NaHCO3. To make a 10 stock solution combine 220 mL of 5 M NaCl, 18.8 mL of 1 M KCl, 3.2 mL of 0.2 M NaH2PO4 l 2H2O, 8.16 mL of 1 M CaCl2 l 2H2O, and 500 mL of ultrapure water in a 1 L graduated cylinder. Add 2 g of NaHCO3 last to prevent precipitation. Add ultrapure water to a final volume of 1 L. Cover graduated cylinder with parafilm and invert 5 times to mix. Filter sterilize through a 0.1 μm PES bottle-tip filter into an autoclaved 1 L glass bottle. Solution keeps many months at room temperature if the bottle is tightly sealed (wrap cap with parafilm). Dilute 1:10 with ultrapure water for 1 Becker Ringer’s solution, store at room temperature. 2.2 Live Imaging of Drosophila Testes

1. Insulin stock solution: Dissolve powdered bovine pancreas insulin in acidified water (1 μL concentrated HCl + 1 mL ultrapure water) at a final concentration of 10 mg/mL. Store 50 μL aliquots in 1.5 mL microcentrifuge tubes at 20  C. 2. Drosophila tissue culture media: 15 % fetal bovine serum (v/v), 0.5 penicillin/streptomycin in Schneider’s media, pH 7. Adjust Schneider’s media to pH 7, sterilize using a 0.1 μm PES syringe filter (this can be stored for many months at 4  C). Combine 2111.25 μL of pH 7 Schneider’s media, 375 μL of fetal bovine serum, and 13.75 μL of 100 penicillin/streptomycin (10,000 U/mL of penicillin and 10,000 μg/ mL streptomycin in 10 mM citrate buffer) in 15 mL sterile conical tube. Vortex briefly, store at 4  C. Make fresh every week.

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3. Live imaging solution: 0.2 mg/mL insulin in Drosophila tissue culture media. Add 10 μL of the 10 mg/mL insulin stock solution to 500 μL of Drosophila tissue culture media. Make fresh on the day of imaging (see Note 3). 4. Poly-L-lysine: 1 mg/mL in 0.1 M Trizma buffer pH 8.5. Dissolve 10 mg of Poly-L-lysine in 10 mL 0.1 M filter-sterilized Trizma buffer pH 8.5. Store 200 μL aliquots in 1.5 mL microcentrifuge tubes at 20  C. 5. Imaging dish: 35-mm glass-bottom Petri dish with 10-mm microwell. 6. Laser scanning or spinning disc confocal microscope (e.g., Zeiss LSM 780 confocal microscope with Zen software or a similar microscope). 2.3 Immunostaining Testes After Live Imaging

1. 900 and 5.7500 glass Pasteur pipettes. 2. 1 Phosphate Buffered Saline (PBS): 137 mM NaCl, 2.7 mM KCl, 1.5 mM KPO4 monobasic, 8.1 mM NaPO4 dibasic anhydrous. To make 10 PBS combine 80 g NaCl, 2 g KCl, 2 g KPO4 monobasic, 11.44 g NaPO4 dibasic anhydrous, and 700 mL of ultrapure water in a 2 L beaker and stir with a magnetic stir bar. When dissolved, transfer solution to a 1 L graduated cylinder and add ultrapure water to a final volume of 1 L. Filter sterilize (PES bottle-tip filter, 0.1 μm pore size) into a 1 L autoclaved glass container and store at room temperature or at 4  C. Dilute 1:10 with ultrapure water to make 1 PBS. 3. 1 Phosphate Buffered Saline with 0.1 % Triton X-100 (v/v) (PBX): Add 1 mL of Triton X-100 to a 2 L beaker containing 1 L 1 PBS. Stir with a magnetic stir bar for 20 min. Store in 1 L glass bottle at room temperature or 4  C. 4. Fixative solution: 4 % paraformaldehyde (v/v) in PBX. Dilute 16 % paraformaldehyde (open a fresh ampule each week) in PBX to a final concentration of 4 %. Make fresh each time. 5. Block solution: 3 % BSA (w/v), 0.02 % NaN3 (w/v) in PBX. Add 15 g bovine serum albumin (BSA) to a 1 L beaker containing 500 mL PBX. Mix with a magnetic stir bar until dissolved. Add 500 μL 20 % sodium azide, filter sterilize (PES filter, 0.1 μm pore size), and store at 4  C. This solution can be used for several months. 6. Normal goat serum. Store 300 μL aliquots in 1.5 mL microcentrifuge tubes at 20  C. Working aliquot can be stored at 4  C. 7. Primary antibodies. 8. Secondary antibodies with conjugated fluorophores. 9. Vectashield mounting medium.

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Methods All methods are performed at room temperature unless stated otherwise.

3.1 Preparing Imaging Dish

1. Thaw 200 μL aliquot of 1 mg/mL Poly-L-Lysine and pipette onto the coverslip portion of the imaging dish. 2. Cover dish and incubate at room temperature on the bench for at least 1 h. 3. Remove Poly-L-Lysine from coverslip with a micropipettor and return to the original tube. Poly-L-Lysine can be stored at 4  C and reused for 1–2 weeks. 4. Wash coverslip: Add 200 μL of sterile distilled water to coverslip, pipette up and down three times, and discard. Repeat three times. 5. Pipette 200 μL of sterile distilled water onto coverslip, replace the cover on the dish, and keep at room temperature while dissecting testes. Leave water on the coverslip until ready to directly transfer testes.

3.2 Dissecting and Mounting Whole Testes

1. Clean all forceps, dissecting dishes, CO2 pad, and dissecting area with ethanol before dissecting. 2. Anesthetize adult flies using CO2. Using the stereomicroscope, collect 20–50 males with the paintbrush and leave them on the CO2 pad to be dissected. 3. Pour approximately 20 mL 1 Becker Ringer’s solution into dissecting dish. 4. Hold the blunter pair of forceps in your nondominant hand, and use them to grasp a fly by the thorax. Anchor fly to the bottom of the dissecting dish so that it is completely submerged in Becker Ringer’s solution. Without releasing the fly, use the finer pair of forceps in your dominant hand to gently puncture the cuticle near the middle of the fly’s abdomen to partially release the testes into the media. Grasp the posterior cuticle that contains the testes and associated structures, and gently pull until it separates from the rest of the fly (Fig. 2a). Anchor the posterior cuticle to the bottom of the dish using the blunter pair of forceps and gently use one prong of the finer pair of forceps to dissociate the testes from the cuticle. Separate the testes from the accessory glands and ejaculatory duct but not the seminal vesicle, by severing the connection between the seminal vesicle and the accessory gland (see Note 4) (Fig. 2a0 , b, b0 ). Testes

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Fig. 2 Testes dissected from an adult male Drosophila are shown attached to the cuticle (a), completely detached from the cuticle, accessory glands, and ejaculatory duct (a0 ), and mounted on the circular coverslip portion of an imaging dish (a00 ). Yellow arrow (a) indicates a rupture in one of the testes, making it unfit for imaging. Mounted testes (a00 ) are flat on the coverslip and do not overlap. Diagram (b) and image (b0 ) of Drosophila testes, accessory glands, and ejaculatory duct. Testes with seminal vesicles attached should be separated from the accessory glands and ejaculatory duct (red arrow) prior to imaging

can be left in the Becker Ringer’s solution in the dissecting dish until all testes are dissected. 5. Once finished dissecting, discard any ruptured testes (Fig. 2a). 6. Coat 900 Pasteur pipette with 1 Becker Ringer’s solution by pipetting up and down. 7. Remove all the water from imaging dish using a micropipettor. Immediately transfer testes from the dissecting dish to the coverslip portion of the imaging dish using the coated Pasteur pipette (see Note 5). 8. Using the tip of the blunter pair of forceps, carefully press testes onto the coverslip so their apical ends adhere and lie flat against

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the coverslip. There should be no overlap between samples (see Note 6) (Fig. 2a00 ). 9. Remove all of the Becker Ringer’s solution with a Pasteur pipette. Immediately add back enough fresh Becker Ringer’s solution to coat testes, preventing samples from drying out. This will cause testes to strongly adhere to the coverslip allowing them to withstand several media changes if applicable. 10. When ready to image, remove Becker Ringer’s solution and immediately add live imaging solution using a micropipettor (see Note 7). 3.3 Overnight TimeLapse Imaging of the Testis Niche

Imaging instructions are based on using a Zeiss LSM 780 confocal microscope with Zen software. 1. Mount covered imaging dish onto movable stage of the microscope (see Note 8). 2. Use a 60 objective oil lens that will be in contact with the coverslip. For a wider field of view, a 40 objective oil lens can be used. 3. Locate testes using bright field or fluorescent channels (see Note 9). Adjust the laser power, exposure time, and gain for each wavelength used in order to obtain an optimal signal without photobleaching. This will differ depending on the microscope and fluorophore used. The more fluorophores imaged, the greater the chance for photobleaching. 4. Evaluate the fluorescent signal for each testis and select about seven that are favorable for imaging (see Note 10). Save the position of those selected to image at the center plane of the Z range. 5. Once each testis position is selected, set the Z stack to range around the center of the field of interest. Typically a 30 μm range with 1.25 μm steps will suffice. 6. Set the total time for imaging testes and the time between each scan. Typically 25-min intervals over 12.5 h will be enough to follow cellular behaviors (see Note 11). 7. Germline divisions should be seen throughout the entire movie indicating viability of the tissue during the imaging period (Fig. 3a). Nuclear markers can be used to detect cell death since DNA will condense causing bright dense signals. While some spermatogonial death is expected to occur in wild-type testes (Fig. 3a) [10], massive amounts of cell death, especially stem cell and hub cell death which normally does not occur in wild-type testes [11, 12], are indicative of a nonviable tissue (Fig. 3b).

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Fig. 3 Single time point images from overnight movies of a viable (a) and nonviable (b) testis expressing histone-RFP (red) and germline-GFP (green). (a) Viable testes show germline divisions throughout the imaging period (yellow arrow) with some cell death seen in spermatogonia (white arrow). (b) In nonviable testes, cell death is seen in most cells including germline stem cells (yellow arrow) and hub cells (white arrowhead). Dying cells can be detected through the condensation of nuclei as indicated by the dense red signal from the histone-RFP marker 3.4 Immunolocalization After Time-Lapse Imaging

Once live imaging is complete, testes can be fixed and stained directly in the dish for additional markers. 1. Remove all imaging media from the imaging dish using a Pasteur pipette (see Note 12). 2. Quickly add 600 μL of fixative solution onto the coverslip portion of the imaging dish. Incubate at room temperature for 22 min. 3. Remove formaldehyde with Pasteur pipette and discard. Rinse testes twice briefly with ~1 mL PBX. 4. Wash testes three times for 10 min each with ~1 mL PBX and allow the dish to remain stationary at room temperature. Keep the dish covered to minimize evaporation. 5. Remove PBX and add 500 μL of 2 % Normal goat serum in block solution to the imaging dish. Testes should remain covered and stationary for at least 1 h at room temperature or overnight at 4  C to reduce any nonspecific antibody binding (see Note 13). 6. Remove Normal goat serum in block solution and add 500 μL of primary antibody diluted in block solution to an empirically determined working concentration. Incubate dish for 1–2 h at room temperature or overnight at 4  C. Keep dish covered and stationary during incubation. 7. Remove primary antibody with a Pasteur pipette and quickly add ~1 mL PBX (see Note 14).

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8. Rinse twice with ~1 mL PBX then wash three times for 10 min with ~1 mL PBX as in steps 3 and 4. 9. Remove PBX, and add 500 μL of secondary antibody diluted in block solution to an empirically determined working concentration. Keep imaging dish covered and stationary and incubate dish for 1–4 h at room temperature or overnight at 4  C (see Note 15). Place dish inside a foil-covered box to protect samples from light during the secondary antibody incubation and ALL subsequent steps. 10. Remove secondary antibody solution and rinse twice briefly, then twice for 10 min with ~1 mL PBX. 11. Remove PBX then rinse once briefly and once for 10 min with ~1 mL PBS. 12. Remove PBS and immediately cover testes with a drop of Vectashield. 13. Store covered imaging dish with fixed and stained testes on a horizontal surface in a foil-covered box at 20  C until ready to image. 14. Image fixed and stained testes with the same microscope used for live imaging, if possible, to eliminate variation between instruments. 15. Using the map of testes’ locations (or tile scan), locate testes that were previously imaged live. It is helpful to compare the last frame of the movie to fixed testes to ensure they correspond (Fig. 4).

Fig. 4 Comparison of a live (a) and fixed (b) testis after 12.5 h of imaging. Staining after imaging allows for the use of cell specific markers. Cells tracked live that are histone-RFP positive (red), germline-GFP negative (nongreen), and close to the hub can be accurately identified as CySCs through ZFH1 staining (white) (compare a and b yellow arrow). Hub cells (white asterisk) are encircled by the germline stem cells, but can also be identified by Fasciclin 3 staining (b, blue)

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Notes 1. Not all fluorescent reporters or fluorescently tagged proteins are strong enough for live imaging. Those that work well in adult testes include but are not limited to: nos::Moe-GFP to visualize the germline (from R. Lehmann) [13], His2Av-RFP to visualize nuclei of all cells (Bloomington Stock Center), G-TRACE reporter for lineage tracing (Bloomington Stock Center) [14], and FUCCI reporter for cell cycle analysis (Bloomington Stock Center) [15]. Flies should be raised in incubators at constant temperature (18 or 25  C) and humidity (65 %), and shifted to the appropriate temperatures (29 or 31  C) prior to dissection if applicable to the experiment. 2. To make lined dissecting dishes, add 9 mL of elastomer base and 1 mL curing agent from a Sylgard 184 Silicone Elastomer Kit (Dow Corning) to a disposable 15 mL conical tube. Add a pinch of 100–400 mesh activated charcoal powder (Sigma Aldrich) and invert gently to mix. Continue to add charcoal until the elastomer is opaque, and then pour solution into a clean beveled edge watch glass. Remove bubbles from elastomer surface with a Pasteur pipette. Place dish in a covered container and cure overnight at room temperature. 3. Dyes such as MitoTracker Red CMXRos (1 μM) (Molecular Probes) can be added to the imaging media as well as various drugs. Test toxicity of new dyes or drugs to the tissue over a 12.5-h period prior to use in live imaging. 4. To separate the testis from the cuticle use both prongs of the forceps to pull the testis off the accessory gland or place one prong of the forceps inside the spiral of the testis (careful to not puncture the testis) and pull with the anchor hand. Testes should also be separated from each other. 5. If there is excess debris in the media following dissection, testes can be gently transferred to a separate dissecting dish containing fresh Becker Ringer’s solution, rinsed a few times with fresh Becker Ringer’s solution, and then transferred to the imaging dish. Try to minimize handling of testes and number of transfers as this could damage the tissue. 6. Testes will loosely adhere to the coverslip portion of the imaging dish due to the Poly-L-Lysine. If repositioning of testes is necessary, be extremely gentle since too much force can damage the tissue. 7. Media will spill over the coverslip portion of the dish. Try to keep as much media over the testes as possible. Too little media will lead to evaporation causing the testes to dry out during imaging. If desired, incubate testes in media containing dye or drug prior to imaging.

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8. Using a microscope fitted with an environmental chamber is desirable to further stabilize the dish on the stage. Utilizing the heat or humidity controls is optional. 9. Using paper, map the location of each testis on the imaging dish prior to mounting the dish on the microscope. Alternatively, create a tiled image using the 10 objective to get an overview of where each testis is located. This helps to ensure that each testis is evaluated before choosing which to image. 10. Select testes in which the fluorescent signal is strong, indicating the field of interest is closest to the coverslip, and there is little movement due to muscle contractions in order to prevent blurred images. If using multiple fluorophores, ensure all have a strong signal. 11. The interval between scans should allow for enough time for each testis to be imaged before the next cycle begins. On a LSM 780 microscope, 7–8 testes imaged with GFP and RFP with 30 μm stacks take approximately 15 min to image. This allows for 10 min between time points that the testes are not being exposed to the laser thus helping to maintain tissue viability and minimize photobleaching. 12. It is easiest to remove solutions from the dish using a long 900 Pasteur pipette and to add in fresh solutions when washing testes using a shorter 5.7500 Pasteur pipette. Position pipette so that it isn’t directly over any testes since too much suction can cause testes to dissociate from the coverslip. 13. If evaporation of the solution during overnight incubations is a concern, covered dishes can be kept in a sealed container containing a wet Kimwipe. 14. Many primary antibodies can be saved, stored at 4  C and reused. 15. A nuclear counterstain such as 40 -6-diamidino-2-phenylindole (DAPI) can be added to the secondary antibody solution at a final concentration of 1 μg/mL.

Acknowledgements The authors thank Steve DiNardo and Kari Lenhart for sharing adaptations to the protocol, and Margaret de Cuevas and Miriam Akeju for comments on the manuscript. This work was supported by NIH Grants HD040307 and HD052937 (EM). The Zeiss LSM 780 confocal microscope used in this study was funded by NIH Grant S10 OD016374. LJG was supported by T32 GM007445.

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References 1. Sheng XR, Matunis E (2011) Live imaging of the Drosophila spermatogonial stem cell niche reveals novel mechanisms regulating germline stem cell output. Development 138:3367–3376 2. Hara K, Nakagawa T, Enomoto H, Suzuki M, Yamamoto M, Simons BD, Yoshida S (2014) Mouse spermatogenic stem cells continually interconvert between equipotent singly isolated and syncytial states. Cell Stem Cell 14:658–672 3. de Cuevas M, Matunis EL (2011) The stem cell niche: lessons from the Drosophila testis. Development 138:2861–2869 4. Prasad M, Jang AC, Starz-Gaiano M, Melani M, Montell DJ (2007) A protocol for culturing Drosophila melanogaster stage 9 egg chambers for live imaging. Nat Protoc 2:2467–2473 5. Prasad M, Montell DJ (2007) Cellular and molecular mechanisms of border cell migration analyzed using time-lapse live-cell imaging. Dev Cell 12:997–1005 6. Morris LX, Spradling AC (2011) Long-term live imaging provides new insight into stem cell regulation and germline-soma coordination in the Drosophila ovary. Development 138:2207–2215 7. Cheng J, Hunt AJ (2009) Time-lapse live imaging of stem cells in Drosophila testis. Curr Protoc Stem Cell Biol 2: Unit 2E.2 8. Lenhart KF, DiNardo S (2015) Somatic cell encystment promotes abscission in germline

stem cells following a regulated block in cytokinesis. Dev Cell 34:192–205 9. Ashburner M (1989) Drosophila: a laboratory manual. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY, p 376 10. Yacobi-Sharon K, Namdar Y, Arama E (2013) Alternative germ cell death pathway in Drosophila involves HtrA2/Omi, lysosomes, and a caspase-9 counterpart. Dev Cell 25:29–42 11. Brawley C, Matunis E (2004) Regeneration of male germline stem cells by spermatogonial dedifferentiation in vivo. Science 304:1331–1334 12. Hasan S, He´tie´ P, Matunis EL (2015) Niche signaling promotes stem cell survival in the Drosophila testis via the JAK-STAT target DIAP1. Dev Biol 404:27–39 13. Sano H, Renault AD, Lehmann R (2005) Control of lateral migration and germ cell elimination by the Drosophila melanogaster lipid phosphate phosphatases Wunen and Wunen 2. J Cell Biol 171:675–683 14. Evans CJ, Olson JM, Ngo KT, Kim E, Lee NE, Kuoy E, Patananan AN, Sitz D, Tran P, Do MT, Yackle K, Cespedes A, Hartenstein V, Call GB, Banerjee U (2009) G-TRACE: rapid Gal4-based cell lineage analysis in Drosophila. Nat Methods 6:603–605 15. Zielke N, Korzelius J, van Straaten M, Bender K, Schuhknecht GF, Dutta D, Xiang J, Edgar BA (2014) Fly-FUCCI: a versatile tool for studying cell proliferation in complex tissues. Cell Rep 7:588–598

Chapter 5 RNA Isolation from Early Drosophila Larval Ovaries Dana Gancz and Lilach Gilboa Abstract In the fruit fly Drosophila melanogaster, ovarian germline stem cells (GSCs) and their niches form during larval development. This process is poorly studied partly due to technical difficulties in isolating early larval ovaries. In addition, purifying RNA from larval ovaries proves to be more challenging than purifying it from other organs. Here we describe a technique for dissecting ovaries from early larvae and advise on how to extract RNA with maximum yield and purity. RNA isolation allows assaying gene expression in a direct and quantitative manner, which is invaluable for understanding molecular events underlying ovarian niche formation and GSC establishment. Key words RNA isolation, TRIzol/TRI reagent, Drosophila, Larvae, Ovary

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Introduction Stem cells and their niches develop from precursor cells during gonadogenesis. Understanding how they form is imperative for our understanding of tissue homeostasis, disease, and regeneration. Since normal development of stem cells and niches depends on interactions with neighboring cells as a part of organ development, analyzing this complex process must be performed in vivo. Ovarian germline stem cells (GSCs) in the fruit fly Drosophila melanogaster have been studied extensively as a model of how adult stem cells interact with their somatic niche cells. However, only a few studies have been dedicated to understanding how GSCs and niche cells form from their precursor cells: primordial germ cells (PGCs) and somatic lineages, respectively [1–10]. The somatic niches begin to form within the larval ovary 2–3 days after egg laying. The study of larval ovary development is hampered by the fact that these organs are small (a diameter of ~50 μm when niches start forming), transparent, and embedded in a white fat body, making their isolation physically challenging.

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Despite their small size, RNA purification coupled with quantitative methods, such as RNA sequencing (RNAseq) or quantitative PCR (qPCR), can be used to assay gonadal gene expression in a direct and quantitative manner. This can open a window into the molecular events underlying ovarian niche formation and GSC establishment. Successful isolation of intact RNA requires the immediate inactivation of endogenous RNases by strong denaturants, such as guanidinium isothiocyanate or guanidinium hydrochloride, that simultaneously disrupts cells, solubilizes their components, and denatures proteins [11]. Commercial kits based on guanidinium isothiocyanate and designed for isolating RNA from a small number of cells are available and work well with larval organs such as salivary glands, imaginal discs, and brain. However, in our experience, using TRIzol (Invitrogen) gives the highest yield and quality when purifying RNA from larval ovaries. TRIzol or the equivalent TRI reagent (Sigma-Aldrich) is a monophasic solution of phenol and guanidinium isothiocyanate [12], which are based on the single-step RNA isolation method developed by Chomczynski and Sacchi [13]. After sample homogenization in TRIzol/TRI reagent, the addition of chloroform followed by centrifugation separates the solution into an aqueous phase and an organic phase. RNA remains exclusively in the aqueous phase and can be recovered by precipitation with isopropyl alcohol. This protocol, which can be easily scaled up or down, allows rapid purification of high-yield RNA from multiple samples, retrieving high-abundance as well as low-abundance RNA isoforms. The purified RNA is of a wide range of sizes, including small RNAs, such as microRNAs, piRNAs, or small interfering RNAs [6, 14, 15]. However, some bias might occur when assaying a very small number of cells [16]. Here we describe a technique for isolating ovaries from midthird instar larvae (96 h after egg laying, when adult niche markers are first expressed) followed by RNA purification with TRIzol/TRI reagent. We offer technical solutions for locating the ovaries and separating them from the fat body and indicate variations to the manufacturer’s protocol designed to extract RNA with maximum yield and purity specifically from this tissue. The amount of ovaries to be isolated differs according to developmental timing and the end use. For simple qPCR of mid-third instar larval ovaries, 20 ovaries should suffice while an RNA expression array will require about 150–200 ovaries.

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Materials Diligently follow all waste disposal regulations when disposing of waste materials.

2.1

Ovary Dissection

1. Ringer’s buffer: 128 mM NaCl, 2 mM KCl, 1.8 mM CaCl2, 4 mM MgCl2, 35.5 mM sucrose, 5 mM Hepes pH 6.9. 2. Dumont biology tweezers 5 dumstar polished (Fine Science Tools). 3. A plastic plate. 4. Dissecting dish: 9-Well plates 85  100 mm, 22 mm O.D.  7 mm deep. 5. Shallow dissecting dish: Microscope slides with cavities 76  26 mm, 15 mm O.D.  0.8 mm deep. 6. Stereo Microscope MZ 16.5 with a standard transmitted light base TL ST (Leica Microsystems). 7. Nickel plated pin holder and stainless steel minutien pins 0.1 mm diameter, 10 mm long. 8. Liquid nitrogen.

2.2 RNA Purification Reagents

1. TRIzol (Life technologies) or TRI reagent (Sigma). 2. 1 ml Syringe with a 21 G 1½ in., Nr. 2, (0.8  40 mm) needle. 3. H2O, RNase-free or DEPC-treated H2O. 4. Chloroform. 5. Ethanol: 70 % in RNase-free H2O. 6. Glycogen, RNA grade. 7. Isopropanol.

3

Methods Wear gloves at all times and follow good laboratory and safety practices when handling the tissue to avoid RNase contamination.

3.1 Dissection of Larvae

1. Flood a vial (see Note 1) with tap water and gently scratch the surface of the food with tweezers (forceps) until the larvae float, and then pour the water into a plastic plate. 2. Pick larvae from the water (see Note 2) using fine biological tweezers and place them in a dissecting dish containing Ringer’s buffer at room temperature (RT) (see Notes 3 and 4). 3. Under a microscope separate female larvae from males. The gender can be distinguished by looking at the fat body approximately one-third from the posterior end. Male testes appear as

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Fig. 1 An illustration of larval ovary dissection technique. (a) Larval head should be removed posterior to the brain. (b) The fat body is extracted by holding the larva by the posterior spiracles (with the trachea facing down) with one pair of forceps and slowly squeezing the fat body out with the other pair, leaving behind the cuticle and trachea. (c) The ovaries can be found approximately one-third from the posterior, surrounded by a “flower” of fat body (arrows). (d) Removing the fat body surrounding the gonad can be achieved by crossing two pins between fat body and gonad all around the gonad

big clear ovals, while female ovaries, located at the same place, are much smaller, clear, round spheres. 4. Transfer females into a shallow dissecting well containing Ringer’s buffer and remove the head by holding the larva posterior to the brain with one pair of forceps and pulling the head with a second pair of forceps. 5. Hold the remaining part of the larva by the posterior spiracles (with the trachea facing down) with one pair of forceps and slowly squeeze the fat body out with the other pair, leaving behind the cuticle and trachea (Fig. 1). Discard the cuticle (see Note 5). 6. Use two nickel-plated pin holders, holding 0.1 mm diameter pins, to isolate the gonads, which are surrounded by a “flower”-shaped fat body (Fig. 1d). 7. Sever the connections between the fat body and the gonad by crossing the pins between the two tissues. Repeat this in all places where the fat body is attached to the gonad until no fat body remains. Separate the isolated gonad away from the fat body residue (Fig. 1d, see Notes 6 and 7). 8. Aspirate the ovaries with a minimal volume of Ringer’s buffer (see Note 8) using a pipet with a 1–10 μl filter tip. 9. Place the ovaries in an RNase-free 1.7 ml Eppendorf tube and snap freeze in liquid nitrogen (see Notes 9 and 10).

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RNA Purification

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This variation of the manufacturer’s protocol is best suited for extracting RNA from larval ovaries. 1. Cool a centrifuge to 4  C and heat RNase-free H2O to 60  C. 2. Establish an RNase-free working environment (see Note 11). 3. Take the ovaries out of the liquid nitrogen and immediately add 500 μl TRIzol/Tri Reagent (see Notes 12–14). 4. Homogenize the sample by passing ten times through a 1 ml syringe with a 21 G needle. 5. Add 20 μg RNA-grade glycogen that will serve as a carrier (see Note 15) and vortex the sample gently for 15 s. 6. Incubate for 5 min at RT (see Note 16). 7. Add 100 μl chloroform and vortex vigorously for 15 s (see Note 17). 8. Incubate for 15 min at room temperature. 9. Centrifuge the samples at 12,000  g for 15 min at 4  C (see Note 18). 10. Collect the aqueous phase (see Note 19) and transfer to a new tube. 11. Add 250 μl isopropanol and vortex the sample vigorously for 15 s. 12. Incubate at room temperature for 10 min (see Note 20). 13. Centrifuge the samples at 12,000  g for 15 min at 4  C. 14. Carefully remove the supernatant without touching the RNA pellet (see Note 21). 15. Wash the pellet with 500 μl cold ( 20  C) 70 % ethanol (see Note 22). 16. Centrifuge the samples at 12,000  g for 10 min at 4  C. 17. Carefully remove the supernatant without touching the loose RNA pellet. 18. Centrifuge briefly (spin down) and completely remove the supernatant. 19. Air-dry pellet for 3–7 min (see Note 23). 20. Dissolve the pellet in 10–13 μl warm (60  C) RNase-free H2O, by gently passing the solution a few times through a pipette tip, and then immediately place on ice (see Notes 24–26).

3.3 Assessing RNA Yield and Quality

1. Measure RNA concentration and purity using a NanoDrop 1000A Spectrophotometer. RNA of good quality should have a A260/A280 ratio of ~2 and A260/A230 ratio of >1.5 (see Notes 27 and 28).

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Fig. 2 An output of a Bioanalyzer analysis. RNA was extracted from 150 early larval ovaries and examined using an Agilent 2100 Bioanalyzer. The RNA profile shows the 18S (1995 nt), 5.8S (123 nt), and 28S (3945 nt) rRNA. Due to a “hidden break” Drosophila 28S rRNA is processed into two fragments that migrate in a similar manner to the 18S rRNA [18, 19]. FU fluorescence intensity, nt nucleotides

2. Measure RNA integrity using a Bioanalyzer. The RNA profile should show the 28S, 18S, and 5S ribosomal RNA (Fig. 2, see Note 29).

4

Notes 1. For correct timing of larvae, it is important to keep the cultures under-crowded (about 30 eggs/25 mm vial), at 25  C with 60–80 % humidity. Flies are allowed to lay eggs for 2–4 h and then 96 h after egg laying the larvae reach the non-wandering third instar stage (mid-L3). 2. Avoid leaving the larvae in the water for more than a few minutes, as this stressful, hypoxic condition may alter gene expression. 3. Ringer’s solution is favorable for in vitro experiments on organs or tissues; however, PBS can also be used. 4. To prevent gene expression changes prior to freezing the ovaries, buffers and ovaries should be kept at room temperature. 5. Try to keep the fat body intact and attached to the gut as this will help in recognizing the location of the gonads. 6. Care should be taken when manipulating the pins not to poke or stretch the ovary, as this will lead to cell stress. 7. The dissected ovaries tend to stick to the glass well. Coating the well with repulsive covers, such as SIGMACOTE, can reduce adhesiveness. 8. The total sample volume should not exceed 1/10 of the TRIzol/TRI reagent volume. 9. To prevent RNA degradation, the entire dissection procedure (from larval head removal to placing the dissected ovaries in liquid nitrogen) should take no longer than 20 min.

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10. At this stage the ovaries can be stored at 80  C; however, in our experience, RNA quality and yield are better when proceeding immediately to RNA purification. 11. To ensure successful purification of intact RNA it is important to maintain an RNase-free environment. Clean work surfaces, pipettes, and all other equipment with an RNase decontamination solution, such as RNaseZAP (Life Technologies). Pipettes and automatic pipettes should be reserved for RNA work to prevent cross-contamination with RNases from shared equipment. Use RNase-free disposable plastic-ware or nondisposable glassware that was decontaminated. Use commercial RNase-free H2O or DEPC-treated H2O. Wear disposable gloves at all times and use sterile technique when handling the reagents used for RNA isolation or analysis. 12. TRIzol/TRI reagent contains phenol (toxic and corrosive) and guanidine isothiocyanate (an irritant); therefore subsequent steps should be performed in a fume hood wearing a lab coat and gloves. 13. To prevent RNA degradation, samples should not be allowed to thaw before the addition of the TRIzol reagent. 14. If the sample is divided into several tubes, they should be combined at this stage. 15. The glycogen remains in the aqueous phase and is coprecipitated with the RNA. The glycogen concentration should not exceed 4 mg/ml. 16. This incubation allows the complete dissociation of nucleoprotein complexes. 17. Chloroform addition results in phase separation. It is important that the chloroform does not contain any additives such as isoamyl alcohol. 18. The mixture separates into three phases: a lower red phenol/ chloroform phase, an interphase that contains DNA and proteins, and an upper clear phase that contains RNA. 19. Care should be taken not to disturb the lower phases as this will result in phenol contamination. The aqueous phase should be about 50 % of the total volume; however, it is recommended to leave behind 20 % of it. 20. The sample can also be incubated overnight (or at least a few hours) at 20  C. In our hands, longer incubation times increase the yield but reduce the quality of the obtained RNA and is therefore not recommended. 21. After centrifugation the RNA should look like a white pellet at the bottom and side of the tube. 22. Do not vortex samples to avoid losing the small and loose pellet.

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23. Be careful not to completely dry the pellet as this will greatly decrease its solubility. 24. Heating the water to 60  C facilitates dissolving of the RNA. 25. For use in polymerase chain reaction (PCR), it is recommended to treat the RNA with DNaseI when the two primers lie within a single exon. However, this might significantly reduce RNA yield. 26. At this stage the RNA can be stored in 80  C. However, if possible, we recommend continuing immediately with downstream applications such as cDNA synthesis. 27. Contamination with proteins, phenol, or glycogen used for precipitation will result in a lower A260/A280, hampering the accurate quantification of RNA. However, 20 ovaries should yield enough RNA for qPCR and 150 ovaries can be used for a microarray. 28. A regular spectrophotometer can also be used. An aliquot of the RNA should be dissolved in 1 mM Na2HPO4, with a pH above 7.5 The reading at 260 nm allows calculation of the concentration of RNA in the sample. An optical density of 1 corresponds to approximately 40 g/ml of single-stranded RNA. 29. The gene regions of Drosophila rDNA encode four individual rRNAs: 18S (1995 nt), 5.8S (123 nt), 2S (30 nt), and 28S (3945 nt) [17]; however due to a “hidden break” Drosophila 28S rRNA is processed into two fragments that migrate in a similar manner to the 18S rRNA [18, 19]. References 1. Gancz D, Gilboa L (2013) Insulin and Target of rapamycin signaling orchestrate the development of ovarian niche-stem cell units in Drosophila. Development 140(20):4145–4154 2. Gancz D, Lengil T, Gilboa L (2011) Coordinated regulation of niche and stem cell precursors by hormonal signaling. PLoS Biol 9(11): e1001202 3. Gilboa L, Lehmann R (2004) How different is Venus from Mars? The genetics of germ-line stem cells in Drosophila females and males. Development 131(20):4895–4905 4. Gilboa L, Lehmann R (2006) Soma-germline interactions coordinate homeostasis and growth in the Drosophila gonad. Nature 443 (7107):97–100 5. Lengil T, Gancz D, Gilboa L (2015) Activin signaling balances proliferation and differentiation of ovarian niche precursors and enables adjustment of niche numbers. Development 142(5):883–892

6. Dvorak Z, Pascussi JM, Modriansky M (2003) Approaches to messenger RNA detection comparison of methods. Biomed Pap Med Fac Univ Palacky Olomouc Czech Repub 147 (2):131–135 7. Sarikaya DP, Extavour CG (2015) The Hippo pathway regulates homeostatic growth of stem cell niche precursors in the Drosophila ovary. PLoS Genet 11(2):e1004962 8. Song X, Call GB, Kirilly D, Xie T (2007) Notch signaling controls germline stem cell niche formation in the Drosophila ovary. Development 134(6):1071–1080 9. Zhu CH, Xie T (2003) Clonal expansion of ovarian germline stem cells during niche formation in Drosophila. Development 130 (12):2579–2588 10. Hodin J, Riddiford LM (1998) The ecdysone receptor and ultraspiracle regulate the timing and progression of ovarian morphogenesis

RNA Isolation from Early Larval Ovaries during Drosophila metamorphosis. Dev Genes Evol 208(6):304–317 11. Chirgwin JM, Przybyla AE, MacDonald RJ, Rutter WJ (1979) Isolation of biologically active ribonucleic acid from sources enriched in ribonuclease. Biochemistry 18(24):5294–5299 12. Rio DC, Ares M Jr, Hannon GJ, Nilsen TW (2010) Purification of RNA using TRIzol (TRI reagent). Cold Spring Harb Protoc 2010(6): pdb prot5439 13. Chomczynski P, Sacchi N (1987) Single-step method of RNA isolation by acid guanidinium thiocyanate-phenol-chloroform extraction. Anal Biochem 162(1):156–159 14. Sarkies P, Selkirk ME, Jones JT, Blok V, Boothby T, Goldstein B, Hanelt B, Ardila-Garcia A, Fast NM, Schiffer PM, Kraus C, Taylor MJ, Koutsovoulos G, Blaxter ML, Miska EA (2015) Ancient and novel small RNA pathways compensate for the loss of piRNAs in multiple independent nematode lineages. PLoS Biol 13 (2):e1002061 15. Ahmann GJ, Chng WJ, Henderson KJ, PriceTroska TL, DeGoey RW, Timm MM, Dispenzieri A, Greipp PR, Sable-Hunt A, Bergsagel L,

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Fonseca R (2008) Effect of tissue shipping on plasma cell isolation, viability, and RNA integrity in the context of a centralized good laboratory practice-certified tissue banking facility. Cancer Epidemiol Biomarkers Prev 17 (3):666–673 16. Kim Y-K, Yeo J, Ha M, Kim B, Kim VN (2012) Retraction notice to: cell adhesion-dependent control of microRNA decay. Molecular Cell 43, 1005-1014; September 16, 2011 (2012). Mol Cell 46(6):896 17. Tautz D, Hancock JM, Webb DA, Tautz C, Dover GA (1988) Complete sequences of the rRNA genes of Drosophila melanogaster. Mol Biol Evol 5(4):366–376 18. Melen GJ, Pesce CG, Rossi MS, Kornblihtt AR (1999) Novel processing in a mammalian nuclear 28S pre-rRNA: tissue-specific elimination of an ‘intron’ bearing a hidden break site. EMBO J 18(11):3107–3118 19. Macharia RW, Ombura FL, Aroko EO (2015) Insects’ RNA profiling reveals absence of “Hidden Break” in 28S ribosomal RNA molecule of onion thrips, thrips tabaci. J Nucleic Acids 2015:965294

Chapter 6 Live-Cell Imaging of the Adult Drosophila Ovary Using Confocal Microscopy Nevine A. Shalaby and Michael Buszczak Abstract The Drosophila ovary represents a key in vivo model used to study germline stem cell (GSC) maintenance and stem cell daughter differentiation because these cells and their somatic cell neighbors can be identified at single-cell resolution within their native environment. Here we describe a fluorescent-based technique for the acquisition of 4D datasets of the Drosophila ovariole for periods that can exceed 12 consecutive hours. Live-cell imaging facilitates the investigation of molecular and cellular dynamics that were not previously possible using still images. Key words Drosophila ovary, Live-cell imaging, Confocal microscopy, Molecular dynamics, Cellular dynamics

1

Introduction Studies in the Drosophila ovary have provided important insights into stem cell biology. Each adult female has a pair of ovaries with ~16 ovarioles. At the anterior tip of the ovariole is a structure called the germarium where 2–3 germline stem cells (GSC) reside adjacent to 6–8 somatic cap cells that comprise the GSC niche (Fig. 1). When a GSC divides it gives rise to two daughter cells. Daughter cells positioned adjacent to the cap cells remain stem cells, while daughters displaced away from the niche undergo four incomplete mitotic divisions to give rise to a 16-cell cyst. One of the 16 cells becomes the oocyte, while the other 15 cells develop into nurse cells. Somatic follicle cells envelop this 16-cell cyst to form an egg chamber, which continues to grow and develop until it is released as a mature egg [1]. The key to the success of the Drosophila ovary as a model for stem cell biology is the development of molecular genetic tools that enable the identification and functional characterization of genes required during GSC maintenance and daughter differentiation [2, 3]. These studies have led to the establishment

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Fig. 1 (a) Dissect the ovary out of the female abdomen and separate out individual ovarioles. (b) Using very fine forceps, carefully remove the muscle sheath from individual ovarioles

of well-defined molecular markers that allow us to identify different cell types within the ovariole, at single-cell resolution. Excitingly, more recent advances in the use of live-cell imaging now allow one to study the molecular and cellular dynamics of female GSCs and their neighbors. In addition, live-cell imaging revealed the asymmetric inheritance of RNA particles during GSC divisions [4], as well as the asymmetric recovery of the Pol I regulator, Udd, within the nucleoli of GSCs and their daughters [5]. Further characterization of the dynamic expression and distribution of specific proteins and different aspects of cell-cell interactions will provide a much richer understanding of germline stem cell biology. This chapter describes a simple technique for the imaging of ovarioles within the Drosophila ovary for at least 12 consecutive hours, using available transgenic lines that contain fluorescently tagged proteins. This protocol is written for proficient users of a confocal microscope.

2

Materials

2.1 DrosophilaRearing Components

1. Drosophila lines with a constitutively active fluorescent tag in the cells of interest. For example, nanosGal4, UAS-RFP, or Vasa::GFP. Alternatively lines can be generated that express your specific protein of interest tagged with a fluorescent marker (not covered in this chapter).

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2. Fly-rearing bottles with freshly made fly food. The standard cornmeal, Bloomington fly food recipe, is sufficient (http:// flystocks.bio.indiana.edu/Fly_Work/media-recipes/ bloomfood.htm). 3. 18  C Incubator. 4. Wet yeast: Mix dry yeast with enough water to make a paste. 2.2 Drosophila Ovary-Dissecting Components

1. Dissecting dish with at least 2 wells, deep enough to cover the entire ovary.

2.3 Media Components

1. Pieces of 4 % low-melting agarose: Prepare 4 % low-melting agarose in Nano pure water. For example, measure 2 g of lowmelting agarose in 50 ml of Nano pure water, and briefly bring to a boil in a microwave. Polymerize in a sterile container (e.g., a petri dish), cut into 1 cm2 pieces, and place pieces in a 1 L bottle with Nano pure water to dialyze. Change water every 12 h for at least 48 h. Transfer pieces to a 50 ml conical with Nano pure water. Store at 4  C.

2. Two pairs of dissecting forceps; one pair does not need to have sharp tips because these will be used to dissect the ovary out of the abdomen (e.g., Dumont # 5). While the second pair must have very sharp tips because these will be used to separate the ovarioles from each other and carefully remove the muscle sheath (e.g., Dumont # 55).

2. Prepare media: 1:10 Fetal bovine serum (FBS), 200 μg/ml human insulin, 1:100 penicillin/streptomycin (stock: 10,000 IU/ml penicillin, 10 mg/ml streptomycin), in Schneider’s Drosophila medium. Aliquot in 200 μl aliquots and store at 4  C. 3. Two heat blocks, one set to 45  C and the other set to 32  C. 2.4 Mounting and Microscopy

1. Microscope slides. 2. Invisible tape. 3. Tungsten needles. 4. 22  22 mm Cover slips. 5. Sealant glue. 6. Confocal laser scanning microscope with a resonant scanner (e.g., Leica TCS SP8) or with a spinning disc (e.g., Andor spinning disk confocal), equipped with laser lines and filters for the fluorescent proteins in use.

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

3.1

Preparing Flies

1. For best results, flies should be raised at 18  C in nonovercrowded bottles (see Note 1). To do this, place 20 males and 20 females, of the desired genotype, in a fresh bottle with a small spoon of wet yeast on the side of the bottle. Allow them to mate and lay eggs for 48 h at 25  C. Then, remove the adult flies and place the bottle with eggs and larvae at 18  C until they eclose. The idea is to prevent overcrowding of the bottles, which leads to smaller larvae and adult flies. You may need to adjust the number of parents added, depending on the fertility of the genotype you are using (see Note 2). 2. Approximately 20 days later, take 5–10 adult day-0 females and place them in a fresh vial with wet yeast and place the vial at 18  C for 48 h to fatten the ovaries.

3.2 Preparing Ovaries

1. Mix media with agarose: Warm two 200 μl tubes of media prepared above in the heat block set to 45  C. Melt a small piece of agarose in the microwave using the medium setting on the microwave. Add 25 μl of the molten agarose to 200 μl of warmed media. (You may find it easier to pipette the agarose by cutting off the end of the tip to create a larger hole.) Mix well and place in the heat block set to 32  C (see Note 3). 2. Dissect ovaries: Add 200 μl of media to a dissecting well. Wait 30 s for it to cool before dissecting the ovaries. Using the less sharp forceps, dissect out one ovary pair from a single female (Fig. 1). Remove the rest of the fly keeping only the pair of ovaries in the well. Using very fine forceps, carefully separate out several ovarioles from the ovary (Fig. 1). Then to each individual ovariole, remove the muscle sheath by pinching it from the more developed egg chambers and slowly pulling it off towards the germarium (Fig. 1). When the muscle sheath has been completely removed the ovariole looks like a “string of beads.” Do not touch the ovariole directly with your forceps. Repeat this for ~10 ovarioles. Note: Do not use the sharp forceps for the initial dissection of the ovary out of the fly, as the cuticle on the fly will very quickly lead to blunt forceps.

3.3 Mounting Ovarioles

1. Add 200 μl of media + agarose (warmed at 32  C) to a well. Let cool for a few seconds, and then transfer separated ovarioles, using a pipette tip, to the well. Transfer 5–10 ovarioles. 2. Transfer to a slide: Add invisible tape to a slide to bridge the cover slip. The purpose of the tape is to create a gap between

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the cover slip and slide, preventing the specimen from getting “squashed.” Using a pipette tip, transfer ovarioles with media + agarose to the slide. Separate the ovarioles out with tungsten needles. Bring them closer to the surface of the media. Place a 22  22 mm cover slip carefully on top, and seal the slide with glue. 3.4

Imaging

1. Place the prepared slide on the microscope stage. 2. Use bright-field illumination to locate an ovariole. Avoid using epifluorescence illumination to search for ovarioles as this may lead to photobleaching of the fluorescence in the ovariole. 3. Visually inspect the ovariole to ensure that it has not been damaged during dissection, and select a different one if necessary. 4. Center the ovariole and switch to the desired objective. 5. Similar to still images, before starting the laser scanner, set the laser power down, adjust the pinhole to Airy units 1–2 (depending on the desired resolution and strength of the epifluorescence), and adjust the zoom to 1.0. 6. Turn on the laser scanners and adjust the focal plane to visualize the cells. Adjust the zoom (for observing GSCs at the tip of the germarium, use 63 objective and zoom in as necessary), laser power (keeping it as low as possible, and the gain as high as possible, to avoid photobleaching and damaging the ovariole), and pinhole. Observe the cells for a minute at a “fast scan” mode to check for healthy movements of the cells. This may take several trials before you can recognize healthy cells versus unhealthy ones. 7. After all the necessary microscope parameters have been determined (as you would adjust for still images), start the 4D adjustments. Adjust the Z-stack parameters and then determine how often the microscope should take a Z-stack. This technique works for a “continuous” scan, as well as a scan that is taken less often, such as every 30 min or 1 h (see Note 4). 8. Figure 2 is an example of the use of live-cell imaging to track the rate of translation across the germarium. In this experiment, mEos was used as a tool to study translation by observing the recovery rate of GFP protein following photoconversion with UV light [6]. We generated a fly carrying nanos-Gal4, UASp-GAP43.mEos. Both transgenic lines were acquired separately from the Bloomington Drosophila Stock Center and the single fly carrying both lines was generated in the laboratory. The resultant fly expresses GAP43::mEos in nanos-Gal4-expressing cells. The germarium was exposed to 5 s of UV light which was sufficient to convert GFP to RFP. Z-stacks were taken every 30 min for 12 consecutive hours, to monitor the recovery rate of GAP43::mEos in nanos-Gal4-expressing cells.

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Fig. 2 Timeline of a photoconversion experiment in the Drosophila adult ovary. A germarium of the genotype nanos-Gal4, UASp-Gap43::mEos was exposed to 5 s of UV light for photoconversion of GFP to RFP. Z-stacks were taken every 30 min for 12 consecutive hours to monitor the recovery of translation across the germarium. In this particular case, expression of unconverted Gap42::mEos appeared to occur quicker in the GSC compared to its daughter cells (GSC germline stem cells)

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Notes 1. Ovaries from female flies raised at 18  C seemed to survive the dissection and long imaging sessions better than older flies or flies raised at 25  C. 2. Several genotypes may interfere with cell survival during long imaging sessions. Raising them at 18  C may help to alleviate this issue, as well as decrease the laser power during the imaging session. 3. Be sure to use low-melting agarose. High-melting agarose will damage your sample. 4. It may take several imaging sessions to be able to differentiate between healthy-looking cells versus unhealthy ones. Therefore, be sure to monitor the session frequently for the first few sessions before leaving it overnight or unmonitored for long time periods.

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Acknowledgement We would like to thank P. Robin Hiesinger and Neset Ozel for their input and advice while developing this protocol. References 1. Eliazer S, Buszczak M (2011) Finding a niche: studies from the Drosophila ovary. Stem Cell Res Ther 2:45 2. Gilboa L (2015) Organizing stem cell units in the Drosophila ovary. Curr Opin Genet Dev 32:31–36 3. Slaidina M, Lehmann R (2014) Translational control in germline stem cell development. J Cell Biol 207:13–21 4. Fichelson P, Moch C, Ivanovitch K et al (2009) Live-imaging of single stem cells within their niche reveals that a U3snoRNP component

segregates asymmetrically and is required for self-renewal in Drosophila. Nat Cell Biol 11:685–693 5. Zhang Q, Shalaby NA, Buszczak M (2014) Changes in rRNA transcription influence proliferation and cell fate within a stem cell lineage. Science 343:298–301 6. Nienhaus GU, Nienhaus K, Holzle A et al (2006) Photoconvertible fluorescent protein EosFP: biophysical properties and cell biology applications. Photochem Photobiol 82:351–358

Chapter 7 Measurement of mRNA Poly(A) Tail Lengths in Drosophila Female Germ Cells and Germ-Line Stem Cells Aymeric Chartier, Willy Joly, and Martine Simonelig Abstract mRNA regulation by poly(A) tail length variations plays an important role in many developmental processes. Recent advances have shown that, in particular, deadenylation (the shortening of mRNA poly (A) tails) is essential for germ-line stem cell biology in the Drosophila ovary. Therefore, a rapid and accurate method to analyze poly(A) tail lengths of specific mRNAs in this tissue is valuable. Several methods of poly (A) test (PAT) assays have been reported to measure mRNA poly(A) tail lengths in vivo. Here, we describe two of these methods (PAT and ePAT) that we have adapted for Drosophila ovarian germ cells and germ-line stem cells. Key words Deadenylation, Drosophila, Germ line, Germ-line stem cell, Ovary, PAT assay, Poly(A) tail, Polyadenylation

1

Introduction The regulation of developmental processes and cellular activities largely relies on mRNA regulation, including control of mRNA localization, stability, and translation. An important mechanism of this regulation, which can affect these three levels of control, involves changes in mRNA poly(A) tail lengths by deadenylation or cytoplasmic polyadenylation [1]. These modes of control are particularly important in oocytes and early embryos, where transcription is repressed and development depends on such regulation of maternal mRNAs [2, 3]. Recent studies in Drosophila have shown that poly(A) tail length regulation has also important functions earlier in the germ line, including in the biology of ovarian germ-line stem cells (GSCs). The CCR4-NOT deadenylation complex, is the predominant deadenylase in all biological systems. Drosophila CCR4-NOT

Aymeric Chartier and Willy Joly contributed equally. Michael Buszczak (ed.), Germline Stem Cells, Methods in Molecular Biology, vol. 1463, DOI 10.1007/978-1-4939-4017-2_7, © Springer Science+Business Media New York 2017

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is composed of eight proteins NOT1–3, NOT10, NOT11, CAF40, and two deadenylases, CCR4 and POP2 [4–6]. The CCR4-NOT complex has the capability to shorten mRNA poly(A) tails without specificity; however, specificity is achieved through the recruitment of the complex to mRNAs by specific RNA binding proteins. The CCR4-NOT complex is involved in GSC self-renewal in the Drosophila ovary [7]. In these cells, CCR4 acts in complex with two major translational repressors, Nanos and Pumilio, which play a key role in GSC maintenance through the translational repression of mRNAs encoding differentiation factors [8, 9]. mei-P26 mRNA has been identified as an important target of CCR4/Nanos/Pumilio-mediated translational repression for GSC self-renewal [7]. meiP26 encodes a protein of the Trim-NHL tumor suppressor family that has conserved functions in stem cell lineages. Translational derepression of mei-P26 mRNA in twin mutant GSCs (twin encodes Drosophila CCR4) correlates with mei-P26 elongated poly(A) tails [7]. Several subunits of the CCR4-NOT complex were also identified in an RNAi screen for their role in either self-renewal or differentiation of GSCs [10]. The specificity of the CCR4-NOT complex for mRNAs depends on RNA binding proteins, and therefore, its dual role in GSC self-renewal and differentiation could result from its interaction with different RNA binding proteins. Consistent with this, a recent study revealed the role of CCR4 and POP2 deadenylases together with Pumilio and Brain Tumor (Brat) in GSC differentiation. This complex acts in cystoblasts (the differentiating daughter cells of GSCs), to repress translation of selfrenewal mRNAs encoding transcriptional regulators [11]. The miRNA pathway also plays a crucial role in GSC selfrenewal [12–14]. A major mechanism of silencing by miRNAs involves deadenylation by the CCR4-NOT complex, through its recruitment by the GW182 protein linked to Argonaute1 (for review see [15]). Therefore, the CCR4-NOT complex is likely to play a role in miRNA-mediated translational repression in the GSCs, thus making this complex a central effector of translational repression in the GSCs. Recent studies have reported methods allowing transcriptomewide measurements of mRNA poly(A) tails through deepsequencing approaches (see for example [3, 16, 17]). However, rapid methods to measure poly(A) tails of specific mRNAs from small quantities of tissues are very useful to address the role of mRNA regulation by poly(A) tail length in GSCs and early germ cells. Here, we described two methods allowing poly(A) tail measurements of specific mRNAs in a few hours from GSCs, cystoblasts, or early germ cells. LM-PAT: The ligation-mediated poly(A) test (LM-PAT or PAT) was set up by Salle´s et al. [18]. It is an RT-PCR-based method using an oligo(dT)-anchor and a gene specific upstream primer. It was

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designed to avoid preferential amplification of shorter poly(A) tails and favor priming at the very end of the poly(A) tail. mRNA poly (A) tails are coated with oligo-(dT)12-18 primers at 42  C, in the presence of T4 DNA ligase. Annealing at this temperature tends to leave short single-stranded 30 oligo(A) overhangs, which are too short to hybridize with oligo-(dT)12-18. This reaction is followed by annealing of the (dT)-anchor primer to the overhanging remaining oligo(A) at 12  C (the lower temperature allows hybridization) and its ligation. This ligated primer is then used as the primer for reverse transcription, which is followed by PCR using (dT)-anchor and a gene specific primer. ePAT: The extension poly(A) test (ePAT) was designed by J€anicke et al. [19]. It is also based on RT-PCR using an oligo(dT)-anchor and a gene specific primer. Here, the (dT)-anchor primer is first annealed to mRNA poly(A) tails at 37  C and used as template for mRNA extension with Klenow polymerase. This reaction is then switched to 55  C to dissociate annealings that had not been extended by Klenow polymerase, and followed by reverse transcription and PCR using (dT)-anchor and a gene specific primer. Both methods result in PCR fragments with a size distribution reflecting the different poly(A) tail lengths of the specific mRNA plus the 30 UTR region that was included in the PCR. Visualization of these PCR products on 2 % agarose gel appears as smears. In our hands, the ePAT produced better results for most genes. For both methods, PCR specificity can be an issue since the upstream primer is the only gene specific primer. This can be solved by using nested specific primers, or by blotting the PCR products and hybridizing with specific probes. Control RNase H digestion can be performed in the presence or the absence of oligo-(dT)12-18, prior to PAT or ePAT, to verify that the smears correspond to poly(A) tails (see [20]).

2

Materials All solutions used for RNA extraction, DNase digestion or PAT synthesis should be made with RNase-free water (commercial RNase-free water). All other precautions to prevent RNase contaminations should be applied: keep solutions used for RNA work in unique area when stored at room temperature, 4  C or 20  C; always use sterile disposable plastic ware and filter tips; wear disposable gloves; and make all steps with care.

2.1 GSC and Cystoblast Generation and Dissection

1. Plastic vials containing standard cornmeal/agar medium. 2. Water bath at 37  C. 3. Stainless steel No. 5 forceps. 4. Microscope slide (SuperFrost®).

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5. Dissecting microscope. 6. 1 Phosphate Buffered Saline (PBS). 7. Fly stocks with the following genotypes: ry506 e1 bamΔ86/TM3, ryRK Sb1 Ser1 ([21]; BL5427 at the Bloomington Drosophila Stock Center). w1118; hs-bam11d/TM3, Sb1 ([22]; BL24637). c587-Gal4 [23]. w*; UAS-dpp ([24]; BL1486). 2.2 Total RNA Extraction and DNase Digestion

1. Trizol® reagent (Invitrogen). Store at 4  C. 2. TURBO DNase (2 U/μL) (Ambion). Store at 3. Phenol–chloroform–isoamyl (Sigma). Store at 4  C.

alcohol

20  C.

mixture

(25:24:1)

4. Chloroform–isoamyl alcohol (24:1) (Fluka). Store at 4  C. 5. 5 M ammonium acetate, 100 mM EDTA. Store at

20  C.

6. Isopropanol. Store at room temperature. 7. 70 % ethanol. Store at room temperature. 2.3 PAT and ePAT Assays

Store all reagents at

20  C

1. DNA Polymerase I, Large (Klenow) Fragment (New England Biolabs). 2. SuperScript® III reverse transcriptase and corresponding buffer (Invitrogen). 3. T4 DNA Ligase (40 U/μL) (New England Biology). 4. dNTP mix (dATP, dGTP, dCTP, dTTP at 10 mM each) from set of deoxyribonucleotides at 100 mM (Promega). 5. 10 mM ATP solution. 6. 50 μM oligo-(dT)-anchor (50 GCGAGCTCCGCGGCCGCG TTTTTTTTTTTT30 ). 7. 0.1 μg/μL oligo-(dT)12-18 (Invitrogen). 8. 50 μM gene specific primer (see Note 1). 9. Recombinant RNasin® Ribonuclease Inhibitor (Promega). 10. Taq DNA Polymerase (cloned) and corresponding buffer (GE Healthcare). 11. Thermocycler (Eppendorf ).

2.4

PAT Visualization

1. 3 μg of 50 bp DNA ladder (New England Biolabs). Store at 20  C. 2. 6 PAT Loading Solution: 0.25 % bromophenol blue, 30 % glycerol.

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Methods

3.1 Generation of GSCs, Cystoblasts, and Early Germ Cells

1. GSC-like cells are obtained using the bamΔ86 mutant (Bag of marble (Bam) is the major GSC differentiation factor [21]). bamΔ86 newly eclosed homozygote females are collected and kept for 7 days (to increase the number of GSCs) in vials with wild-type males on standard medium supplemented with fresh yeast (30 females per tube) at 25  C. Ovaries are then dissected. GSCs can also be produced from females bearing the transgenes c587-Gal4 and UAS-dpp (see Note 2). 2. The production of cystoblasts is induced by the expression of Bam in bamΔ86 homozygous females carrying the hs-bam transgene, following heat shock [22]. We use the stock hs-bam11d e1 bamΔ86/TM3 Sb to generate bamΔ86 homozygote females carrying one copy of the hs-bam transgene. To perform the heat shock, vials containing 7 day-old females kept at 25  C with wild-type males on standard medium supplemented with fresh yeast are plunged in a water bath at 37  C for 2 h. The vials are then placed back in the 25  C incubator for 18 h and ovaries are dissected. Cystoblasts can also be produced from females bearing the three transgenes c587-Gal4, UAS-dpp and hs-bam following heat shock (see Note 3). 3. To produce early germ cells, newly eclosed females are collected and kept on standard medium supplemented with fresh yeast for 4–6 h. Ovaries are then dissected. Ovaries from these 1 day-old females contain ovarian stages from germarium to stage 8.

3.2 Dissection, RNA Extraction and DNase Digestion

1. Dissection: females are anesthetized, placed on a microscope slide and ovaries are dissected in cold 1 PBS with forceps, collected in 1.5 mL tubes and immediately frozen on dry ice. 100–200 ovaries are used for total RNA extraction. 2. RNA extraction: ovaries are crushed in 1 mL TRIzol® and total RNA is extracted using manufacturer’s instructions. 3. DNase digestion: Dissolve the RNA pellet in 26 μL of H2O and add 3 μL of 10 DNase Buffer and 1 μL TURBO DNase (see Note 4). 4. Vortex quickly. 5. Incubate 30 min at 37  C.

3.3 Phenol– Chloroform Extraction and Isopropanol Precipitation

1. 30 μL of RNA treated with DNase is mixed with 15 μL of 5 M ammonium acetate, 100 mM EDTA and the volume is set to 150 μL with H2O (see Note 5). 2. Add 150 μL of phenol–chloroform and vortex vigorously. 3. Centrifuge at 12,000  g for 2 min in a microcentrifuge.

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4. Discard lower phase (organic phase) by aspiration. 5. Add 150 μL of chloroform on aqueous phase and vortex vigorously. 6. Centrifuge at 12,000  g for 2 min in a microcentrifuge. 7. Discard lower phase (organic phase) by aspiration. 8. Add 150 μL of chloroform on aqueous phase and vortex vigorously. 9. Centrifuge at 12,000  g for 2 min in a microcentrifuge. 10. Discard lower phase (organic phase) by aspiration. 11. Centrifuge the aqueous phase at maximum speed for 30 s in a microcentrifuge. This separates the remaining chloroform at the bottom of the tube. The excess of chloroform is discarded. 12. Transfer the aqueous phase in a new tube and add 1 volume of isopropanol. 13. Incubate at

20  C for 15 min.

14. Centrifuge at maximum speed for 25 min at 4 microcentrifuge.



C in a

15. Wash RNA pellet with at least 500 μL of 70 % ethanol. 16. Centrifuge at 8000  g for 5 min at 4  C in a microcentrifuge. 17. Air-dry the RNA pellet and dissolve the pellet with 10 μL of H2O. 18. Determine the RNA concentration by spectrophotometry. 3.4 LM-PAT cDNA Synthesis

1. Mix on ice 1 μg of total RNA digested by the DNAse with 1 μL of 0.1 μg/μL oligo-(dT)12-18 and H2O to obtain a RNA mixture with a final volume of 7 μL. 2. For each sample, mix on ice 4 μL of H2O, 4 μL of 5 SuperScript® III Reverse transcriptase buffer, 1 μL 0.1 M DTT, 1 μL 10 mM dNTP mix, 1 μL 10 mM ATP, 1 μL recombinant RNasin® Ribonuclease Inhibitor (40 U/μL), and 1 μL T4 DNA Ligase to obtain a BUFFER mixture with a final volume of 13 μL. 3. Incubate the RNA mixture for 5 min at 65  C in a thermocycler and cool down the mixture at room temperature. 4. Centrifuge the RNA mixture at maximum speed for 15 s in a microcentrifuge. 5. Warm the RNA mixture and BUFFER mixture separately for 2 min at 42  C. 6. At 42  C, transfer 13 μL of BUFFER mixture into the RNA mixture; mix by pipetting up and down and incubate for 30 min at 42  C. 7. Add 1 μL of 50 μM oligo-(dT)-anchor.

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8. Incubate the reaction for 2 h at 12  C. 9. Incubate the reaction for 2 min at 42  C. 10. Add 1 μL of SuperScript® III reverse transcriptase, mix by pipetting and incubate at 42  C for 1 h. 11. Inactivate the enzyme at 70  C for 10 min. 12. Store on ice for the PCR or store at 3.5 ePAT cDNA Synthesis

20  C for further use.

1. Mix on ice 1 μg of total RNA digested by the DNAse with 2 μL of 50 μM oligo-(dT)-anchor and H2O to obtain a RNA mixture with a final volume of 12 μL. 2. For each sample, mix on ice 4 μL of 5 SuperScript® III Reverse transcriptase buffer, 1 μL 0.1 M DTT, 1 μL 10 mM dNTP mix, and 1 μL recombinant RNasin® Ribonuclease Inhibitor (40 U/μL) to obtain a BUFFER mixture with a final volume of 7 μL. 3. Incubate the RNA mixture for 5 min at 68  C in a thermocycler and cool down the mixture at room temperature. 4. Centrifuge the RNA mixture at maximum speed for 15 s in a microcentrifuge. 5. Warm the RNA mixture and BUFFER mixture separately for 2 min at 37  C. 6. At 37  C, transfer 7 μL of BUFFER mixture into the RNA mixture; mix by pipetting up and down. 7. Add 1 μL of DNA Polymerase I, Large (Klenow) Fragment (5 U/μL) and mix by pipetting at 37  C. 8. Incubate the reaction for 1 h at 37  C. 9. Inactivate the DNA Polymerase I for 20 min at 70  C. 10. Centrifuge at maximum speed for 15 s in a microcentrifuge. 11. Add 1 μL of SuperScript® III reverse transcriptase, mix and incubate at 55  C for 1 h. 12. Inactivate the enzyme at 70  C for 10 min. 13. Store on ice for the PCR or store at

3.6

PCR

20  C for further use.

1. For each sample to be analyzed, set up a 50 μL reaction mix containing 1 μL of PAT- or ePAT-cDNAs from Subheadings 3.4 or 3.5, 200 μM dNTP, 500 nM of oligo-(dT)-anchor primer, 500 nM of gene specific primer, 1 Taq Buffer with MgCl2 and 0.5 μL of Taq DNA Polymerase (5 U). 2. Appropriate conditions have to be determined for each gene analyzed (see Notes 1 and 6), but most often, the following conditions with an annealing temperature of 58  C can be used: 30 s 94  C; (10 s 94  C, 15 s 58  C, 45 s 72  C) 35 cycles; 5 min 72  C; and cooling down to 15  C.

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3. Mix 25 μL of PCR reaction with 5 μL of 6 PAT Loading Solution. 4. Prepare a 1 TBE agarose gel with ethidium bromide (0.9 μg/ mL ethidium bromide in 2 % agarose gel). A 2 % agarose and 24 cm-length gel is sufficient to separate the PCR products (see Note 7). 5. Load a lane with 3 μg of 50 bp DNA ladder and the samples in the following lanes. 6. Run the gel in 1 TBE up to 60–80 % of the gel length at 5 V/cm. 7. Visualize the DNA using a UV table.

4

Notes 1. The gene specific primer is designed following the PCR primer rules. The primer must be 19–23 nt long and the melting temperature around 58–60  C. Primer design can be done with the help of online software, such as Primer3Plus (http:// www.bioinformatics.nl/cgi-bin/primer3plus/primer3plus.cgi/) [25], which gives a primer list for a specific DNA fragment. The matching region of the primer in the 30 UTR of a specific gene must be localized 150–300 nucleotides upstream of the poly(A) site. When several poly(A) sites have been identified in a gene, the different poly(A) tails can be visualized with a single gene specific primer if these poly(A) sites are close. If they are too far apart (more than 300 nucleotides), a primer has to be designed for each poly(A) site. 2. GSC-enriched ovaries can also be obtained from flies carrying the transgenes c587-Gal4 and UAS-dpp. In ovaries of these flies, Dpp is overexpressed in the somatic cells surrounding the stem cell niche resulting in accumulation of GSCs (through the prevention of bam expression) [26]. It is noteworthy that bamΔ86 females produce a higher purity of GSC content since no GSCs can differentiate in bam mutant females, whereas differentiation is possible in c587-Gal4/þ; UAS-dpp/þ females. Therefore, when using females of this genotype, it is crucial to only dissect the very anterior part of ovarioles to have a pure population of GSCs. 3. Cystoblast-enriched ovaries can be obtained from flies containing the three transgenes c587-Gal4, UAS-dpp and hs-bam following heat shock. These females are produced by crossing c587-Gal4; hs-bam with UAS-dpp stocks. The heat-shock is identical as for hs-bam11d e1 bamΔ86/bamΔ86 females and the dissection is also after 18 h-recovery at 25  C following heat shock. As for Dpp-expanded GSCs, the purity of cystoblasts is not complete with this genotype and the very anterior part of ovarioles must be dissected to enrich in cystoblasts.

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4. TURBO DNase can be supplied as TURBO DNase-free™ kit working with DNase Inactivation reagent. However, in our hands, this reagent inhibits the PAT-cDNA synthesis, therefore DNase inactivation is replaced by phenol–chloroform extraction and precipitation. 5. Sodium acetate and ethanol precipitation can be performed instead of ammonium acetate and isopropanol precipitation. In that case modify the sodium acetate and ethanol volumes for RNA precipitation. For highly concentrated RNA solution, an RNA clean up kit (Qiagen) can be used directly after the DNase digestion. In that case refer to the manufacturer’s instructions. 6. The annealing temperature can vary between gene specific(dT)-anchor primer pairs from 58 to 61  C. 7. Standard agarose, such as SeaKem® LE Agarose (Lonza) can be used routinely but if a higher resolution is required, use a higher resolution agarose such as UltraPure™ Agarose 1000 (Invitrogen). The percentage of agarose can vary from 2 to 4 %.

Acknowledgements Work in the Simonelig lab is supported by the CNRS UPR1142, ANR (ANR-2010-BLAN-1201 01 and ANR-15-CE12-0019-01), FRM (Equipe FRM 2013 DEQ20130326534), and AFMTelethon (No 17110). References 1. Weill L, Belloc E, Bava FA, Mendez R (2012) Translational control by changes in poly(A) tail length: recycling mRNAs. Nat Struct Mol Biol 19:577–585 2. Benoit P, Papin C, Kwak JE, Wickens M, Simonelig M (2008) PAP- and GLD-2-type poly(A) polymerases are required sequentially in cytoplasmic polyadenylation and oogenesis in Drosophila. Development 135:1969–1979 3. Subtelny AO, Eichhorn SW, Chen GR, Sive H, Bartel DP (2014) Poly(A)-tail profiling reveals an embryonic switch in translational control. Nature 508:66–71 4. Temme C, Simonelig M, Wahle E (2014) Deadenylation of mRNA by the CCR4-NOT complex in Drosophila: molecular and developmental aspects. Front Genet 5:143 5. Temme C, Zaessinger S, Meyer S, Simonelig M, Wahle E (2004) A complex containing the CCR4 and CAF1 proteins is involved in mRNA

deadenylation in Drosophila. EMBO J 23:2862–2871 6. Temme C, Zhang L, Kremmer E, Ihling C, Chartier A et al (2010) Subunits of the Drosophila CCR4-NOT complex and their roles in mRNA deadenylation. RNA 16: 1356–1370 7. Joly W, Chartier A, Rojas-Rios P, Busseau I, Simonelig M (2013) The CCR4 deadenylase acts with Nanos and Pumilio in the fine-tuning of Mei-P26 expression to promote germline stem cell self-renewal. Stem Cell Rep 1:411–424 8. Gilboa L, Lehmann R (2004) Repression of primordial germ cell differentiation parallels germ line stem cell maintenance. Curr Biol 14:981–986 9. Wang Z, Lin H (2004) Nanos maintains germline stem cell self-renewal by preventing differentiation. Science 303:2016–2019

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10. Yan D, Neumuller RA, Buckner M, Ayers K, Li H et al (2014) A regulatory network of Drosophila germline stem cell self-renewal. Dev Cell 28:459–473 11. Newton FG, Harris RE, Sutcliffe C, Ashe HL (2015) Coordinate post-transcriptional repression of Dpp-dependent transcription factors attenuates signal range during development. Development 142:3362–3373 12. Jin Z, Xie T (2007) Dcr-1 maintains Drosophila ovarian stem cells. Curr Biol 17:539–544 13. Park JK, Liu X, Strauss TJ, McKearin DM, Liu Q (2007) The miRNA pathway intrinsically controls self-renewal of Drosophila germline stem cells. Curr Biol 17:533–538 14. Yang L, Chen D, Duan R, Xia L, Wang J et al (2007) Argonaute 1 regulates the fate of germline stem cells in Drosophila. Development 134:4265–4272 15. Braun JE, Huntzinger E, Izaurralde E (2012) A molecular link between miRISCs and deadenylases provides new insight into the mechanism of gene silencing by microRNAs. Cold Spring Harb Perspect Biol 4(12) 16. Chang H, Lim J, Ha M, Kim VN (2014) TAILseq: genome-wide determination of poly(A) tail length and 30 end modifications. Mol Cell 53:1044–1052 17. Harrison PF, Powell DR, Clancy JL, Preiss T, Boag PR et al (2015) PAT-seq: a method to study the integration of 30 -UTR dynamics with gene expression in the eukaryotic transcriptome. RNA 21:1502–1510 18. Salles FJ, Richards WG, Strickland S (1999) Assaying the polyadenylation state of mRNAs. Methods 17:38–45

19. Janicke A, Vancuylenberg J, Boag PR, Traven A, Beilharz TH (2012) ePAT: a simple method to tag adenylated RNA to measure poly(A)-tail length and other 30 RACE applications. RNA 18:1289–1295 20. Chartier A, Klein P, Pierson S, Barbezier N, Gidaro T et al (2015) Mitochondrial dysfunction reveals the role of mRNA poly(A) tail regulation in oculopharyngeal muscular dystrophy pathogenesis. PLoS Genet 11:e1005092 21. McKearin D, Ohlstein B (1995) A role for the Drosophila bag-of-marbles protein in the differentiation of cystoblasts from germline stem cells. Development 121:2937–2947 22. Ohlstein B, McKearin D (1997) Ectopic expression of the Drosophila Bam protein eliminates oogenic germline stem cells. Development 124:3651–3662 23. Manseau L, Baradaran A, Brower D, Budhu A, Elefant F et al (1997) GAL4 enhancer traps expressed in the embryo, larval brain, imaginal discs, and ovary of Drosophila. Dev Dyn 209:310–322 24. Staehling-Hampton K, Hoffmann FM (1994) Ectopic decapentaplegic in the Drosophila midgut alters the expression of five homeotic genes, dpp, and wingless, causing specific morphological defects. Dev Biol 164:502–512 25. Untergasser A, Nijveen H, Rao X, Bisseling T, Geurts R et al (2007) Primer3Plus, an enhanced web interface to Primer3. Nucleic Acids Res 35:W71–W74 26. Kai T, Williams D, Spradling AC (2005) The expression profile of purified Drosophila germline stem cells. Dev Biol 283:486–502

Chapter 8 Identification of Germ-Line Stem Cells in Zebrafish Bruce W. Draper Abstract Both male and female zebrafish have a population of germ-line stem cells that produce gametes throughout the life of the fish. These cells localize to specific regions in the gonads and can be identified because they uniquely express the nanos2 gene, which encodes a conserved regulator of translation. A method is presented here for identifying germ-line stem cells in the ovary and testis using a combined protocol for whole-mount fluorescent RNA in situ hybridization to detect nanos2 mRNA and immunofluorescence to detect the pan-germ cell marker Vasa. Key words Zebrafish germ-line stem cells, Oogonial stem cell, Spermatogonial stem cell, Zebrafish ovary, Zebrafish testis, nanos2, Vasa, Fluorescent RNA in situ hybridization, Confocal immunofluorescence

1

Introduction The zebrafish has become an excellent model organism for studying vertebrate development. Zebrafish have a rapid generation time and are very amenable to both forward and reverse genetics. While most research has been focused on embryonic development, many labs have begun to investigate events that do not occur until later in life, such as the specification and maintenance of germ-line stem cells (GSC). Unlike mammals, where GSCs are only present in the adult testis, GSCs in zebrafish are present in both the adult testis and ovary [1, 2]. Members of the Nanos family of conserved translational regulators are involved in the development and maintenance of germ-line stem cells in most, if not all, organisms. Three Nanos ortholog-encoding genes are present in the genomes of most vertebrates (nanos1-3), and in several of these, including zebrafish, nanos2 has been shown to specifically mark germ-line stem cells [2–4]. The GSCs in zebrafish are only a small fraction of the total number of premeiotic germ cells that are present in both the testis

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and ovary. However, they can be easily identified because they localize to specific regions within each tissue, have a characteristic nuclear morphology and uniquely express nanos2. The organizational unit of the zebrafish testis is the lobule, which is a somewhat spherical unit that has premeiotic spermatogonia and early meiotic spermatocytes localizing to the surface of the sphere while the mature sperm localize to its center. In ovaries, the GSCs localize to the germinal zone, which is a discrete circumferential band of cells on the medial and lateral ovarian surface [1, 2]. Because the GSCs in both testes and ovaries are located on the surface of the tissue, it is not necessary to section the tissue prior to staining. Instead, the GSC can be identified in whole-mount tissue preparations. Below we describe a method for the identification of GSCs that is a combination of whole-mount fluorescent RNA in situ hybridization, to visualize nanos2 expressing cells, and immunofluorescence, to visualize the pan-germ cell marker protein Vasa (Fig. 1). The in situ protocol has been adapted from Thisse and Thisse [5] and the Vasa immunofluorescences protocol from Draper [6]. This procedure will take 6 days to complete.

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Materials

2.1 Tissue Fixation and Permeabilization

1. 10 Phosphate Buffered Saline (PBS): 1.37 M NaCl, 27 mM KCl, 100 mM Na2HPO4, 17.6 mM KH2PO4. Autoclave and store at room temperature. Prepare a working solution by diluting tenfold with sterile water. 2. 8 % Paraformaldehyde (PFA) Stock: 40 g paraformaldehyde in 400 ml water. While stirring heat to 60  C, add 1 N NaOH drop-wise until paraformaldehyde is dissolved. Adjust to 500 ml, filter through a 45 μm vacuum filter, aliquot into 50 ml conical tubes and store frozen at 20  C for up to 1 year. Once thawed, use within 1 week. 3. 1 Fixative: 4 % PFA in 1 PBS. 4. Sylgard-coated Petri dishes: Mix Sylgard 184 Silicone Elastomer (Fisher) parts A and B in 10:1 ratio (w/w). Fill Petri dishes (10m and 3.5 cm) one-third full and incubate at 50  C overnight, or until hardened. 5. Dumont No. 5 forceps. 6. Angled scissors: e.g., Supercut 10 cm angled scissors. 7. Proteinase K: 10 mg/ml proteinase K stock solution (PCR grade) in PBT. Store 500 μl aliquots at 20  C. Final working concentration will be 50 μg/ml. 8. 100 % methanol.

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Probe Synthesis

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1. Purified DNA template: nanos2 cDNA clone pBD77 [2], linearized with SpeI enzyme (see Note 1). 2. PCR purification kit (Thermo Scientific). 3. 10 DIG RNA labeling mix (Roche). 4. T7 RNA polymerase (Roche). 5. Dithiothreitol (DTT). 6. RNase inhibitor (Roche). 7. 0.5 M EDTA, pH 8.0. 8. tRNA (RNase free): Resuspend tRNA, Type V, from wheat germ, in water and extract 3 with phenol–chloroform to remove proteins (or until no precipitate is visible at the interphase). Precipitate tRNA by adding 1/10th volume 5 M LiCl, 3 volume 100 % ethanol, and store at 20  C for 30 min before pelleting in a centrifuge. Resuspend at 50 mg/ml in RNase-free water. 9. 5 RNA loading dye: 0.25 % bromophenol blue (w/v), 0.25 % xylene cyanol FF (w/v), 1 % sodium dodecyl sulfate (SDS; v/v), and 30 % glycerol (v/v). 10. 1 % Agarose 0.5 TBE gel.

2.3 nanos2 In Situ Hybridization

1. PBT: 1 PBS, 0.1 % (v/v) Triton X-100. 2. 20 SSC: Dissolve 175.3 g NaCl, 88.2 g citric acid trisodium salt in 1 l water. Autoclave. 3. 50 mg/ml heparin: Heparin sodium salt from porcine intestinal mucosa, dissolved in sterile water. 4. Prehybridization solution: 50 % deionized formamide, 5 SSC, 50 μg/ml heparin, 500 μg/ml RNase-free tRNA, 0.1 % Tween, pH to 6.0 by adding 460 μl 1 M citrate/50 ml pre-hyb. Store indefinitely at 20  C. 5. Hybridization solution: 1.8 ml prehybridization solution, 200 μl 50 % dextran sulfate (5 % final concentration; see Note 1) and 10 μl nanos2 RNA probe (1:200 final dilution). Preheat to 65  C before use. 6. Antibody Blocking Solution: PBT, 2 % sheep serum (v/v), 2 mg/ml bovine serum albumin, fraction V, 99 % pure. 7. AP-Anti-DIG: Sheep anti-DIG FAb conjugated to alkaline phosphatase. 8. Alkaline coloration buffer: 100 mM Tris-HCl, pH 9.5, 50 mM MgCl2, 100 mM NaCl, 0.1 % Tween 20.d 9. SigmaFast™ Fast Red TR/Naphthol AS-MX tables (Sigma). 10. Labeling solution: Dissolve one Fast Red tablet in 1 ml alkaline coloration buffer just before use.

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2.4 Vasa Immunofluorescence

1. Primary antibody: rabbit anti-Vasa polyclonal sera [7]. 2. Secondary antibody: Alexa Fluor 488 goat anti-rabbit IgG. 3. 40 ,6-diamidino-2-phenylindole (DAPI): 10 mg/ml stock in water, stored in dark at 4  C.

2.5 Mounting and Confocal Microscopy

1. Graded glycerol series (30, 50, and 75 % diluted in PBT). 2. Vacuum grease. 3. 25  75  1 mm microscope slides. 4. 22  22 mm coverslips.

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Methods In adult zebrafish the ovarian germ-line stem cells (GSCs) localize to the germinal zone, a discrete circumferential band that runs along the lateral and medial surfaces of the ovary, oriented parallel to the anterioposterior axis [1, 2]. However, because of the large size of the adult gonad it can be difficult to reliable identify the germinal zone. By contrast, GSCs can more easily be identified in the juvenile ovary (40–50 dpf) because it is relatively planer and most of the GSCs localize to its margins, the presumed precursor to the germinal zone, and thus are easier to image using the confocal microscope [2]. In males, the germ cells are organized into spherical “lobules” and the germ-line stem cells as well as clones of various stage spermatocytes, localize predominantly to the surface of the lobules, whereas mature sperm are found interior [2]. In preparation for this procedure it is important to note that the zebrafish ovary, in particular, is very fragile so it is necessary that they be fixed while still in the body cavity in order to maintain their ultrastructural. While the testes are less delicate, they tend to curl if dissected prior to fixation, making imaging more challenging. For these reasons it is recommended that both testes and ovaries be fixed while in situ. The method outlined below details how to identify nanos2expression GSCs using fluorescent RNA in situ hybridization, coupled with an anti-Vasa antibody staining to label all premeiotic germ cells and early stage oocytes. In addition, all nuclei are counterstained with the DNA dye DAPI, which allows accurate germ cell staging based on nuclear morphology.

3.1 Preparation of Template DNA

1. If using the nanos2 cDNA plasmid as a template, linearize 10 μg by digestion with SpeI restriction enzyme in a volume of 50 μl. To ensure complete digestion, incubate at 37  C for a minimum of 2 h. Assess the completeness of digestion by running 1 μl of the digest on a 0.8 % TBE gel with un-digested plasmid as a control. Alternatively, a DNA template can be produced by PCR (see Note 2). 2. Purify linearized template using a PCR purification kit. Elute in 30 μl RNase-free elution buffer and then dilute to 500 ng/μl.

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1. Assemble the components listed in Table 1 in the designated order in an RNase-free 1.7 ml snap-cap tube (see Note 3). 2. Incubate at 37  C for 1 h. 3. Add an additional 1 μl of T7 RNA polymerase and incubate for an additional 1 h at 37  C. 4. Stop the transcription reaction by adding 0.8 μl 0.5 M EDTA, pH 8.0. 5. Remove 1 μl of the reaction to run on a 1 % agarose TBE gel containing ethidium bromide (or appropriate nucleic acid stain), using the RNA loading buffer, and an appropriate DNA molecular weight marker (e.g., 1 kb ladder). Because this is not a denaturing gel, the RNA will usually run as multiple bands, with one band being significantly brighter than the others. All RNA bands will run below the template DNA band, and the brightest RNA band should be at least 3 brighter than the DNA template band. Importantly, all bands should be discrete, and you should not see a “smear,” which would indicate degraded RNA. The nanos2 RNA probe made with the pBD77 plasmid template has a major band at 2 kb, and a minor band at 3.5 kb. 6. While the gel is running, precipitate the RNA probe as follows: Add 1 μl 50 mg/ml RNase-free tRNA, 2 μl 5 M LiCl, 75 μl 100 % ethanol. Gently mix and incubate at 20  C for 30 min. Pellet precipitated RNA using a 4  C-cooled microcentrifuge (20 min at 14K RPM/20Kg). Rinse pellet with 75 % ethanol. Let pellet slightly dry for 5 min at room temperature. Resuspend the RNA probe in 100 μl prehybridization solution and store at 80  C indefinitely.

Table 1 In vitro transcription reaction component for in situ probe synthesis Component

Volume

H2O (RNase-free)

11.5 μl

10 transcription buffer

2 μl

10 DIG labeling mix

2 μl

Linearized DNA template (500 ng/μl)

2 μl

100 mM DTT

1 μl

RNase inhibitor

0.5 μl

T7 RNA polymerase

1 μl

Vt

20 μl

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3.3 Preparation of Gonads for In Situ Hybridization and Indirect Immunostaining

Day 1

1. Fix gonads in situ overnight in 1 PBS/4 % paraformaldehyde as follows: First, euthanize fish by immersion in 0  C ice water for a minimum of 5 min (hypotermal shock). Next, decapitate fish by cutting just posterior to the gills using angled scissors. Finally, cut open the body cavity along the ventral midline, starting at the anterior and proceeding to the cloaca. To ensure that the tissue is properly and rapidly fixed, place no more than ten 40–50-day-old fish/30 ml of fixative, or no more than three adults/30 ml of fixative. Day 2

2. Following fixation, wash fish 3 10 min in PBT. 3. Dissect gonads from the body cavity, using a Sylgard-coated petri dish flooded with PBT and No. 5 Dumont forceps. It is useful to stabilize the fish by impaling one set of forceps into the tail just posterior to the cloaca. Using the other set of forceps, grasp the skin and body wall muscle close to the midline and peel skin and muscle off of torso to expose the gonads. The ovaries and testes are bilaterally located dorsal to the stomach and liver and ventrolateral to the swim bladder. Both ovaries and testes may have a layer on lipid covering their dorsal surface after dissection. This should be gently removed so that the tissue will sink rather than float. 4. Wash dissected ovaries in PBT 3 15 min. 5. Methanol permeabilization: Rinse ovaries 2 5 min in 100 % MeOH, then add fresh 100 % MeOH and incubate for 1 h at 20  C. Next, rehydrate ovaries in a graded MeOH series in (70, 50, 30 % MeOH in PBT) 10 min each for 40–50 dpf gonads and 20 min each for adult gonads (see Note 4). Wash 3 5 min in PBT. 6. Proteinase K permeabilization: Place ovaries in a solution of 10 μg proteinase K/ml PBT for 10 min at 37  C (see Note 5). 7. Inactivate the proteinase K by refixing ovaries in 4 % PFA/1 PBS for 20 min. 8. Wash ovaries 3 10 min in PBT. 3.4 Whole-Mount In Situ Hybridization

1. Prehybridization: Place permeabilized gonads into a 1.7 ml snap-cap tube and add 1 ml of room temperature prehybridization solution. If large adult-size ovaries are being hybridized, place them into 5 ml snap-cap tubes (Falcon) and use 2–3 ml prehybridization solution (enough to cover tissue). Place tubes into a floating rack in a 65  C circulating water bath. Incubate for a minimum of 1 h. 2. Hybridization: After removing and discarding the prehybridization solution from the tubes, add enough preheated

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hybridization solution to the tubes to cover all of the tissue (typically 500 μl–1 ml). Replace tubes into the 65  C water bath and incubate overnight (minimum 16 h). Because the hybridization solution contains dextran sulfate the tissue will initially float to the top of the solution, but will eventually sink as it equilibrates. Day 3

3. Make the solutions in Table 2 and preheat to 65  C: 4. Remove and save the hybridization solution. Carefully remove the hybridization mix and add back to any mix that remained in the probe stock tube (see Note 6). Store at 20  C. 5. Wash the gonads as outlined in Table 2. Once the washes reach the room temperature steps, the tubes should be gently rocked using a rotating platform. 3.5 Antibody Staining

1. Incubate the gonads in Antibody Blocking Solution for 1 h. 2. 1 antibody labeling: Incubate gonads overnight at 4  C in Antibody Blocking Solution containing goat anti-DIG FAb (1:5000 dilution) and rabbit anti-Vasa antibody (1:2000 dilution). Tubes should be gently rocked on a rotating platform. Day 4

3. Remove 1 antibody and discard. Wash 2 quickly, then 4 15 min, then 2 30 min in PBT at room temperature with gentle rocking. 4. Incubate the gonads in Antibody Blocking Solution for 1 h. 5. 2 antibody labeling: Incubate gonads overnight at 4  C in Antibody Blocking Solution containing goat anti-rabbit Alexa Fluor 488-labeled antibody (1:300 dilution). Table 2 Post-hybridization washes Wash duration

Solution

Temperature

1 5 min

1:2 mixture of Prehybridization: 2 SSC

65  C

1 5 min

2:1 mixture of Prehybridization: 2 SSC

65  C

1 5 min

2 SSC

65  C

2 20 min

0.2 SSC

65  C

1 20 min

0.1 SSC

65  C

1 5 min

1:1 mixture 2 SSC; PBSTw

RT

2 5 min

PBSTw

RT

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Day 5

6. Remove 2 antibody and discard. Wash 2 quickly, then 4 15 min, then 2 30 min in PBT at room temperature with gentle rocking. 7. Develop the nanos2 in situ: Wash 3 10 min in Coloration buffer. 8. Dissolve one tablet of SigmaFast Fast Red in 1 ml of Coloration buffer, and then add this to gonads. While the reaction is developing, the tubes should be kept in the dark, either in a drawer or wrapped in foil. 9. Allow the in situ reactions to develop in the dark for a minimum of 2 h and a maximum of 4 h. Monitor the reactions using a dissecting microscope with top lighting. At this stage it is useful to transfer the gonads to a watch glass. The signal will appear as punctate red dots. However, because germ-line stem cells are a very small fraction of the total cell population, it may be difficult to identify them at low magnification. 10. Stop the reaction by washing 2 quickly in water, then 4 15 min, then 2 30 min in PBT þ 2 μg/ml DAPI at room temperature with gentle rocking. 11. Place gonads in 30 % glycerol in PBT þ 2 μg/ml DAPI, and incubate overnight at 4  C. This step assures that the nuclei are properly stained with DAPI. 3.6 Mounting Ovaries for Confocal Analysis

Day 6

1. To mount ovaries for microscopy, the gonads are cleared by further dehydrating in a glycerol series, as follows (see Note 7): (a) 50 % glycerol in PBT þ 2 μg/ml DAPI, 1 h at room temp. (minimum). (b) 75 % glycerol in PBT, 1 h at room temp. (minimum). The gonads will initially float in the 50 % glycerol solution, but will sink as they equilibrate. By contrast, after equilibration in the 75 % glycerol solution gonads may not sink. 2. Mount gonads on a microscope slides in 75 % glycerol/PBT as follows: Using a 3 ml syringe filled with vacuum grease, place four spots (“post”) on a microscope slide in a square pattern that is just smaller than the coverslip. Place the gonad in the middle of the posts in a small volume of 75 % glycerol/PBT. Ovaries should be arranged dorsal side up for 40–50 dpf (see Note 8). If mounting an adult ovary, it will be necessary to carefully dissect off the lateral surface while leaving the most mature oocytes behind. This is best done with sharpened tungsten needles. Finally, place a coverslip on the vacuum grease post and, while viewing under a dissection microscope, gently depress coverslip until it is in contact with, and slightly

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depresses, the gonads. Care should be taken to not over-distort the tissue. Add enough 75 % glycerol/PBT to fill underneath the coverslip. 3. Collect images of the stained sample using a confocal microscope equipped with 405, 488, and 543 lasers to visualize the DNA, Vasa protein, and nanos2 mRNA, respectively. Figure 1 shows representative images of nanos2- expression GSCs (red) and anti-Vasa (green) which labels all germ cells. In addition, all nuclei are counterstained with DAPI to allow accurate staging based on nuclear morphology. For staging purposes, it is recommended to present the nuclear staining (DAPI) as a black and white image for maximum contrast (Fig. 1c, f).

Fig. 1 Identification of germ-line stem cells in the zebrafish ovary and testis. Merged panels (a, b, e, f, d) show nanos2 RNA (red) Vasa protein (green) and DNA (blue). Panels c and g show merged nanos2 and DNA only. Panels d and h show DNA only. Low magnification views of nanos2-positive germ-line stem cells (GSC; arrows) that localize to the lateral edge of a 40-day-old ovary (outlined in a) or on the surface of a testicular lobule (outlined in e). Higher magnification views of a single nanos2-positive ovarian (b) or testicular (f) GSC within clusters of nanos2-negative premeiotic germ cells and early meiotic cells. Note that premeiotic germ cells can be identified as having nuclei with a single prominent nucleolus, which appears as holes in the staining. GSC germ-line stem cell, Oo oogonia, Sp spermatogonia, IAp pachytene-stage IA oocytes, IB stage IB oocytes; Scale bar: 20 μm in a, b, f, 50 μm in e

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Notes 1. Because nanos2 is a single exon gene, template DNA can be easily produced by PCR using genomic DNA and the following primers: nos2 Fwd: CATGGGCAAAACACACCTAAAACA/ T7-nos2-Rev: GGCGTAATACGACTCACTATAGGGGAAG CTGCAGACCTGGAAACAAAT (T7 promoter is underlined). For detailed method, see Thisse and Thisse [5]. 2. If using a template that was produced by PCR, it must be first purified using a PCR purification kit. Then use 250 ng PCR product/transcription reaction. 3. Gonads can be stored for several months in 100 % MeOH at 20  C without significantly affecting this procedure. 4. Each batch of proteinase K must be tested for optimal concentration to use in this step. 5. Dextran sulfate is a molecular crowding agent that increases the effective probe concentration in the hybridization solution. This helps to increase the signal without increasing the noise. 6. Hybridization solution can be reused multiple times with little loss of activity. In fact the signal to noise ratio often improves after repeated use. 7. The precipitate that is formed by the alkaline phosphatase enzyme using the Fast Red substrate is slightly water-soluble. Because of this, it is important to image your samples soon after developing them to maintain maximum fluorescence intensity (no more than 2 days post-development). 8. 40–50-day-old ovaries, while mostly planer, have a slight curvature such that the convex and concave surfaces represent the dorsal and ventral surfaces, respectively. Once ovaries are on the slide, they can be flipped and positioned using fine eyelashes glued to tooth picks (hair from some short-haired dog breeds work well also).

Acknowledgments The author would like to thank Holger Knaut for the anti-Vasa antibodies. This work was supported by a NIH grant R01HD081551.

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References 1. Draper BW, McCallum CM, Moens CB (2007) nanos1 is required to maintain oocyte production in adult zebrafish. Dev Biol 305:589–598 2. Beer RL, Draper BW (2013) nanos3 maintains germline stem cells and expression of the conserved germline stem cell gene nanos2 in the zebrafish ovary. Dev Biol 374:308–318 3. Sada A, Suzuki A, Suzuki H, Saga Y (2009) The RNA-binding protein nanos2 is required to maintain murine spermatogonial stem cells. Science 325:1394–1398 4. Nakamura S, Kobayashi K, Nishimura T et al (2010) Identification of germline stem cells in the ovary of the teleost medaka. Science 328:1561–1563

5. Thisse B, Thisse C (2014) In situ hybridization on whole-mount zebrafish embryos and young larvae. In: Nielsen BS (ed) In situ hybridization protocols, vol 1211, Methods in molecular biology. Springer Science þ Business Media, New York, NY, pp 53–67 6. Draper BW (2012) Identification of oocyte progenitor cells in the zebrafish ovary. In: Mace KA, Braun KM (eds) Progenitor cells: methods and protocols, vol 916, Methods in molecular biology. Springer Science þ Business Media, New York, NY, pp 157–165 7. Knaut H, Pelegri F, Bohmann K et al (2000) Zebrafish vasa RNA but not its protein is a component of the germ plasm and segregates asymmetrically before germline specification. J Cell Biol 149:875–888

Chapter 9 Primordial Germ Cell Isolation from Xenopus laevis Embryos Amanda M. Butler, Tristan Aguero, Karen M. Newman, and Mary Lou King Abstract Primordial germ cells (PGCs) are the precursors to the gametes and have the unique ability to retain full developmental potential. However, the mechanism(s) and gene-network(s) necessary for their proper specification and development are poorly understood. This is due, in part, to the challenges that must be overcome in order to identify and isolate PGCs during critical stages of development. Two distinct mechanisms have been characterized to specify the germ cell lineage in vertebrates: induction and inheritance. Regardless of mechanism, there are common developmental features shared among all vertebrates in forming the germ cell lineage. Xenopus offers several advantages for understanding the molecular mechanisms necessary to establish the germ line. Here, we provide detailed methods for isolating live PGCs at different time points: 1) just after they have segregated from the endodermal lineage, and 2) while they are migrating towards the presumptive gonad. Isolation of PGCs at these critical developmental stages will allow for the investigation of the mechanism(s) and gene-network(s) necessary for their proper specification and development. Key words Xenopus, Germ line, Primordial germ cell (PGC), Germ plasm, Maternal-to-zygotic transition (MZT), Migration

1

Introduction PGCs maintain their totipotent potential even upon differentiation into highly specialized gametic cells. Retaining full potential is an essential characteristic of PGCs and is required for the survival of all sexually reproducing species. However, the mechanism(s) and gene-network(s) necessary for their proper specification and development are poorly understood. Defining the PGC specialized gene program that preserves this potential will provide mechanistic insights for the fields of reproductive and stem cell biology. To investigate the ability of PGCs to retain totipotency, methods are necessary to isolate these cells at different stages during development. Isolating PGCs from any organism presents various challenges including their tight association with somatic germ layers and their relatively low numbers compared to somatic cells.

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Two distinct mechanisms have been characterized to specify the germ cell lineage in vertebrates, namely induction and inheritance. Regardless of mechanisms, there is a commonality in the development of the vertebrate germ line including posttranscriptional gene regulation and protection from somatic differentiation [1]. Additionally, several essential genes for germ cell development including vasa, dazl, nanos, oct4, deadend, and tudor are conserved in vertebrates [1]. Therefore, it is useful to study different model systems to understand key concepts in germ line development. Xenopus offers several advantages for understanding the molecular mechanisms necessary to establish the germ line: rapid external development, large embryos containing abundant quantities of RNA and protein, well-characterized developmental stages, ease of creating loss-of-function mutants, and ability to monitor phenotypic changes with in vivo imaging [2]. Germ line specification in Xenopus is established via inheritance of germ plasm, a subcellular matrix containing maternally derived RNAs and proteins. Germ plasm has been shown to be both required and sufficient for germ cell specification [3]. Therefore, cells that inherit germ plasm during development are destined to become PGCs, which ultimately form the gametes. During early oogenesis, germ plasm components are assembled into a distinct, mitochondrial-rich subcellular structure termed the Balbiani body (BB) (also known as the mitochondrial cloud). Germ plasm is then moved to the vegetal pole as oogenesis progresses [4–7]. Somatic determinants are also localized to the vegetal pole [8]. During cleavage, the germ plasm is asymmetrically inherited by one daughter cell termed the presumptive PGC (pPGC). Segregation of the germ line occurs at gastrulation when the germ plasm moves to a perinuclear location and subsequent divisions result in both daughter cells receiving germ plasm, now termed PGCs. After segregation from the endoderm, PGCs initiate their zygotic transcription program. Beginning in the tailbud stage and continuing throughout early tadpole stages, PGCs undergo migration steps and eventually colonize the presumptive gonad. Here, we provide detailed methods for isolating live PGCs: (1) when they segregate from the endoderm lineage, and (2) while they undergo migration towards the presumptive gonad. Isolation of PGCs at these critical developmental stages will allow for the investigation of the mechanism(s) and gene-network(s) necessary for their proper specification and development. With the advent of single cell RNA-sequencing analysis, these procedures open up a new frontier in understanding gene networks in the germ line [9, 10].

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Materials

2.1 Isolating PGCs from St. 12–16 X. laevis Embryos

1. Ca2þ, Mg2þ free medium (CMFM): 50.3 mM NaCl, 0.7 mM KCl, 9.2 mM Na2HPO4, 0.9 mM KH2PO4, 2.4 mM NaHCO3, 1.0 mM EDTA, pH 7.3. 2. 2 % agarose in CMFM. 3. 0.1 Marc’s Modified Ringers (0.1 MMR) Buffer: 0.1 M NaCl, 2 mM KCl, 1 mM MgSO4, 2 mM CaCl2, 5 mM HEPES (pH 7.8). 4. Cysteine: 2 % w/v Cysteine Y in 0.1 MMR (pH 8.0). 5. Saturated solution of DiOC6 (Invitrogen) in DMSO (solution can be used for up to 6 months stored at 4ºC). Bring solution to room temperature before use. 6. Stock DiOC6 solution 2:1000 (v/v) made from the saturated solution in 0.1 MMR buffer (make fresh before use). 7. Working DiOC6 solution (5:1000) made from the stock solution in 0.1 MMR buffer (make fresh before use). 8. 10 and 6 cm clean glass petri dishes (Corning Life Sciences). 9. Dumont #5 forceps (Fine Science Tools), sharpened. 10. Plastic, sterile transfer pipettes (Sarstedt). 11. 10 μL micropipette and sterile pipette tips. 12. Nuclease-free, sterile 1.5 mL snap-cap tubes. 13. Fluorescent Stereo Microscope (Magnifications 1.6 PF, 16-100 zoom, 460-490 nm excitation filter).

2.2 Isolating PGCs from St. 24-42 X. laevis Embryos

1. Ca2þ, Mg2þ free medium (CMFM): 50.3 mM NaCl, 0.7 mM KCl, 9.2 mM Na2HPO4, 0.9 mM KH2PO4, 2.4 mM NaHCO3, 1.0 mM EDTA, pH 7.3. 2. 2 % agarose in CMFM medium. 3. 0.1 Marc’s Modified Ringers (0.1 MMR) buffer: 0.1 M NaCl, 2 mM KCl, 1 mM MgSO4, 2 mM CaCl2, 5 mM HEPES (pH 7.8). 4. Buffered MS222: 1.5 g MS222 (Sigma), 1.5 g sodium bicarbonate, 1 L 0.1 MMR buffer. 5. Cysteine: 2 % w/v Cysteine Y in 0.1 MMR (pH 8.0). 6. 4 % Ficoll (Sigma) in 0.1 MMR. 7. pCS2þ -Venus-DS 30 UTR plasmid [11]. 8. mMESSAGE MACHINE Transcription kit for SP6 or T7 (Life Technologies). 9. mRNA transcribed from the PCS2 þ Venus-DS 30 UTR plasmid using mMESSAGE MACHINE (SP6 or T7 depending on plasmid).

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10. Micro-injector, Nanoject II (Drummond Science Company). 11. 10 and 6 cm glass petri dishes (Corning Life Sciences). 12. Sterile scalpel blades. 13. Plastic, sterile transfer pipettes (Sarstedt). 14. 10 μL micropipette and sterile pipette tips. 15. Nuclease-free, sterile 1.5 mL snap-cap tubes.

3

Methods

3.1 Isolating PGCs from St. 12–16 X. laevis Embryos

Xenopus PGCs initiate their specialized zygotic transcription program after they segregate from the endodermal lineage at gastrulation [12]. Therefore, it is predicted that the PGC transcriptome will diverge from endoderm during gastrulation and early neurula stages (St. 12–16). To investigate the molecular mechanisms necessary to understand this critical juncture in germ line specification, PGCs must first be isolated at these developmental stages. Here, we describe detailed methods, modified from our previous work [13], to isolate PGCs from St. 12–16 embryos. The PGC isolation procedure takes advantage of the unusually high concentration of mitochondria in germ plasm. We use the lipophilic fluorescent dye DiOC6 (3,30 -dihexyloxacarbocyanine), which accumulates only in live mitochondria, to label PGCs in embryos. Using the method described below, we have recently performed RNA-seq analysis on PGCs and stage-paired endoderm cells just after they initiate their zygotic program (St. 12–14). As predicted, our PGC population is significantly enriched with transcripts specific to the germ cell lineage including vasa, dazl, deadend, and nanos, compared to matched endoderm cells (unpublished RNA-seq data sets will be uploaded to NCBI after complete analysis). These data provide compelling evidence that this method is useful for identifying and isolating PGCs at gastrula and neurula stages. Alternatively, PGCs can be identified using the Dria-line of transgenic Xenopus female frogs, harboring enhanced green fluorescent protein fused to the mitochondrial transport signal [14]. The Dria-line is available from the National Xenopus Resource at the Marine Biological Laboratory in Woods Hole, MA. * DiOC6 is light sensitive: all solutions and stained embryos must be protected from light (see Note 1). 1. Follow the Cold Spring Harbor Laboratory Manual in vitro fertilization procedure for producing embryos [15]. 2. Dejelly embryos [15] at 2-cell stage and rinse ten times with 0.1 MMR. 3. When embryos reach the 4-cell stage, submerge the embryos in the DiOC6 working solution for 30 min (see Note 1).

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4. Wash embryos ten times, 3–5 min each wash, in 0.1 MMR (see Note 2). 5. Stain several waves (see Note 1) of embryos as described in steps 3 and 4. Then incubate stained embryos at different temperatures (such as 17–18  C and room temperature) to allow for a range of stages to be available the next morning. 6. When the embryos reach stage 10–11, screen them for stained PGCs in the yolk plug. If the embryo is well stained, you should easily detect 1–5 PGCs in the yolk plug when viewed under a fluorescent stereomicroscope (Fig. 1A, B). 7. Collect only the St. 10-11 embryos where you can clearly detect 1–5 PGCs in the yolk plug (Fig. 1A, B) and put them in a new petri dish with 0.1 MMR. Allow the embryos to develop until they reach the desired stage (St. 12–16). 8. Once the sorted embryos are at the desired stage, under a bright field microscope, use forceps to remove the vitelline membrane from each embryo (see Note 3). Act quickly to reduce the exposure of embryos to light. 9. Under a bright field microscope, use forceps to expose the endodermal core by cutting along the neural tube (Fig. 1C). Next, use forceps to carefully dissect out the endodermal core (Fig. 1D; see Note 3). Again, act quickly to reduce the light exposure. 10. To dissociate the endodermal core into single cells, use a pasture pipette to transfer the endodermal core into a 10 cm, agarose-coated petri dish containing CMFM (see Note 4). Incubate endodermal cores in CMFM for 10 min at room temperature. 11. Using a hair-loop, carefully brush over one of the endodermal cores. This should cause the cells to separate. If the cells do not separate, allow the cores to incubate longer in CMFM. 12. Once the blastomeres in the endodermal core are able to separate, carefully wave the hair-loop through the cells so that all the individual cells contained in the core are visible. Using a fluorescent stereomicroscope, identify the PGCs. Although there will be several blastomeres lightly stained, the PGCs are identified by having the most intense, sub-cellular signal (Fig. 1E) (see Note 5). 13. Using a micropipette, carefully collect the PGCs (you will collect endoderm cells along with PGCs at this step) and move them to a clean area in the dish (Fig. 1F). Next, using a gentle stream of CMFM media from the pipette, move somatic cells away from the PGCs (Fig. 1G). PGCs are isolated in progressive steps, diluting away the contaminating endoderm cells. Sub-cellular localization of germ plasm in PGCs is easily

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Fig. 1 DiOC6-stained and dissociated PGCs in X. laevis embryos. (A) Identification of successfully stained DiOC6 PGCs at St. 111/2. A, bright field; A’, fluorescent images. (B) Representative fluorescent image of the yolk plug of a DiOC6-stained St. 111/2 embryo; note sub-cellular localization of germ plasm (green). Inset, whole embryo, zoomed region identified. (C) Representative DiOC6-stained St. 14 embryo with an incision along the neural tube, exposing the endodermal core. (D) Isolated endodermal core. (E) Dissociated endodermal core cells. E, bright field; E’, fluorescent images. Arrows indicate PGCs (F) Diluted PGCs (arrows) and matched endoderm cells. F, bright field; F’, fluorescent images. (G) Isolated PGC. G, bright field; G’, fluorescent images. (H) Representative image of isolated DiOC6-stained St. 14 embryo embedded in 1 % agarose and imaged using a confocal microscope. H, bright field; H’, merged; H’’, fluorescent images. Note specific staining of perinuclear-localized germ plasm. Arrow indicates nucleus

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detected using a fluorescent stereomicroscope. A confocal microscope may be used to confirm perinuclear location of germ plasm in DiOC6-stained PGCs isolated from St. 12–16 embryos (Fig. 1H). 14. Once the PGCs are isolated from surrounding cells, collect the PGCs (in as little volume as possible) using a micropipette and transfer them into a nuclease-free, sterile 1.5 mL tube containing either RNA or protein extraction buffer (of your choice) on ice. Be sure to use a new, sterile pipette tip after PGCs have been transferred to lysis buffer. 15. Repeat steps 11–14 until the PGCs in each endodermal core have been isolated. 16. Begin at step 8 with the next wave of embryos, once they have reached the desired stage. 17. After all the PGCs have been collected (in the same tube), flashfreeze samples on dry ice then store them at 80  C until you are ready to extract RNA or protein (see Note 6). 3.2 Isolating PGCs from St. 24-42 X. laevis Embryos

Similar to most vertebrates, PGCs in the developing Xenopus embryo undergo a series of cellular migration steps to reach the presumptive gonad. PGC migration begins in the tailbud stage, stage 24 [16–18], and continues throughout early tadpole stages. To study the molecular mechanisms and active gene networks necessary for this critical developmental step, we have designed a method to isolate PGCs as they undergo migration. There are roughly 16–24 PGCs found in St. 24-32 embryos, and 32–48 PGCs in St. 33-42 embryos [17, 19]. Although PGCs will retain DiOC6 fluorescent dye through stage 42 without any adverse effects on migration, the fluorescent signal is more faint and difficult to distinguish at later stages. Here, we describe a method to identify PGCs during migratory stages (St. 24-42) that results in lower background staining and better contrast between PGCs and somatic cells; thereby allowing for successful isolation of PGCs. The method described below involves targeting a reporter construct such as Venus to the germ cells using the 30 UTR of DEADSouth (DS). The DS 30 UTR is degraded following the maternal to zygotic transition (MZT) by miRNAs in somatic cells, but not in PGCs [20]. Alternatively, PGCs can be identified using the Drialine of transgenic Xenopus female frogs, harboring enhanced green fluorescent protein fused to the mitochondrial transport signal [14]. The Dria-line is available from the National Xenopus Resource at the Marine Biological Laboratory in Woods Hole, MA. 1. Follow the Cold Spring Harbor Laboratory Manual for in vitro fertilization procedure for producing embryos [15]. 2. Dejelly embryos [15] at 2-cell stage and rinse ten times with 0.1 MMR.

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Fig. 2 Venus-labeled PGCs in X. laevis embryos. (A) Live 16-32-cell embryo; note: germ plasm is detected as dark regions at vegetal pole (*). Typical injection sites are shown. (B) Cartoon of tailbud-stage embryo; location PGCs should be found is indicated in green; region removed for subsequent dissociation outlined in black panels. (C-E) Live tailbud stage embryos. Arrows indicate embryos with successful PGC staining. C, D, E, bright field; C’, D’, E’, fluorescent images

3. When the embryos reach the 16-32 cell stage, inject 50–100 embryos with Venus-DS 30 UTR transcript into two germ plasm bearing blastomeres (400 pg total, 200 pg/blastomere) per embryo (Fig. 2A; see Note 7). Perform injections in 4 % Ficoll in 0.1 MMR. 4. Inject several waves of embryos as described in step 3. Then incubate injected embryos at different temperatures (such as 17–18  C and room temperature) to allow for a range of stages to be available the next morning. 5. When the embryos reach stage 10–11, screen them for stained PGCs in the yolk plug as described in Subheading 3.1, step 6 (Fig. 1A). 6. Collect only the St. 10-11 embryos where you can detect staining in the yolk plug (see Note 8) and put them in a new petri dish with 0.1 MMR. Allow the embryos to develop until they reach the desired stage (St. 24-42). 7. Once the sorted embryos are at the desired stage, place the embryos in buffered MS222 for ~5 min to slow their movement. Using a fluorescent stereomicroscope, identify the embryos bearing well-stained PGCs (Fig. 2C-E; see Note 9). 8. Collect only the embryos where you can clearly detect PGCs and transfer them to a new petri dish with 0.1 MMR. Of the collected embryos, select 8–12 and transfer them to a petri dish containing buffered MS222 to inhibit movement, and put the others back at 17  C. 9. Using a sterile scalpel blade cut the region containing PGCs (outlined in Fig. 2B). Wash the cut regions in 0.1 MMR then

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transfer them into a 10 cm, agarose-coated petri dish containing CMFM (see Note 4). Incubate cut regions in CMFM for 10 min at room temperature. 10. To isolate PGCs, process the cut regions as described for the endodermal core in Subheading 3.1, steps 11–15. 11. Begin at step 7 with the next wave of embryos. 12. Once all the PGCs have been collected (in the same tube), flash-freeze samples on dry ice then store them at 80  C until you are ready to extract RNA or protein.

4

Notes 1. Waves refer to a series of embryo clutches from a single mother that are fertilized at different time points. Stain no more than 100 embryos at a time. Set up a petri dish for DiOC6 staining by covering the lid of a 6 cm glass petri dish in aluminum foil and put 10 mL of 0.1 MMR in the bottom then add 50 μL of the stock DiOC6 solution and pipette up and down to mix (this is the DiOC6 working solution). When embryos are at the 4cell stage, transfer ~50–100 of them into the DiOC6 working solution and cover with a foil-covered lid for 30 min. *All DiOC6 solutions and DiOC6-stained embryos are protected from light using foil-covered lids as described here. 2. To wash embryos, set up a row of five 10 cm clean glass petri dishes (labeled wash 1–5) followed by five 6 cm clean glass dishes (labeled wash 6–10) each containing fresh 0.1 MMR. Transfer the embryos from the working DiOC6 solution to the first wash dish. After 3–5 min, transfer the embryos to wash #2. Repeat this until all ten washes are completed. 3. To reduce exposure to light, we suggest removing the vitelline membranes from no more than 8–12 embryos at a time. Dissect the endodermal core from these embryos one at a time, keeping the others in a 6 cm glass petri dish covered with foil at 17–18  C. 4. Be sure to leave adequate space around each endodermal core to allow for easier pick-up of individual PGCs. 5. IMPORTANT: Do not expect this protocol to work for every embryo, even after sorting. If there is not ~3–8 PGCs per endodermal core easily identified after dissociation, move on to the next embryo. 6. Each PGC contains roughly 400 pg of RNA. 7. Germ plasm is easily detected in the vegetal pole blastomeres from 8 to 32 cells as a dark shadow (Fig. 2a). This is an effective way of targeting transcripts to the germ line without staining.

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8. Since DS 30 UTR is in the process of being degraded in somatic cells at stage 10–11, there may be more nonspecific staining visible using this method compared to using DiOC6. Therefore, you should be less stringent when selecting for stained PGCs at this step. 9. If the embryo is well stained, you should detect roughly 16–48 PGCs (depending on the developmental stage) in the regions outlined in Fig. 2b, c. If ~16–48 PGCs are not detected, move on to the next embryo. References 1. Yang J, Aguero T, King ML (2015) The xenopus maternal-to-zygotic transition from the perspective of the germline. Curr Top Dev Biol 113:271–303 2. King ML (2014) Germ cell specification in xenopus. In: Kloc M, Kubiak JZ (eds) Xenopus development. Wiley, Hoboken, NJ, pp 75–100 3. Tada H, Mochii M, Orii H, Watanabe K (2012) Ectopic formation of primordial germ cells by transplantation of the germ plasm: direct evidence for germ cell determinant in xenopus. Dev Biol 371:86–93 4. Heasman J, Wylie CC, Hausen P, Smith JC (1984) Fates and states of determination of single vegetal pole blastomeres of X. laevis. Cell 37:185–194 5. Forristall C, Pondel M, Chen L, King ML (1995) Patterns of localization and cytoskeletal association of two vegetally localized rnas, vg1 and xcat-2. Development 121:201–208 6. Kloc M, Etkin LD (1995) Two distinct pathways for the localization of rnas at the vegetal cortex in xenopus oocytes. Development 121:287–297 7. Zhang J, Houston DW, King ML, Payne C, Wylie C, Heasman J (1998) The role of maternal vegt in establishing the primary germ layers in xenopus embryos. Cell 94:515–524 8. King ML, Messitt TJ, Mowry KL (2005) Putting RNAs in the right place at the right time: RNA localization in the frog oocyte. Biol Cell 97:19–33 9. Tang F, Barbacioru C, Bao S, Lee C, Nordman E, Wang X, Lao K, Surani MA (2010) Tracing the derivation of embryonic stem cells from the inner cell mass by single-cell rna-seq analysis. Cell Stem Cell 6:468–478 10. Kolodziejczyk AA, Kim JK, Svensson V, Marioni JC, Teichmann SA (2015) The technology and biology of single-cell RNA sequencing. Mol Cell 58:610–620 11. Kataoka K, Yamaguchi T, Orii H, Tazaki A, Watanabe K, Mochii M (2006) Visualization of the xenopus primordial germ cells using a

green fluorescent protein controlled by cis elements of the 30 untranslated region of the deadsouth gene. Mech Dev 123:746–760 12. Venkatarama T, Lai F, Luo X, Zhou Y, Newman K, King ML (2010) Repression of zygotic gene expression in the Xenopus germline. Development 137:651–660 13. Venkataraman T, Dancausse E, King ML (2004) Pcr-based cloning and differential screening of rnas from xenopus primordial germ cells: cloning uniquely expressed rnas from rare cells. Methods Mol Biol 254:67–78 14. Taguchi A, Takii M, Motoishi M, Orii H, Mochii M, Watanabe K (2012) Analysis of localization and reorganization of germ plasm in xenopus transgenic line with fluorescence-labeled mitochondria. Dev Growth Differ 54:767–776 15. Sive HL, Grainger RM, Harland RM (2000) Early development of xenopus laevis: a laboratory manual. CSHL Press, Cold Spring Harbor, NY 16. Kamimura M, Ikenishi K, Kotani M, Matsuno T (1976) Observations on the migration and proliferation of gonocytes in xenopus laevis. J Embryol Exp Morphol 36:197–207 17. Kamimura M, Kotani M, Yamagata K (1980) The migration of presumptive primordial germ cells through the endodermal cell mass in xenopus laevis: a light and electron microscopic study. J Embryol Exp Morphol 59:1–17 18. Nishiumi F, Komiya T, Ikenishi K (2005) The mode and molecular mechanisms of the migration of presumptive pgc in the endoderm cell mass of xenopus embryos. Dev Growth Differ 47:37–48 19. Whitington PM, Dixon KE (1975) Quantitative studies of germ plasm and germ cells during early embryogenesis of xenopus laevis. J Embryol Exp Morphol 33:57–74 20. Yamaguchi T, Kataoka K, Watanabe K, Orii H (2014) Restriction of the xenopus deadsouth mrna to the primordial germ cells is ensured by multiple mechanisms. Mech Dev 131:15–23

Chapter 10 Single-Cell Lineage Analysis of Oogenesis in Mice Lei Lei and Allan C. Spradling Abstract Lineage analysis is widely used because it provides a very powerful tool for characterizing the developmental behavior of the cells in vivo. In this chapter, we describe a particularly informative variant of lineage analysis that we term “single-cell lineage analysis”. As in traditional lineage analysis, the method employs a Tamoxifen (Tmx)-inducible CAGCreER mouse line, which is crossed to an R26R reporter line that can be activated by Cre-mediated DNA recombination. However, instead of driving CreER at a high level within a subset of cells defined by a particular promoter, CreER is driven with a generic promoter that is active in essentially all cells throughout the lifespan of the mouse. Specificity comes from using only a very low dose of Tmx so that just a few random, widely separated individual cells undergo recombination and become labeled. The growth and behavior of most such initially marked cells can subsequently be followed over time because each one forms a growing clone of marked cells that does not overlap with other clones due to their rarity. Following individual cell growth patterns provides much more information than can be derived from traditional lineage analysis, which relies on promoter specificity and uses high doses of Tmx that cannot resolve the behavior of single cells. We illustrate the value of single-cell lineage analysis using a recent study of fetal germ cell development and a recent search for female germ-line stem cells in adult mouse ovaries. Key words Tamoxifen, CreER, Germ-line stem cells, Ovary, Oocyte, Mouse

1

Introduction Inducible site-specific DNA recombination using CreER-loxP has been used widely to study gene function and for lineage analysis since the first CreER mouse line was established in 1995 [1]. Today, the most commonly employed variant is CreERT, in which a synthetic estrogen receptor (ER) has been fused to the Cre recombinase. The CreER recombinase remains inactive under normal physiological condition, but becomes functional in the presence of 4-hydroxytamoxifenTamoxifen (Tmx) (Tmx), which acts as an ER ligand (Fig. 1a). When CreER is driven by a specific promoter, Cre-mediated DNA recombination is controlled spatially by the promoter’s specificity, and temporally by the timing of Tmx injection ([2], Fig. 1b). However, promoters integrated into

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Fig. 1 Comparison between traditional cell-type specific lineage analysis and single-cell lineage analysis. (a) Marking cells heritably using Tmx-inducible CreER. Upon Tmx injection, Tmx binds to the inactive Cre in the cytoplasm. Activated Cre then relocates into the nucleus to enable YFP expression in the cell. (b) When CreER is driven by a “marker gene” promoter, only a specific type of cell in the tissue may be lineage labeled following Tmx injection. By contrast, single-cell lineage analysis uses a line in which CreER is driven by a generic promoter. All the cells in the mouse have an equal potential to be lineage labeled, and the dose of Tmx determines the number of lineage-labeled cells

cellular genomes often express in patterns that do not precisely mimic their endogenous behavior, complicating interpretation. Despite this potential limitation, CreER-based lineage analysis has been widely used to analyze progenitor cell differentiation and to search for stem cells in many murine organs [3–9]. An even more powerful and reliable approach of lineage analysis is to follow the development of individual cells without relying on the expression properties of a promoter construct [10]. The cytomegalovirus (CMV) early enhancer chicken beta actin (CAG) promoter is broadly and nonspecifically expressed in most or all mouse cells [11]. The suitability and reliability of the system is also

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tested in each experiment. When a proper dose of Tmx is identified, the distribution of lineage marked cells is first analyzed shortly after Tmx injection. It can be easily determined if all cells of interest are labeled randomly and at a suitably low frequency such that widely separated individual marked cells will be generated. Under these conditions, when tissue is analyzed at longer times after the single, initial Tmx injection, groups of labeled cells in the tissue will represent clones, i.e., the progeny of single cells at the time of injection. By contrast, when many cells are lineage marked following a high dose of Tmx injection at the initial time point, the clones of labeled cells they generate will run together, making it impossible to determine the growth kinetics and growth pattern of individual starting cells (Fig. 2).

Fig. 2 Effect of Tmx dose on the labeling behavior of mouse fetal germ cells. (A–B) Test of the lineage labeling efficiency 24 h after Tmx injection. (A, A0 ) In the mouse fetal gonad with a low dose of Tmx injection, only one lineage-labeled germ cell clone was found (arrow head). (B, B0 ) A higher dose of Tmx injection resulted in multiple adjacent lineage-labeled germ cell clones (arrow heads) in the mouse fetal gonad. (A00 ) A single lineage-labeled germ cell clone comprising six primary oocytes 14 days after Tmx injection. (C) Diagram of the single-cell lineage analysis used in searching for female germ-line stem cells (GSCs) in adult mouse ovaries

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Fig. 3 Experimental scheme used for Tmx-induced single-cell lineage analysis

In this chapter we describe in detail methods for single-cell lineage analysis using a mouse line that carries both CAGCreER and R26RYFP (Fig. 3). We have used this system to study several aspects of mouse oogenesis that remained elusive or controversial, including: (1) the differentiation of primordial germ cells (PGCs) into primary oocytes [12]; (2) the stability of the primary oocyte pool that arises during fetal development; and (3) to search for the presence of adult germ-line stem cells (GSCs), and the production of new primary oocytes in adult ovaries, a hypothetical process termed “neo-oogenesis” [13]. This same method of single-cell lineage analysis can be applied to characterize the cellular behavior of many if not most developing mouse tissues throughout development and in adulthood. Moreover, it provides a highly sensitive, unbiased, and reliable method for identifying stem cells in any tissue without the need to know beforehand anything about their number, location, or behavior. Most importantly, it is not necessary to first identify a “marker gene” for such stem cells to use the method. Consequently, we believe that the methods described here have a wide range of uses and that single-cell lineage analysis can be employed to resolve many important issues in mouse developmental and stem cell biology.

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Materials

2.1 Mouse Housing and Breeding

1. A mouse rack in a standard mouse facility with the capacity for about 30 type II cages. 1–5 mice will be housed in each cage. 2. Mouse lines: (a) CAGCreERþ/- mice: B6.Cg-Tg(CAG-cre/Esr1*)5Amc/ J (JAX® Mice stock number: 004682). (b) R26RYFP/YFP mice: B6.129X1-Gt(ROSA)26Sortm1(EYFP) Cos/J (JAX® Mice stock number: 006148)

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3. REDExtract-N-Amp™ Tissue PCR kit (Sigma-Aldrich). 4. Thermal cycler for genotyping PCR. 5. Standard equipment for agarose gel electrophoresis. 2.2 Lineage Label Induction

1. Tamoxifen (T5648, Sigma-Aldrich, see Note 1). 2. Ethyl alcohol (Absolute). 3. Corn oil. 4. 1.5 ml microcentrifuge tubes. 5. 50 ml conical centrifuge tubes. 6. 1 ml syringes (without needle). 7. 26 G needles. 8. Nutating shaker. 9. Freezers ( 20  C and

2.3 Whole-Mount Immunostaining

80  C).

1. 10 PBS stock solution (pH 7.4): dissolve 80 g NaCl, 2 g KCl, 14.4 g Na2HPO4, 2.4 g KH2PO4 in 1 L deionized (DI) water, adjust pH with 10 N HCl or 10 M NaOH. Store at room temperature. 2. 4 % paraformaldehyde (PFA, see Note 2): dilute 32 % PFA (EM grade) with 1 PBS. Store at 4  C for up to 4 weeks. Protect it from light during storage. 3. PBS-T2: 1 PBS, 0.1 % Tween 20, 0.5 % Triton X-100. Store at room temperature. 4. Antibody dilution buffer 1: PBS-T2 with 10 % serum from the animal that secondary antibody is derived from. In our case, we used donkey serum. Make fresh antibody dilution buffer each time before use. Serum should be aliquoted and stored at 20  C. 5. Primary antibodies: rabbit polyclonal to DDX4/MVH (1:400; Abcam), chicken polyclonal to GFP (1:1000; Aves Labs). Store according to the manufacturer’s instruction. 6. Secondary antibodies: Donkey anti-rabbit 568, donkey antichicken 488. Store according to the manufacturer’s instruction (see Note 3). 7. 40 , 6-diamidino-2-phenylindole (DAPI) stock solution: 5 mg/ ml in DI water. Store at 20  C. 8. VECTASHIELD mounting medium (Vector Labs). 9. Dissecting tools (surgical scissors, forceps, etc.). 10. Ice bucket, regular ice. 11. Kimwipes™ wipers. 12. Flat slide holders. 13. Slide boxes.

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Fig. 4 Using a slide box to construct a humid chamber for antibody staining

14. Microscope slides (Superfrost® plus). 15. Coverslips (18  18 mm). 16. Regular clear nail polish. 17. Stereomicroscope. 18. Confocal microscope. 2.4 Frozen Section Immunostaining

1. 30 % sucrose: 30 g sucrose, 100 ml 1 PBS, store at room temperature. 2. PBS-T: 1 PBS, 0.1 % Tween 20, store at room temperature. 3. Antibody dilution buffer 2: PBS-T, 10 % serum from the animal that secondary antibody is derived from. Make fresh each time before use. 4. O.C.T. (Tissue-Tek). 5. Peel-A-Way® Embedding Mold (Polysciences, Inc.). 6. Dry ice. 7. Staining jars. 8. Humid chamber (see Note 4, Fig. 4). 9. ImmEdge hydrophobic barrier pen (Vector Labs). 10. Coverslips (20  60 mm). 11. Cryostat. 12. Upright fluorescence microscope.

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Methods

3.1 Mouse Crossing for Lineage Labeling

1. To set up a breeding pair, one adult male (8 weeks or older) and one adult female (6 weeks or older) are housed in one cage. Breeding pairs for lineage analysis: (a) R26RYFP/YFP (male)  R26RYFP/YFP (female). Adult female progeny will be used for timed mating to CAGCreERþ/-; R26RYFP/YFP (male) and Tmx injection for lineage analysis of fetal germ cell differentiation.

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(b) CAGCreERþ/-; R26RYFP/YFP (male)  R26RYFP/YFP (female). Adult female progeny will be used for Tmx injection and lineage analysis of adult germ-line stem cells. 2. To set up timed mating, two adult female R26RYFP/YFP mice are put into a cage with one adult male CAGCreERþ/-; R26RYFP/YFP mouse in the afternoon. Vaginal plug will be checked every morning after setting up mating. Midday on the day vaginal plug appears is designated as E0.5 (embryonic day 0.5). Birth usually occurs between E19 and E20. The day of birth is designated postnatal day 0 (P0). 3.2 Tamoxifen Stock Preparation

Tamoxifen (Tmx) powder is dissolved in corn oil as the delivery vehicle for injections. Make 25 mg/ml Tmx stocks for long-term storage (see Note 5). 1. Weigh 500 mg Tmx and place it into a 50 ml conical centrifuge tube. 2. Add 1 ml ethyl alcohol (Absolute) to the tube with Tmx powder, shake the tube to suspend Tmx. 3. Add 19 ml corn oil to the tube with Tmx and ethyl alcohol. Cover the tube with foil to protect Tmx from light. 4. Place the tube on a nutating shaker for about 30 min to dissolve Tmx. Tmx stock solution should be clear without any white Tmx crumbs. 5. Aliquot Tmx stock solution into 1.5 ml microcentrifuge tubes with 100 μl per tube or 500 μl per tube (see Note 6). 6. Store Tmx stocks in a 20  C freezer for up to 6 months or a 80  C ultralow freezer for up to 24 months. Once a stock is thawed for injection, discard the remaining Tmx properly (see Note 1).

3.3 Tmx Injection and Lineage Analysis of Single PGC in the Fetal Ovary

To follow the differentiation process of individual PGCs in mouse fetal ovaries, we inject pregnant R26RYFP/YFP females that had been mated to CAGCreERþ/-; R26RYFP/YFP males with a single dose of Tmx intraperitoneally on embryonic day 10.5 (E10.5). The critical step for this experiment is to first identify a proper dose of Tmx for which only one PGC on average is lineage labeled in each gonad. The dose we identified is 0.25 mg Tmx per 40 g body weight (BW) (see Note 7). Control gonads need to be collected as well including: (1) Gonads of the fetus with the genotype of CAGCreER-/-; R26RYFP/YFP from above injection and (2) Gonads of the fetus with the genotype of CAGCreERþ/-; R26RYFP/YFP, but injected with same amount of vehicle (oil with 5 % ethyl alcohol) on E10.5. In control gonads, no YFP-expressing cells should be found.

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1. Thaw a tube of 100 μl Tmx stock at room temperature (see Note 8). Add 400 μl corn oil to make 500 μl of Tmx at the concentration of 0.5 mg/100 μl. 2. Weigh the pregnant females and calculate the volume of Tmx for injection. 3. Draw Tmx from the tube with a 1 ml syringe without needle. 4. Remove any air bubbles in the syringe and wipe clean the tip of the syringe. 5. Put a 26 G needle on the tip of the syringe and push the plunger slowly to further get rid of air in the needle. 6. Inject the mice with Tmx intraperitoneally. Take care not to injure fetus when injecting. 7. Mice that are injected at the same time can be housed in one cage. It is recommended that the mice injected with Tmx at different time points be caged separately (see Notes 9 and 10). 3.4 Collecting Fetal Ovaries

1. Euthanize the mice according to the instruction from institutional Animal Care and Use Committee (see Note 11). 2. After opening the abdomen, dissect two intact uteri with fetuses and place them in a 35-mm petri dish with cold PBS. 3. Cut open the uterus with a fine scissor from one end, release fetuses into cold PBS. 4. Transfer fetuses with a blunt-pointed forceps into a petri dish with fresh cold PBS. 5. Dissect fetal ovaries under stereomicroscope and place each pair of ovaries in a microcentrifuge tube with 4 % PFA. Keep the tubes on ice during dissection. 6. Collect fetal tails in clean microcentrifuge tubes for genotyping. Leave the tails on ice before storing them in a 20 freezer or digesting for genotyping.

3.5 Whole-Mount Immunostaining of Fetal Ovaries

1. After 2–5 h fixation (see Note 12) in PFA, wash fetal ovaries in PBS-T2 on a nutating shaker three times, for at least 20 min each time. Possible stop point: ovaries can be stored in PBS-T2 at 4  C for 2–3 days. 2. Remove as much PBS-T2 as possible (see Note 13). Dilute the primary antibodies in antibody dilution buffer 1. Incubate the ovaries with 100 μl primary antibodies at 4  C overnight. 3. On the next day, remove the primary antibodies from the tubes. Add 1 ml PBS-T2 to each tube and wash three times, for at least 30 min each time.

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4. Remove as much PBS-T2 as possible. Dilute the secondary antibodies in PBS-T2. Incubate the ovaries with 100 μl secondary antibodies at 4  C overnight. 5. On the next day, remove the secondary antibodies from the tubes. Add 1 ml PBS-T2 to each tube and wash three times, for at least 30 min each time. 6. Remove PBS-T2, add 200 μl DAPI (5 μg/ml in PBS) to each tube. Leave the tubes at room temperature for 30 min to stain nuclei. Cover the tubes with foil. 7. Remove DAPI, add 1 ml PBS-T2 to each tube, wash for 30 min. 8. Place a pair of ovaries on a microscope slide by pipetting a small amount of PBS-T2 with ovaries onto the slide. 9. Remove PBS-T2 on the slide with Kimwipes wiper carefully. Do not touch the ovaries with wiper. 10. Put 1–2 drops of mounting medium on the ovaries, and place a coverslip on the top. Do not push down on the coverslip. Wait for about 30 s to let the mounting medium evenly distribute under the coverslip. Get rid of the air bubbles by very gently pushing down on the coverslip near bubbles with a pipet tip. 11. Seal the coverslip with nail polish by applying a small amount of nail polish on each edge of the coverslip. Place sealed slides on a flat slide holder. Do not image the ovaries until the nail polish is totally dry. Stained ovaries can be stored at 4  C for 3–4 weeks. 12. YPF-positive clones in germ cells can be easily distinguished based on cell morphology (Fig. 2). By observing under lower magnification lenses, the number of lineage labeled germ cell clones in each fetal ovary can be counted. 3.6 Tmx Injection for Lineage Analysis of GSCs in Adult Mouse Ovaries

In regard to the presence of GSCs in adult mouse ovaries, two critical questions are: (1) whether there is rapid oocyte turnover; (2) whether newly generated primary oocytes can be observed. To clearly address these two questions, we used single-cell lineage analysis to label only a small number (about 30 out of 2000) of VASA-expressing oocytes that are randomly distributed within each adult ovary (CAGCreERþ/- ; R26RYFP/YFP). These 30 YFP-labeled oocytes are a small enough fraction of the whole germ cell population that two adjacent YFP-positive oocytes were never found in the ovary 24 h after Tmx injection. So any clustered YFP-positive oocytes (two or more YFP-expressing oocytes adjacent to each other) found at later time points after Tmx injection can be safely taken as evidence of germ cell proliferation, a prerequisite for new oocyte production from GSCs (neo-oogenesis). 30 YFP-labeled oocytes/ovary is also a large enough statistical sample to allow us to identify potentially rare stem cells, and to quantify and examine the dynamics of oocyte turnover in adult mouse ovaries. Control

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ovaries are: (1) CAGCreER-/-; R26RYFP/YFP mice injected with same amount of Tmx. (2) CAGCreERþ/-; R26RYFP/YFP mice injected with same amount of vehicle. In control ovaries, no YFPexpressing cells are expected to be found. 1. Thaw a tube Tmx of 500 μl aliquot. The concentration is 2.5 mg/100 μl. 2. Weigh the mice and inject them with Tmx intraperitoneally at 5 mg per 20 g body weight. 3. Ovaries of the injected mice were taken at 24 h after the Tmx injection to assess the efficiency of lineage labeling, and at 4 days, 2 weeks, 6 weeks and 16 weeks after the Tmx injection for clonal analysis and assessing oocyte turnover in adult mouse ovaries. 3.7 Collecting Adult Mouse Ovaries

1. Euthanize the mice. Open the abdomen with surgical tools. After removing the intestine, ovaries can be found embedded in a mass of fat at the end of uteri. Take the fat tissue together with an ovary, and place it in a 35-mm petri dish with cold PBS. 2. Dissect fat tissue and ovarian bursa under a stereomicroscope. 3. Fix a pair of ovaries in 500 μl cold 4 % PFA in a microcentrifuge tube at 4  C overnight. 4. On the next day, wash the ovaries with PBS three times on a nutating shaker, 10 min each time. 5. Remove PBS and add 1 ml 30 % sucrose (dissolved in PBS). Leave the ovaries in sucrose for about 5 h to overnight at 4  C. Possible stop point: ovaries can be stored in 30 % sucrose at 4  C for 2–3 days. 6. Fill an ice bucket with fine-granulated dry ice, about 2 in. deep. 7. Label the embedding molds. Fill each mold with O.C.T. to about 2/3 of the total volume. Avoid making air bubbles in the O.C.T. 8. Transfer an ovary into a mold with O.C.T. with a fine forceps. Place the ovary at the center of the bottom of the mold (see Note 14). 9. Place the mold with an ovary on dry ice. Make sure the mold is level. O.C.T. will turn into a white block. 10. Store the molds with ovaries in a 80  C ultralow freezer. Ovaries can be stored for 1–2 years. Tissues are ready for sectioning as soon as they are frozen.

3.8 Frozen Sectioning Adult Mouse Ovaries

1. Before sectioning, equilibrate the molds containing ovaries in a cryostat chamber for at least 15 min. 2. Remove the mold, and section the block with its ovary. Collect ten consecutive sections on each slide. Section through the whole ovary and collect all sections.

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3. Dry the sections at room temperature for at least 30 min before storing the sections in a 80  C freezer or else start staining (see Note 15). 3.9 Immunohistological Staining of Ovarian Frozen Sections

1. Post-fix sections in cold 4 % PFA for 5 min (see Note 16). 2. Wash sections with PBS-T in a staining jar three times, 5 min each time. 3. Dilute the primary antibodies in antibody dilution buffer 2. 4. Draw a circle around the sections using an IHB pen. Do not let the sections dry. 5. Add the primary antibodies inside the circled area containing sections. Incubate the sections with primary antibodies in a humid chamber at 4  C overnight. 6. The following day, wash slides with PBS-T in a staining jar three times, for at least 5 min each time. 7. Dilute the secondary antibodies in PBS-T, incubate the sections with secondary antibodies in a humid chamber for 1 h at room temperature. 8. Wash slides with PBS-T in a staining jar three times, for at least 5 min each time. 9. Add DAPI (5 μg/ml in PBS) on the sections and stain the nuclei for 5 min. 10. Rinse the slides briefly in PBS-T, add 1–2 drops of mounting medium on each slide. 11. Place a coverslip (20  60 mm) and seal the coverslip by applying a small amount of nail polish on each edge of the coverslip. 12. By using lower magnification lenses, examine each section to find YFP-positive, VASA-positive oocytes. Besides recording whether there are two or more adjacent YFP-positive oocytes on each section, for each YFP-positive oocyte, examine two adjacent sections (the one before and the one after), to see whether there are YFP-positive oocytes in the same region.

4

Notes 1. Tamoxifen (Tmx) is a health hazard. Avoid inhaling or touching with bare hands when handling Tmx. Dispose unused Tmx and the containers used for preparing Tmx properly according the institutional guidelines for biohazard waste disposal.

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2. PFA is carcinogenic. Use a fume hood when handling PFA. Dispose used, old PFA and the containers used for preparing PFA properly according the institutional guidelines for biohazard waste disposal. 3. When incubating tissues with multiple primary antibodies simultaneously, make sure the primary antibodies are derived from different species of animals, and the secondary antibodies are generated in the same species but conjugated with different fluorophores that can be distinguished based on their wavelength. 4. A humid chamber for the immunohistological staining on sections can be made from a regular slide box (100 slides) as shown in Fig. 4. When incubating antibodies, make sure to put the slide box on a level surface so antibody solution will not accumulate on the lower side of slide, which may cause the sections on the higher side of slide to dry up during an overnight antibody incubation. Dried sections will exhibit nonspecific staining. 5. To minimize variation between individual injections, make a relative large batch of Tmx stock according to the particular experimental need, and store it at 80  C. 6. Use 1.5 ml microcentrifuge tubes instead of 0.5 ml small tubes for Tmx stocks. When Tmx is thawed for injection, draw Tmx directly from the tube with a 1 ml syringe without needle. Oil is very viscous and cannot be drawn quickly into a syringe with a needle. 7. To identify a proper dose of Tmx to label single PGCs in the fetal gonad, we injected mice with graded doses of Tmx. The doses tested were: 1 mg/40 g BW, 0.5 mg/40 g BW, 0.25 mg/ 40 g BW, 0.125 mg/40 g BW. Fetal gonads were collected 24 h after Tmx injection to assess the number of lineage-labeled (YFP-positive) PGCs in each gonad by immunostaining. Since PGCs are also actively dividing from E10.5 to E11.5, we looked for the number of YFP-positive germ cell clones (germ cell clusters with YFP expression) in each gonad. Individual labeled germ cell clones can be easily identified (Fig. 2). The key time point for assessing labeling efficiency is 24 h after Tmx injection. 8. When thawing a Tmx stock for injection, white Tmx precipitation usually appears. After adding oil to dilute the Tmx stock to the working concentration, warm the tube in your hands and shake it several times until all precipitated Tmx disappears. 9. It is recommended that mice injected with Tmx at different time points be caged separately, since any Tmx left on the fur could be consumed by other mice and induce unwanted lineage label. 10. The observed rate of miscarriage induced by Tmx injection was calculated as 28 % (n ¼ 353 injections). Because the injection

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was done on E10.5, and the pregnancy of female mice is based on the appearance of vaginal plug, the actual percentage of Tmx-induced miscarriage is even lower, since it is quite normal that a small proportion of female mice with vaginal plug are not pregnant. 11. It takes about 15 min to collect fetal ovaries from one pregnant mouse, and since tissues may start necrosis shortly following an animal’s death, it is recommended to euthanize mice one at a time, and fix their fetal ovaries in as short a time frame as possible. 12. The time of fixation is determined by the size of a tissue. For fetal gonads/ovaries (E11.5–E14.5) and ovaries (E17.5–P4), 2–5 h fixation is enough. Leaving the ovaries in 4 % PFA at 4  C overnight will not cause significant negative effects on staining, although this is not recommended. 13. Fetal ovaries are very small and relatively opaque following fixation and washing with PBS-T2. When changing buffer, make sure the ovaries do not stick to the tube wall before removing the wash buffer from the tube. Spinning tubes briefly with a mini centrifuge helps to settle the ovaries down. 14. When freezing ovaries in O.C.T., opaque liquid O.C.T. soon turns into a white block. To make future sectioning easier, place the ovary at the center of the bottom of the mold. After removing the mold before sectioning, the ovary can be easily identified as a pinkish or brownish tissue in the white O.C.T. block. In our experience, trimming the O.C.T. block with an ovary in the middle into an 8  8 mm block will be helpful in making consecutive sections. Another helpful tip is that after taking the O.C.T. blocks out from the 80  C freezer for sectioning, equilibrating the O.C.T. blocks in a cryostat chamber or a 20  C freezer for 30 min or longer will also help one make good consecutive sections. 15. Sections can be stored at 80  C for 1–2 weeks and still remain suitable for immunostaining. Staining of freshly sectioned tissue is recommended. 16. Post-fix the sections right after the sections are thawed. Do not let the sections dry.

Acknowledgement This work was supported by Howard Hughes Medical Institute. We thank Dr. Matthew Sieber, Dr. Ethan Greenblatt, and Jui-Ko Chang for comments on the chapter.

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References 1. Feil R, Brocard J, Mascrez B et al (1996) Ligand-activated site-specific recombination in mice. Proc Natl Acad Sci U S A 93:10887–10890. 2. Feil S, Valtcheva N, Feil R (2009) Inducible Cre mice. Gene Knockout Protoc 530:343–363. 3. Vasioukhin V, Degenstein L, Wise B et al (1999) The magical touch: genome targeting in epidermal stem cells induced by tamoxifen application to mouse skin. Proc Natl Acad Sci U S A 96:8551–8556. 4. Harfe BD, Scherz PJ, Nissim S et al (2004) Evidence for an expansion-based temporal Shh gradient in specifying vertebrate digit identifies. Cell 118:517–528. 5. Barker N, van Es JH, Kuipers J et al (2007) Identification of stem cells in small intestine and colon by marker gene Lgr5. Nature 449:1003–1007. 6. Nakagawa T, Nabeshima Y, Yoshida S et al (2007) Functional identification of the actual and potential stem cell compartments in mouse spermatogenesis. Dev Cell 12:195–206. 7. Lepper C, Conway SJ, Fan CM (2009) Adult satellite cells and embryonic muscle progenitors have distinct genetic requirements. Nature 460:627–631.

8. Okubo T, Clark C, Hogan BL (2009) Cell lineage mapping of taste bud cells and keratinocytes in the mouse tongue and soft palate. Stem Cells 27:442–450. 9. Zhou BO, Yue R, Murphy MM et al (2014) Leptin-receptor-expressing mesenchymal stromal cells represent the main source of bone formed by adult bone marrow. Cell Stem Cell 15:154–168. 10. Fox DT, Morris LX, Nystul T et al. (2009) Lineage analysis of stem cells. In: Girard L (ed) Stembook. Harvard Stem Cell Institute. http://www.stembook.org/node/542. 11. Hayashi S, McMahon AP (2002) Efficient recombination in diverse tissues by a tamoxifen-inducible form of Cre: a tool for temporally regulated gene activation/inactivation in the mouse. Dev Biol 244:305–318. 12. Lei L, Spradling AC (2013) Mouse primordial germ cells produce cysts that partially fragment prior to meiosis. Development 140:2075–2081. 13. Lei L, Spradling AC (2013) Female mice lack adult germline stem cells, but sustain oogenesis using stable primordial follicles. Proc Natl Acad Sci 110:8585–8590.

Chapter 11 Visualization and Lineage Tracing of Pax7+ Spermatogonial Stem Cells in the Mouse Gina M. Aloisio, Ileana Cuevas, Yuji Nakada, Christopher G. Pen˜a, and Diego H. Castrillon Abstract The precise identity of spermatogonial stem cells—the germline stem cell of the adult testis—remains a controversial topic. Technical limitations have included the lack of specific markers and methods for lineage tracing of Asingle spermatogonia and their subsets. Immunolocalization of proteins in tissue sections has been a standard tool for the in situ identification and visualization of rare cellular subsets. However, these studies are limited by the need for faithful and reliable protein markers to define these cell types, as well as the availability of specific antibodies to these markers. Here we describe the use of a monoclonal antibody to Pax7 as a means to detect spermatogonial stem cells (SSCs) both in tissue sections and in intact seminiferous tubules. Furthermore, we describe methods for lineage tracing as an alternative method to visualize Pax7+ spermatogonial stem cells and their progeny. Key words Pax7, Spermatogonial stem cell, Spermatogenesis, Lineage tracing, Immunostain

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Introduction Spermatogenesis is a complex, multistep process that serves to maintain male fertility throughout life. The functional unit of the mammalian testis, the seminiferous tubule, is a multilayered epithelium that matures from spermatogonial precursors on the basal layer to more advanced cell types that migrate toward the tubular lumen, where spermatozoa are released [1]. Meticulous histologic analyses have identified type A “single” (Asingle) spermatogonia as the ultimate precursors of the entire spermatogenic series [2, 3]. Asingle spermatogonia are believed to sometimes undergo asymmetric cell divisions in which one daughter retains stem identity and the other proceeds to differentiate through progressive mitotic divisions with incomplete cytokinesis to give rise to successive chains of 2, 4, 8, or 16 interconnected cells, which are collectively termed undifferentiated spermatogonia [4]. Such asymmetric cell divisions (Asingle!Asingle(stem)+Asingle(differentiating)) clearly imply heterogeneity

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among Asingle spermatogonia. However, until recently, markers with the potential to distinguish among Asingle subsets were unknown [5]. Formal proof that spermatogonia have the potential to function as stem cells was provided by transplantation studies where spermatogonia from a donor mouse can restore spermatogenesis in a recipient mouse that has undergone ablation of its germ cells via genetic means or cytotoxic drugs [6]. Although combining transplantation with flow sorting (i.e., based on cell surface markers) provided some information into the properties of spermatogonial stem cells (SSCs), the exact identity of the SSC in the mouse testis has remained elusive [7]. In the future, flow sorting based on spermatogonial subsets marked by expression of fluorescent protein reporters (i.e., through transgenic approaches or “knock-in” into specific loci by CRISPR) will undoubtedly prove increasingly useful [8]. However, the destruction of normal cellular relationships (i.e., the reduction of all spermatogonial chains to individual cells during mechanical/ enzymatic dissociation) and the artificial ablation of the germline in the host (which leads to a highly abnormal somatic microenvironment) will always represent significant limitations in the use of transplantation to identify and study SSCs [9–11]. Thus, methods to precisely identify Asingle subsets in situ and to conduct lineage tracing are needed. In three studies published in 2014, the Oatley, Hamra, and Castrillon laboratories provided direct evidence for heterogeneity among Asingle spermatogonia through analyses of diverse markers, challenging the dogma that all Asingle cells are equivalent, but in agreement with some prior studies suggesting that only a subset of Asingle spermatogonia function as stem cells in transplantation assays [12]. The three markers reported to distinguish among Asingle subsets in these 2014 studies were Erbb3, Id4, and Pax7. Abid et al. (2014) found that Erbb3 was expressed in only a small fraction of Asingle spermatogonia in rats, defining a novel population of Asingle spermatogonia with unique properties [13]. Chan et al. followed up on a previous study characterizing Id4 as a marker of Asingle spermatogonia, demonstrating that these Id4+ spermatogonia represented a subset of Asingle spermatogonia, and furthermore that these Id4+ cells functioned efficiently as stem cells in transplantation studies [8]. Aloisio et al. described a rare subset of Asingle spermatogonia defined by expression of Pax7, and showed by formal lineage-tracing studies using a Cre reporter knocked into the Pax7 locus that these rare cells function as bona fide stem cells in both steady-state spermatogenesis and during regeneration following chemotherapeutic ablation of germ cells [14]. Another protocol in this volume will address the use of Erbb3 as a spermatogonial marker in rats. Here, we describe three methods for visualizing Pax7+ spermatogonia and their descendants by (1) immunodetection in tissue sections, (2) immunofluorescence of intact seminiferous tubules, and (3) in vivo lineage tracing.

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Materials

2.1 Immunohistochemistry (IHC)

1. 10 % Neutral buffered formalin (Fisher) (see Note 1). 2. Super Frost Plus slides. 3. Xylene (see Note 2). 4. 100 % Ethanol. 5. 1 M Sodium citrate in deionized H20. Adjust pH to 6.0 with HCl and then sterilize through a 0.2 μm filter. 6. 30 % Hydrogen peroxide. 7. PBS (144 mg/L KH2PO4, 9000 mg/L NaCl, 795 mg/L Na2HPO4l7H2O) 8. 10 % BSA in PBS. 9. Hydrophobic barrier pen (Vector Labs). 10. Pax7 monoclonal antibody (Hybridoma Bank, bioconcentrate, see Note 3). 11. 20 TBST: 200 mL 1 M TRIS pH 7.5, 600 mL 5 M NaCl, and 20 mL Tween-20, then add deionized H20 to 1 L. 12. Horseradish peroxidase (HRP)-conjugated anti-mouse secondary detection reagent (Immpress). 13. 3,30 -Diaminobenzidine (DAB) chromagen (Dako) (see Note 4). 14. Hematoxylin 2 (Thermo Scientific), undiluted. 15. Cover slips, 24  60 mm.

2.2 Immunofluoresence (IF)

1. 10 % Neutral buffered formalin (see Note 1). 2. Super Frost Plus slides. 3. Xylene (see Note 2). 4. 100 % Ethanol. 5. 1 M Sodium citrate in deionized H2o. Adjust pH to 6.0 with HCl and then sterilize through a 0.2 μm filter. 6. 1 M Tris–glycine in Tris-buffered saline (TBS), pH 9.0. 7. 1 M NH4Cl in deionized H2O. 8. Blocker reagent: 1 % BSA in PBS. 9. 10 % Normal goat serum (NGS). 10. Hydrophobic barrier pen. 11. Mouse-on-mouse (MOM) block (Vector Laboratories, see Note 5). 12. Alexa Fluor (488, 565, or 647)-conjugated anti-mouse secondary antibody.

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13. 40 ,6-Diamidino-2-phenylindole (DAPI) working solution: 5 mg/mL DAPI stock (dissolved in DMSO) diluted to 1:10,000 in 1 PBS. Store at 4  C (see Note 6). 14. Vectashield mounting solution. 15. Cover slips, 24  60 mm. 2.3

Whole-Mount IF

1. 16 % Paraformaldehyde, in 10 mL ampules. 2. PBT: 1 PBS + 0.1 % Tween 20. 3. Methanol. 4. NP-40 detergent solution: 0.2 % NP-40 in 1 PBS. This can be stored at room temperature for up to 3 months. 5. 10 % BSA in PBS. 6. MOM block. 7. 40 ,6-Diamidino-2-phenylindole (DAPI) working solution: 5 mg/mL DAPI stock (dissolved in DMSO) diluted to 1:10,000 in 1 PBS. Store at 4  C (see Note 6). 8. Vectashield mounting solution. 9. Cover slips, 24  60 mm.

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

3.1 IHC on ParaffinEmbedded Tissues

1. Dissect out testis and puncture the tunica albuginea with a needle. Fix tissue in at least 5 volume of formalin, overnight (12–16 h) at 4  C. Paraffin-embed per standard histology laboratory methods. 2. Use microtome to cut 5 μm sections onto Super Frost plus slides and allow slides to dry completely. 3. Deparaffinize the sections in two changes of xylene for 10 min each in 250 mL tubs (see Note 7). 4. Rehydrate the sections in an ethanol series of 100 % ethanol (2  3 min), 95 % ethanol (2  3 min), and in deionized H2O for 3 min (in 250 mL tubs). 5. For antigen retrieval, place slides in gently parboiling 600 mL of 10 mM sodium citrate pH 6.0 in a 1 L beaker on a hot plate for 15 min. Then remove the beaker from heat and allow the solution to cool for 20 min. 6. To block endogenous peroxidase, place slides in 3 % H2O2 (diluted in deionized H2O) for 30 min (in 250 mL tubs). 7. Place slides in Blocker reagent for 15 min.

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8. Use a Kimwipe to carefully dry the slide around the tissue. Draw a circle around the tissue with a hydrophobic barrier pen (see Note 8). 9. Incubate the Pax7 antibody diluted 1:500 in Blocker reagent at 4  C overnight in a sealed wet chamber (to prevent evaporation). 10. Wash slides in TBST 4  5 min with gentle rocking. 11. Completely cover the entire tissue and incubate with the Immpress HRP-conjugated anti-mouse secondary (undiluted) for 30 min at room temperature in a wet chamber. 12. Wash slides in TBST 4  5 min with gentle rocking. 13. Add sufficient DAB solution to completely cover the tissue. It should take approximately 30 s for the signal to appear but this may need to be adjusted based on detection methods (Fig. 1) (see Note 9). 14. Rinse the slides in deionized H2O for 20 s to remove excess DAB. As each slide completes the DAB exposure, it can be placed in the deionized H2O until all slides have been completely developed. 15. To counterstain, place the slides in hematoxylin for 5 s with gentle agitation by moving the slide holder to ensure even counterstain. Avoid over-staining, which can obscure DAB signals. 16. Rinse slides in 2–3 changes of deionized H2O, until water ceases to turn purple. 17. Allow slides to dry completely. 18. Using a transfer pipette, add enough Permount (approximately 200 μL per tissue section) to completely cover the tissue when the cover slip is mounted. To reduce air bubbles between the cover slip and the slide, carefully place one of the long edges of the cover slip on the slide first, and then allow the cover slip to fall onto the slide. Any remaining air bubbles can be pushed out by using the edge of a pipette tip (see Note 10). 3.2 IF on ParaffinEmbedded Tissues

1. Perform fixation, embed, cut sections, deparaffinize, rehydrate, and perform antigen retrieval as in steps 1–5 in Subheading 3.1. 2. Block in 100 mM Tris–glycine in PBS for 10 min. 3. Use a Kimwipe to carefully dry the slide around the tissue, and then draw a circle around the tissue with a hydrophobic barrier pen (see Note 8). 4. Block in 100 mM NH4Cl in 2 % BSA (diluted from 10 % BSA) in PBS for 10 min. 5. Block 1 h in 5 % NGS + 2 % BSA in PBS.

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Fig. 1 Immunodetection of Pax7+ SSCs. (a) Pax7 IHC of adult skeletal muscle. Pax7+ satellite cells are located beneath the basal lamina, and make up approximately 5 % of adult skeletal muscle nuclei [33]. (b) Pax7+ germ cells in the PD3 testis are relatively abundant. The staining is nuclear, and these cells make up approximately 25 % of the germline at this time point [14]. Note the nonspecific staining of the interstitium, but lack of nonspecific staining within tubules. Asterisks denote nonspecific staining of Leydig cells. (c) A complete cross section of adult testis, stained for Pax7, and inset showing a single Pax7+ SSC within a tubule. Again, there is nonspecific staining within the interstitium (right panel), but not within the seminiferous tubules. Inset shows specific nuclear staining of Pax7 in basal spermatogonia. In formalin-fixed testes, retraction artifact due to dehydration of tubules leads to some separation of tubules in the ensuing sections (asterisk); however, this does not affect interpretation of immunostaining. (d) Pax7 IF on formalin-fixed paraffin-embedded tissues of PD7 testis. Some background interstitial staining is apparent with Pax7 IF (asterisk). Again, the tubule interiors are free of background staining. (e) Whole-mount IF of Pax7 and Foxo1 in the PD3 testis. Pax7 stains only a fraction of germ cells at this time point. In whole-mount immunofluorescence, the nonspecific staining of the basement membrane is again apparent. A second cell stains positive for Pax7 in the adjacent tubule, slightly out of focus. Scale bars ¼ 25 μm for all figures except 100 μm for (b) adult testis

6. Block in one drop MOM in 2.5 mL 2 % BSA in PBS for 1 h. 7. Add sufficient solution of Pax7 antibody diluted 1:200 in 5 % NGS + 2 % BSA in PBS to completely cover tissue section and incubate overnight at 4  C in a sealed wet chamber (to prevent evaporation).

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8. Wash slides in PBS for 3  10 min with gentle rocking. 9. Add Alexa-conjugated anti-mouse secondary diluted 1:300 in 5 % NGS + 2 % BSA in PBS. Incubate in wet chamber for 1 h at room temperature (see Note 9). 10. Wash slides in PBS for 3  10 min with gentle rocking. 11. Add DAPI working solution and incubate for 10 min. 12. Rinse off excess DAPI by dipping the slides in PBS for a few seconds. 13. Add one drop of Vectashield per slide and cover slip. 14. To seal the edges of the cover slip to the slide, apply nail polish (see Note 11). 3.3

Whole-Mount IF

1. Prepare 4 % PFA from 16 % PFA in ampules by diluting in PBS. 2. Place testis in PBS. 3. Using sharp needle-nosed tweezers, remove the tunica albuginea, and carefully separate the seminiferous tubules. 4. Transfer the tubules into an Eppendorf tube and fix overnight in 4 % PFA in PBS at 4  C. 5. Wash tubules in PBS 2  5 min. 6. Remove PBS and add 25 % MeOH in PBT (see Note 14). 7. Remove 25 % MeOH and add 50 % MeOH in PBT. Incubate for 5 min on ice with gentle rocking. 8. Remove 50 % MeOH and add 75 % MeOH in PBT. Incubate for 5 min on ice with gentle rocking. 9. Remove 75 % MeOH and add 100 % MeOH. Incubate for 5 min on ice with gentle rocking. 10. Wash in 100 % MeOH for a second time. Store at 20  C for at least 2 h (see Note 15). 11. Remove 100 % MeOH and add 75 % MeOH in PBT. Incubate for 5 min with gentle rocking. 12. Remove 75 % MeOH and add 50 % MeOH in PBT. Incubate for 5 min with gentle rocking. 13. Remove 50 % MeOH and add 25 % MeOH in PBT. Incubate for 5 min with gentle rocking. 14. Remove 25 % MeOH and add PBS. Incubate for 5 min with gentle rocking. 15. Incubate with 0.2 % NP-40 for 20 min (see Note 16). 16. Wash for 5 min in PBS. 17. Block 2 h in 2 % BSA in PBS with gentle rocking. 18. Block in one drop MOM in 2.5 mL 2 % BSA in PBS for 1 h with gentle rocking.

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19. Add Pax7 Ab at 1:100 in 2 % BSA in PBS and incubate overnight at 4  C on a rocker. Subsequent steps are at room temperature (see Note 17). 20. Wash in PBS 3  10 min with gentle rocking. 21. Add Alexa-conjugated anti-mouse secondary at 1:300 in 2 % BSA in PBS for 2 h. 22. Wash in PBS 3  10 min with gentle rocking. 23. Add DAPI solution and incubate for 10 min. 24. Rinse the tubules in PBS for a few seconds. 25. Arrange the tubules on a slide with the needle-nosed tweezers (see Note 18). 26. Add 1–3 drops of Vectashield to the slide to cover the tubules, and cover slip (see Note 19). 27. To seal the edges of the cover slip to the slide, apply nail polish (see Note 20). 3.4

Lineage Tracing

Here we describe lineage tracing methods for Pax7+ SSCs. A precise protocol is not provided, as methods relating to mouse breeding have been well described, but we include a conceptual overview important to a successful lineage tracing experiment. These concepts should also apply to other known or future markers of Asingle subsets such as Erbb3 and Id4 when appropriate genetic tools become available. Multiple Pax7-Cre lines have been generated, and all are freely available through Jackson Labs. These include Pax7tm1(cre)Mrc/J (B6;129-Pax7tm1(cre)Mrc/J, stock #010530) made by Mario Capecchi [15], and two tamoxifen-inducible lines, Pax7tm2.1(cre/ERT2)Fan/ J (B6;129-Pax7tm2.1(cre/ERT2)Fan/J, stock #012476) made by Chen-Ming Fan [16], and Pax7tm1(cre/ERT2)Gaka/J (B6.CgPax7tm1(cre/ERT2)Gaka/J, stock #017763) made by Gabrielle Kardon [17]. In all three lines Cre expression faithfully mirrors Pax7 expression in both the testis and skeletal muscle, with varying degrees of recombination efficiency as determined with diverse Cre tester lines ([14], unpublished). Pax7tm1(cre)Mrc/J was made by inserting an IRES-Cre cassette into the 30 UTR after exon 10. This does not interfere with normal function of Pax7, as mice can be maintained as a homozygous stock [15]. Pax7tm1(cre)Mrc/J is about 70 % efficient in Pax7+ spermatogonia at postnatal day 7 (unpublished data), with the tamoxifen-inducible lines being less efficient (Fig. 2). The structure of the Pax7tm2.1(cre/ERT2)Fan/J allele is a knock-in of CreERT2 followed by an IRES-DsREd inserted into exon 1, creating a hypomorphic allele. Therefore, these mice must be maintained as a heterozygous stock as Pax7 is an essential locus (90 % of homozygous offspring die before puberty) [16]. Although the Pax7tm2.1(cre/ERT2)Fan allele was designed for DSred

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Fig. 2 Lineage tracing of Pax7 descendants using Pax7-Cre lines (a) Number of labeled clones 6 weeks after tamoxifen administration in the two CreERT2 lines. Mice were administered 2 mg tamoxifen daily for 3 days at 6 weeks of age and aged for an additional 6 weeks. The Pax7tm2.1(cre/ERT2)Fan/J allele had fewer labeled clones than the Pax7tm2.1(cre/ERT2)Gaka/J, demonstrating that the latter has a higher efficiency in the germline. (b) Whole testis of a Pax7tm2.1(cre/ERT2)Fan/J; mT/mG mouse, showing large labeled clones on the green channel, and broader (but mutually exclusive) expression on the red channel. (c) Seminiferous tubule from a Pax7tm2.1 (cre/ERT2)Fan /J; Rosa26R mouse, showing dot-like localization of β-galactosidase. Note that at this magnification, chromatoid bodies cannot be clearly resolved, but much of the signal within the germ cell cytoplasm localizes to this structure. (d) Single seminiferous tubule from Pax7tm2.1(cre/ERT2)Fan/J; mT/mG mouse, as viewed on a confocal microscope. Signal is endogenous fluorescence. (e) Testis of a Pax7tm2.1(cre/ERT2)Fan/J; mT/mG mouse in cross section, stained with an eGFP-specific antibody. Individual labeled cells can be readily identified as to cell type based on cellular morphology and location. Endogenous reporter protein fluorescence can be quenched upon fixation; indirect immunofluorescence of fixed tissue sections is a viable alternative. (f) Seminiferous tubule from Pax7tm2.1(cre/ERT2)Fan/J; TdTomato mouse, showing strong endogenous nuclear signal. Tubules were isolated 6 weeks after tamoxifen administration except (f) 3 weeks after tamoxifen administration. Scale bars ¼ 500 μm (b) and 100 μm (c, d, e, f)

expression in Pax7+ cells, the signal is not detectable via immunostaining nor by FACs, likely due to inefficient ribosomal entry at the IRES ([16], and personal observations). Pax7tm1(cre/ERT2)Gaka/J was made by inserting an IRES-CreERT2 cassette 8 bp downstream of the endogenous stop codon. The IRES in this line appears to function with much greater efficiency, and gives expression of functional Pax7 protein. Thus, the allele can be homozygosed.

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In this protocol, we present some common criteria to evaluate linage tracing of SSCs. For studies of putative Asingle subsets, it is essential to first demonstrate that the marker of interest is indeed expressed exclusively within a subset of Asingle spermatogonia. Lineage tracing initiated with a Cre driver expressed in broad subsets of spermatogonia (e.g., all undifferentiated spermatogonia, or more restricted subsets such as Asingle-Aaligned4) would initiate labeling in large numbers of spermatogonia, only a few of which represent actual stem cells. Demonstration that labeling indeed begins within an Asingle population can be readily accomplished by confocal microscopy of intact tubules concurrently labeled for a marker of undifferentiated (Asingle!Aaligned16) spermatogonia such as Plzf, or Foxo1 [18, 19]. Examination of z-stacks permits confirmation that cells labeled by the lineage trace are indeed a subset of Asingle spermatogonia, and not part of larger chains (Apair!Aaligned16). This documentation of the initial patterns of labeling should be conducted within a week of induction of CreERT2 activity by tamoxifen, as longer intervals would make it increasingly difficult to distinguish the initially labeled cells from their descendants. Another criterion to evaluate lineage tracing with respect to the long-term stem potential of the cell type under investigation is the eventual full-linage maturation of clones (i.e., spermatogonia to motile spermatozoa) with long-term perdurance. This ensures that the cells under investigation are in fact stem cells, and not merely transit-amplifying cells whose progeny would become diluted out and eventually disappear. We previously demonstrated that Pax7-derived clones perdure for at least 16 weeks with no decrease in the number of clones. The duration of spermatogenesis in a mouse is estimated at 40 days. Therefore, the 16-week time point encompasses multiple rounds of spermatogenesis. Although the precise number is itself somewhat subjective, we believe that thorough lineage tracing studies of spermatogenesis require multiple time points up to the 16-week range [14, 20]. Several Cre reporter strains are attractive options for lineagetracing studies of SSCs. We have tested Rosa26 LacZ (FVB.129S4 (B6)-Gt(ROSA)26Sortm1Sor/J, stock #003309,[21]), TdTomato (B6.Cg-Gt(ROSA)26Sortm9(CAG-tdTomato)Hze/J, stock #007909, [22]), and a membrane-bound Tomato, membrane-bound eGFP line (mT/mG, Gt(ROSA)26Sortm4(ACTB-tdTomato-EGFP)Luo/J, stock #007576, [23]). These three lines have each proven useful in distinct ways. For unknown reasons, β-galactosidase localizes to the chromatoid body within germ cells. Thus, when a β-galactosidase Cre reporter is used, care must be taken to interpret staining patterns within germ cells (see [24] for description). When tubules are to be visualized following formalin fixation and embedding in paraffin, steps should be taken to minimize time in xylene, which dissolves the blue product [18, 21, 24]; otherwise, the blue signal is

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stable and permanent, facilitating histologic analyses. Since the detection of the β-galactosidase requires a special staining process, it is not ideal for colocalization with other antibodies. The second reporter, TdTomato, yields a very bright nuclear signal that lends itself well to FACS, and is preferred when IF will be performed simultaneously. Although the endogenous TdTomato signal is strong, it is subject to significant photobleaching during lengthy microscopy sessions. Therefore, we sometimes use a dsRed antibody (Living Colors) to detect TdTomato when doing colocalizations. This dsRed antibody does not cross-react with GFP. Finally, the mT/mG reporter clearly delineates cell borders, as the signal is localized to the cell membrane. The reporter is bright enough to detect endogenous fluorescence in whole-mount preparation of tubules, as well as in OCT sections. Many investigators have reported difficulty in the localization of fluorescent reporter signals in frozen sections of diverse mouse tissues [25–27]. On the one hand, fixation and processing clearly can abolish reporter protein fluorescence. A wide variety of conditions have been blamed for this quenching of signal, from the fixation method to the mounting media [25–30]. On the other hand, when unfixed tissues are cryosectioned, the fluorescent proteins readily diffuse from one labeled cell to non-labeled cells, obscuring specific signals and making visualization difficult if special steps are not taken [25]. To detect endogenous fluorescence, we recommend (based on [26]) cryosectioning unfixed tissues, allowing tissues to dry, and then fixing for 30 min in 4 % formalin, 7 % picric acid, and 20 % sucrose at 4  C. For the best resolution of labeled clones in tissue sections, we fix testes in 10 % neutral-buffered formalin, and perform IF as above using a GFP antibody (Aves) or a dsRed antibody that detects the TdTomato protein (Living Colors, 1:100). Whereas some antibodies can cross-react with diverse fluorescent protein derivatives, these antibodies are especially useful because of their specificity to GFP and DsRed, respectively [23] (see Note 21).

4

Notes 1. Formalin and its vapors are toxic, and care should be taken when fixing tissues. Waste should be disposed of separately. 2. Xylene can be reused for up to 40 slides. 3. The Pax7 antibody was donated to the Hybridoma Bank by Kawakami [31], and can be ordered in a few different forms. The “supernatant” is the least concentrated. The “concentrate” is well suited for detection of Pax7 in tissue sections and wholemount immunofluorescence. The “bioconcentrate” is nearly 10 as concentrated, and is the preparation of antibody used

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in these protocols. The epitope of this antibody has been identified [14] and therefore a corresponding blocking peptide could be used as a negative control. 4. DAB chromogen is a carcinogen and should not be allowed to come into contact with bare skin. 5. When using a mouse primary antibody, significant background can be due to the secondary anti-mouse antibody binding to both the primary antibody and endogenous mouse Ig antibodies. The MOM block reduces the binding of the secondary anti-mouse antibody to the endogenous mouse Igs. 6. DAPI is light sensitive and should be kept in the dark. Discard working solution after 2 weeks. 7. Once the slides have been submerged in xylene, do not let the slides to dry at any point in the experiment until the slides are stained with hematoxylin. 8. Pap pens can save on the amount of reagent (blocking solutions, antibody solutions, etc.) used by limiting the total volume needed to cover the slide. 200 μL is sufficient to cover most tissues. To keep the Pap pen functional for as long as possible, thoroughly dry the area where the barrier will be drawn, and store the pen with the tip down. 9. Other detection reagents can be used, but concentration of the Pax7 antibody and detection time will have to be determined by the user. 10. Overall, Pax7+ spermatogonia are exceedingly rare in adult testis. Before attempting to detect them in the adult, protocols should be optimized using neonatal testis, where Pax7+ germ cells are much more readily identified, as they make up about 25 % of germ cells [14]. In adult testis, Pax7+ germ cells are present in about one tubule in a complete cross section through the midportion of the testis. Thus, they are difficult to find, requiring meticulous scanning at high power. Nonetheless, several criteria aid in their identification. Pax7 staining is always nuclear in postnatal germ cells. Therefore, any non-nuclear staining should be ignored. Also, we sometimes observe nonspecific staining with the Hybridoma Bank Pax7 antibody in the tubular basement membrane and in the interstitium, particularly in Leydig cells. Both the basement membrane and Leydig cell staining represent nonspecific cross-reactivity to unknown antigens and should be ignored. However, within the seminiferous tubules, no nonspecific staining should be observed; that is, we do not observe nonspecific signals within more advanced spermatogenic subtypes or Sertoli cells. We have documented the specificity of this Pax7 antibody in germ cells by the lack of intratubular signals in Vasa-Cre Pax7f/f (i.e., germ cell-conditional knockout of Pax7) [14].

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Omission of primary antibody is a helpful negative control; with the recommended detection reagents and conditions, no nonspecific signals should be observed. Blocking peptide could also be used as a control in principle, since a blocking peptide corresponding to the Pax7 monoclonal antibody epitope has been defined [14]. Although the blocking peptide is very effective as a control in Western blotting, we have not used this reagent for IF or IHC protocols. 11. The Pax7 antibody and this protocol can be used in rodents, dogs, cats, monkeys, baboons, and humans, as the epitope recognized by the Pax7 antibody is conserved across mammals [14]. In our experience, detection of positive spermatogonia in humans was relatively difficult due to relatively faint signals, which may reflect non-optimal fixation in the archival clinical specimens we employed. With most other species, staining in testes was comparable to that observed in mouse. 12. After the addition of the fluorescent secondary antibody, the slides are light sensitive and should be kept in the dark as much as possible. The FITC channel tends to exhibit more endogenous background fluorescence following formalin- or paraformaldehyde-based fixation. Pax7 localization is thus easier to conduct using the red channel. 13. With IF on paraffin-embedded formalin-fixed tissues, the endogenous autofluorescence can be high. Use of Tris glycine in step 2 and the ammonium chloride in step 3 helps quench autofluorescence due to aldehyde groups in the fixed tissue sections but does not completely eliminate background autofluorescence. The principal advantage of IF over IHC is that it permits simultaneous detection of multiple markers. For optimal visualization and photomicrographs, use a confocal microscope. 14. To reduce crushing of the tubules, the majority of the protocol is performed in the same Eppendorf tube. Solutions can be changed by using a P1000 pipette while carefully avoiding pipetting tubules by placing the pipette tip at the very bottom of the tube. Check the pipette tip to ensure that tubules have not been aspirated. 15. The tubules can be kept at 20  C for up to 6 months in methanol. Portions can be removed for individual experiments as needed. 16. NP-40 is a mild detergent capable of solubilizing the cytoplasmic membrane and therefore allowing better intracellular penetration of antibodies. However, this step can reduce or abolish cytoplasmic staining of some proteins, and therefore caution should be used for colocalization with cytoplasmic proteins [32].

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17. One testis can be additionally divided up at this step into multiple reactions by using needle-nosed tweezers to place a small number of tubules into different Eppendorf tubes with different antibodies. The use of a rocker during the overnight antibody stain ensures even staining of all tubules. 18. It is difficult to visualize tubules that are on top of other tubules. Therefore, the more separated the tubules are, the better the resulting images. If the tubules begin to dry out while transferring to the slide or while separating, use one drop of PBS to rehydrate. 19. The Vectashield used will vary with the amount of tubules on the slide. Add enough to slightly leak out the slides when the cover slip is added. Lightly place the cover slip on top of the slide but do not apply pressure to the top so as not to crush the tubules. 20. Of the three methods to detect Pax7+ SSCs, whole-mount IF is the most labor intensive at the microscope (due to the difficulty of scanning large areas to identify rare Pax7+ spermatogonia). The main advantage of whole-mount immunofluorescence is that it is the only way to distinguish Asingle spermatogonia from other subsets. Nonspecific staining using this method includes the basement membrane, which on the other hand serves as a useful morphological landmark. Again, we recommend that initial efforts be conducted with tubules from neonatal testis due to the greater ease of locating Pax7+ spermatogonia. When staining adult testis, separate the tubules as much as possible on the slide to facilitate imaging with a confocal microscope, as it is difficult to focus on tubules that are on top of one another. 21. Due to extensive phagocytosis of germ cells by Sertoli cells, small blebs of green signal within Sertoli cell cytoplasm should not be misconstrued as evidence of ectopic Cre-mediated recombination within Sertoli cells. We have not observed any evidence of ectopic recombination with any of the described Pax7 lines. References 1. Oatley JM, Brinster RL (2008) Regulation of spermatogonial stem cell self-renewal in mammals. Annu Rev Cell Dev Biol 24:263–286. doi:10.1146/annurev.cellbio.24.110707. 175355 2. de Rooij DG (2001) Proliferation and differentiation of spermatogonial stem cells. Reproduction 121(3):347–354 3. de Rooij DG, Russell LD (2000) All you wanted to know about spermatogonia but were afraid to ask. J Androl 21(6):776–798 4. Oakberg EF (1971) Spermatogonial stem-cell renewal in the mouse. Anat Rec 169 (3):515–531. doi:10.1002/ar.1091690305

5. Yang QE, Oatley JM (2014) Spermatogonial stem cell functions in physiological and pathological conditions. Curr Top Dev Biol 107:235–267. doi:10.1016/B978-0-12416022-4.00009-3 6. Brinster RL, Avarbock MR (1994) Germline transmission of donor haplotype following spermatogonial transplantation. Proc Natl Acad Sci U S A 91(24):11303–11307 7. Nagano MC, Yeh JR (2013) The identity and fate decision control of spermatogonial stem cells: where is the point of no return? Curr Top Dev Biol 102:61–95. doi:10.1016/ B978-0-12-416024-8.00003-9

Pax7+ Spermatogonial Stem Cells 8. Chan F, Oatley MJ, Kaucher AV, Yang QE, Bieberich CJ, Shashikant CS, Oatley JM (2014) Functional and molecular features of the Id4+ germline stem cell population in mouse testes. Genes Dev 28(12):1351–1362. doi:10.1101/gad.240465.114 9. Nakagawa T, Sharma M, Nabeshima Y, Braun RE, Yoshida S (2010) Functional hierarchy and reversibility within the murine spermatogenic stem cell compartment. Science 328 (5974):62–67, doi:science.1182868 [pii] 10.1126/science.1182868 10. Zhang Z, Shao S, Meistrich ML (2007) The radiation-induced block in spermatogonial differentiation is due to damage to the somatic environment, not the germ cells. J Cell Physiol 211(1):149–158. doi:10.1002/jcp.20910 11. Yoshida S (2012) Elucidating the identity and behavior of spermatogenic stem cells in the mouse testis. Reproduction 144(3):293–302. doi:10.1530/REP-11-0320 12. Nagano MC (2003) Homing efficiency and proliferation kinetics of male germ line stem cells following transplantation in mice. Biol Reprod 69(2):701–707. doi:10.1095/ biolreprod.103.016352 13. Abid SN, Richardson TE, Powell HM, Jaichander P, Chaudhary J, Chapman KM, Hamra FK (2014) A-single spermatogonia heterogeneity and cell cycles synchronize with rat seminiferous epithelium stages VIII-IX. Biol Reprod 90 (2):32. doi:10.1095/biolreprod.113.113555 14. Aloisio GM, Nakada Y, Saatcioglu HD, Pena CG, Baker MD, Tarnawa ED, Mukherjee J, Manjunath H, Bugde A, Sengupta AL, Amatruda JF, Cuevas I, Hamra FK, Castrillon DH (2014) PAX7 expression defines germline stem cells in the adult testis. J Clin Invest. doi:10. 1172/JCI75943 15. Keller C, Hansen MS, Coffin CM, Capecchi MR (2004) Pax3:Fkhr interferes with embryonic Pax3 and Pax7 function: implications for alveolar rhabdomyosarcoma cell of origin. Genes Dev 18(21):2608–2613. doi:10.1101/ gad.1243904 16. Lepper C, Conway SJ, Fan CM (2009) Adult satellite cells and embryonic muscle progenitors have distinct genetic requirements. Nature 460(7255):627–631. doi:10.1038/nature 08209 17. Murphy M, Kardon G (2011) Origin of vertebrate limb muscle: the role of progenitor and myoblast populations. Curr Top Dev Biol 96:1–32. doi:10.1016/B978-0-12-3859402.00001-2 18. Buaas FW, Kirsh AL, Sharma M, McLean DJ, Morris JL, Griswold MD, de Rooij DG, Braun

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32. Goldenthal KL, Hedman K, Chen JW, August JT, Willingham MC (1985) Postfixation detergent treatment for immunofluorescence suppresses localization of some integral membrane proteins. J Histochem Cytochem 33(8):813–820 33. Seale P, Sabourin LA, Girgis-Gabardo A, Mansouri A, Gruss P, Rudnicki MA (2000) Pax7 is required for the specification of myogenic satellite cells. Cell 102(6):777–786

Chapter 12 Transplantation as a Quantitative Assay to Study Mammalian Male Germline Stem Cells Aileen R. Helsel and Jon M. Oatley Abstract In mammals, the activities of spermatogonial stem cells (SSCs) provide the foundation for continual spermatogenesis throughout a male’s reproductive lifetime. At present, the defining characteristics of SSCs and mechanisms controlling their fate decisions are not well understood. Transplantation is a definitive functional measure of stem cell capacity for male germ cells that can be used as an assay to provide an unequivocal quantification of the SSC content in an experimental cell population. Here, we discuss the procedure for mice and provide protocols for preparing donor germ cell suspensions from testes directly or primary cultures of spermatogonia for transplantation, enriching for SSCs, preparing recipient males, microinjection into recipient testes, and considerations for experimental design. Key words Spermatogonial stem cell, Transplantation, SSC enrichment, Mouse

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Introduction In mammals, testes are made up of seminiferous tubules wherein the production of millions of genetically unique spermatozoa occurs daily throughout the reproductive lifetime of a male. The process of sperm production, termed spermatogenesis, occurs in three phases. First, diploid spermatogonia, located in the basal compartment near the basement membrane, undergo mitotic proliferation to expand the population while gaining the competence for initiation of meiotic prophase, thereby yielding primary spermatocytes. Second, spermatocytes proceed through two meiotic divisions to produce haploid spermatids. Third, spermatids undergo drastic morphological changes in a process referred to as spermiogenesis that yields an elongate form, eventually being released into the lumen of the seminiferous tubules. The activities of the spermatogonial population provide the foundation for robust and continual spermatogenesis [1, 2]. This population is heterogeneous, consisting of undifferentiated and differentiating subtypes (Fig. 1). The undifferentiated population

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Fig. 1 Schematic of the heterogeneous undifferentiated spermatogonial population in mammalian testes. The stem cell (SSC) pool is maintained via symmetric or asymmetric self-renewal divisions. From this pool, progenitor spermatogonia arise that transiently amplify in number before transitioning from an undifferentiated to differentiating state at periodic intervals and under the influence of retinoic acid. Collectively, these actions provide the foundation for both continuity and robustness of spermatogenesis

is labeled as type A spermatogonia and can be divided further into stem cell and progenitor subsets. Self-renewal by spermatogonial stem cells (SSCs) maintains a foundational pool from which progenitors arise that divide to bolster the population numbers. A round of spermatogenesis is initiated when a majority of the progenitors transition to a differentiating state under the influence of retinoic acid signaling that occurs at periodic intervals (Fig. 1). Thus, the progenitor spermatogonia are truly a transit-amplifying population. The SSC pool serves as a reservoir from which the next cohort of progenitors arise. Thus, the actions of SSCs are the ground state for continual spermatogenesis. The differentiating spermatogonia can be labeled as type A, intermediate, and type B. The type A subset undergoes mitotic divisions, A1–A4 in the mouse, and then transition to the intermediate state. The type B spermatogonia arise from division of intermediate spermatogonia and transition from mitotic to meiotic divisions, thereby becoming preleptotene spermatocytes. A major bottleneck in the field of mammalian male germ cell biology has been the lack of morphological and molecular markers

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that distinguish the subsets of undifferentiated spermatogonia. In most mammals, the undifferentiated type A spermatogonia can be distinguished from differentiating type A spermatogonia based on nuclear morphology [3, 4]. In addition, spermatogonia can be classified based on “chain” or “cohort” identity. In the undifferentiated population, spermatogonia are present as single (As), paired (Apr), or aligned cohorts of 4–16 cells (Aal4–16) [5]. Importantly, type A differentiating spermatogonia can also be found as interconnected chains of 2–16 cells [6, 7]. Indeed, at periodic intervals, every 8.6 days in mice, most Apr and Aal undifferentiated spermatogonia transition to a differentiating state and retain their chain identity until the next division [8–10]. Because the As undifferentiated spermatogonia do not transition to a differentiating state [8], the population has traditionally been considered to represent the SSC pool [5–7, 11]. Although the features of nuclear morphology and chain identity allow for broad classification or spermatogonial subtypes, they do not distinguish SSC from progenitor. Only recently have molecular markers that can distinguish subsets of undifferentiated spermatogonia begun to be defined [12]. During proliferation of the undifferentiated spermatogonia population, divisions of the As spermatogonia have two potential outcomes. Either the daughter cell completes cytokinesis and forms another As, or the two cells remain connected by an intercellular cytoplasmic bridge to form Apr. Upon the next division, Apr produce a cohort of Aal4 and in the absence of retinoic acid signaling the next divisions produce Aal8 and Aal16 cohorts. Undifferentiated spermatogonial chains of greater than Aal16 have not been reported for any mammalian species. Traditionally, the As population has been considered to be the SSC pool and a division that leads to formation of Apr represents transition to the progenitor state and ultimately commitment to the differentiating pathway. From a fundamental perspective, the label stem cell is applied to cells with the capacity to regenerate a cycling cell lineage. Thus, the SSC label should be applied to spermatogonia with a capacity to regenerate the spermatogenic lineage. While the average mouse testis contains 35,000 As spermatogonia, only 3000 spermatogonia possess regenerative capacity [8, 13]. Therefore, the SSC pool is likely a subset of the As population. Furthermore, while identification of As spermatogonia may be helpful in histological applications, isolation of As from a testis digestion is not possible. Due to the rapid cycling of the spermatogenic lineage from puberty until old age, existence of stem cells in the male germline has been widely accepted for decades. However, functional evidence was not provided until 1994 when the laboratory of Dr. Ralph Brinster published seminal findings about the transplantation of cells isolated from the testes of a donor male into the seminiferous tubules of a recipient male in which colonies of donor-derived spermatogenesis were generated [14, 15]. Because

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each donor-derived colony of spermatogenesis in a recipient testis is clonally derived from a single SSC [16, 17], transplantation provides a robust quantitative assay for determining the SSC content of an experimental cell population (Fig. 2). However, several aspects

Fig. 2 The spermatogonial transplantation assay for mice. (a) A donor germ cell suspension can be collected from testes directly or from primary cultures and microinjected into the seminiferous tubules of recipients that are depleted of endogenous germline. If the donor cells possess a ubiquitously expressed reporter transgene (e.g., LacZ), colonies of donor-derived spermatogenesis are clearly observable several months after transplantation. Because each colony is clonally derived from a single donor spermatogonial stem cell (SSC), colony numbers are a direct measure of SSC content in the transplanted cell suspension. (b) Recipient testes that were transplanted with no SSCs, an unfractionated donor testis cell suspension, or a donor testis cell suspension that was subjected to enrichment for SSC content. The differences in donor-derived colonies are related directly to differences in SSC content of the microinjected cell population. Reprinted from the Annual Review of Cell and Developmental Biology, Volume 24, by Annual Reviews, http://www.annualreviews.org

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must be conducted in an appropriate manner for valid and interpretable data to be generated. By far, the transplantation procedure has been most refined for the mouse. Below, we describe methodology utilized in our lab to assay for the stem cell content of a variety of experimental cell populations using the mouse model. Recently, we utilized the methodology to demonstrate that expression of the transcriptional regulator inhibitor of DNA binding 4 (ID4) is a distinguishing feature of SSCs in the male mouse germline [12].

2

Materials

2.1 Recipient Selection

Transplantation is a powerful tool for the study of germline stem cells when used appropriately. Without proper selection of donor and recipient animals, accurate interpretation of outcomes can be challenging. Endogenous germ cells must be depleted or eliminated in recipient testes for efficient engraftment of donor SSCs. Two major recipient models that have been used over the last 20+ years are genetically sterile mutants and wild-type males treated with chemotherapeutic agents or irradiation [14, 15, 18–21]. The use of genetic mutants such as the W/Wv mouse strain in which survival of precursor germ cells is impaired but the soma remain functionally intact provides an optimal environment for donor SSC engraftment and regeneration of spermatogenic colonies [18]. W/WV animals possess a mutation in the c-Kit gene that leads to impaired migration and survival of primordial germ cells during embryogenesis [22]. Thus, the germline in testes of W/Wv males is lacking at birth. An advantage of using this model is the total absence of endogenous germline; thus any regions of spermatogenesis that develop post-transplantation must be derived from donor SSCs, thereby eliminating the need for donor cells to be marked with expression of a transgene. Although qualitative assessment of SSCs in an experimental cell population can be made with donor cells lacking a marker transgene, quantitative assessment of SSC number is not valid for reasons described in sections below. Treatment of mice with chemotherapeutic agents such as busulfan leads to depletion of the endogenous germline including a portion of the SSC pool within 6–8 weeks after treatment [19, 20, 23]. However, complete elimination of the SSC pool is rare and some endogenous spermatogenesis returns over time. For this reason, use of donor cells that are marked by expression of a reporter transgene is critical for distinguishing regions of spermatogenesis that are derived from transplanted vs. endogenous SSCs when using this recipient model.

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Another key aspect of recipient selection is immunological compatibility with an SSC donor. In our hands, an F1 cross of C57BL/6J and 129ScvP is compatible with cells from donor mice that have varying degrees of genetic background from the parental strains. Similarly, the W/Wv mutant is typically maintained as a mixed background primarily from the C57BL/6J strain. 2.2

Donor Selection

2.3

Reagents

Although the SSC content of both freshly isolated testis cell populations and primary cultures of undifferentiated spermatogonia can be analyzed using transplantation, quantitative assessment requires the capacity to determine the boundaries of spermatogenic colonies derived from donor SSCs. A straightforward means to achieve this is by using donor cells that are marked by constitutive expression of a reporter transgene. Use of W/WV recipients allows for qualitative analysis of unmarked donor cells because the endogenous germline is lacking; thus any regions of spermatogenesis must be donor derived. However, endogenous spermatogenesis does return in seminiferous tubules of animals treated with chemotherapeutic agents (e.g., busulfan); thus neither quantitative nor qualitative analysis of unmarked donor cells is valid when using this recipient model. For these reasons, we recommend using donors that possess a constitutively expressed reporter transgene that is easily detectable (e.g., LacZ or fluorescent proteins) whenever possible. In our experience, donor cells marked by expression of a LacZ reporter transgene are ideal because donor-derived colonies of spermatogenesis in recipient seminiferous tubules stain intense blue following incubation with X-Gal which allows for clear distinction of each colony. Also, the recipient testes can be manually disassociated with relative ease to accurately count colony numbers and the tissue preserved for a long period of time. Typically, we utilize the B6.129S7-GtROSA26Sor/J mouse line as a donor or a hybrid F1 version produced by crossbreeding with another line of interest. 1. DPBS-S: 1 % Fetal bovine serum (FBS), 10 mM HEPES, 1 mM pyruvate, 1 mg/ml glucose, 1  104 U/ml penicillin, and 1  10 μg/ml streptomycin in 1 phosphate-buffered saline (PBS) solution (see Note 1). 2. Mouse serum-free culture media (mSFM): 0.2 % Bovine serum albumin (BSA), 5 μg/ml insulin, 10 μg/ml iron-saturated transferrin, 7.6 μeq/L free fatty acids, 3  10 8 M H2SeO3, 50 μM 2-mercaptoethanol, 10 mM HEPES, 60 μM putrescine, 2 mM glutamine, and antibiotics in minimum essential medium alpha (MEMα) (see Note 1). 3. 4 % Paraformaldehyde (PFA): 4 % w/v PFA in 1PBS, pH 7.4 (see Note 2). 4. Trypsin-EDTA solution: 0.25 % Trypsin, 1 mM EDTA.

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5. DNase I solution: 7 mg/ml DNase I in Hanks’ Balanced Salt Solution. 6. Fetal bovine serum (FBS). 7. 1 mg/ml Collagenase in HBSS. 8. Thy1 magnetic beads (Miltenyi Biotec). 9. MACS column (Miltenyi Biotec). 10. LacZ Rinse Buffer: 0.2 M Sodium phosphate, 2 mM magnesium chloride, 0.02 % Triton X-100, and 0.01 % sodium deoxycholate in water. 11. LacZ Staining Solution: 20 mM Potassium ferricyanide, 20 mM potassium ferrocyanide, 1 mg/ml 5-bromo-4-chloro2-indolyl-β-D galactosidase (X-Gal) in LacZ rinse buffer.

3

Methods

3.1 Donor Cell Preparation

3.2 Preparation of a Single-Cell Suspension from Neonate or Pup Testes

Cell suspensions for microinjection into recipient testes can be prepared freshly from testes of donor males at a variety of developmental stages including embryonic (6–13 days post-conception), fetal (14–18 days post-conception), neonatal (postnatal days, PD, 0–2), pup (PD 3–8), prepubertal (PD 9–35), and adult (> PD 35), or from primary cultures of undifferentiated spermatogonia. Typically, neonatal or pup testis cell suspensions that have not been enriched for spermatogonia or SSCs generate 3 and 14 colonies of donor-derived spermatogenesis in testes of busulfan-treated recipients per 105 cells microinjected, respectively [24]. For testes of adult donor males, 3 colonies of spermatogenesis are generated per 105 cells and cell suspensions prepared from a cryptorchid condition produce 18 colonies per 105 cells [25]. 1. Testes are collected into Hanks’ balanced salt solution (HBSS) and the tunica albuginea is removed with fine forceps. It is helpful to use a dissecting microscope for this step. 2. Testes are transferred to a 35 mm petri dish containing 4.5 ml of trypsin-EDTA solution and 0.5 ml DNase I solution followed by gentle pipetting with a 1 ml serological pipette to disassociate the seminiferous cords. 3. The dish is incubated at 37  C for 5 min followed by addition of another 0.5 ml of DNase I solution and the suspension is gently pipetted again. 4. The dish is returned to 37  C for 5 min and step 3 is repeated until all tissue clumps have become disassociated. 5. FBS is added to the suspension at 10 % of the volume to stop the digestion process and gentle pipetting with a P1000 tip is used to ensure that a single-cell suspension has been achieved.

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6. The single-cell suspension is passed through a 40 μm cell strainer that has been prewashed with 1 ml HBSS into a 50 ml conical centrifuge tube. The filter is then washed twice with 2 ml of HBSS. 7. Cells are then pelleted by centrifugation at 600  g for 7 min at 4  C. At this point, cells can be washed twice in mSFM, suspended at a concentration of 1–10  106 cells/ml, and transplanted as a non-enriched cell population. Alternatively, the pelleted cells can be subjected to enrichment for spermatogonia and SSCs using methods described below. 3.3 Preparation of Adult Testis Single-Cell Suspension

When utilizing adult donors, several extra steps are required to deplete interstitial cells that are present in between seminiferous tubules (note that complete elimination of interstitial cells is not possible). 1. Testes are collected into HBSS and the tunica albuginea is removed using fine forceps. Seminiferous tubules are then gently separated using forceps and transferred to a 50 ml conical tube containing an enzyme solution for further disassociation. The protocol described here is sufficient for two donor testes. 2. The crudely disassociated tubule masses are incubated in a 10 ml enzyme solution of collagenase (1 mg/ml in HBSS) and 1 ml DNase I solution is added. The tissue is then drawn into a 5 ml serological pipette several times to facilitate initial disassociation. 3. The digestion tube is then incubated in a 37  C water bath for 10–20 min with gentle swirling every 5 min until the tubule masses have completely separated. Care must be taken to keep the individual tubules intact. 4. The digestion tube is then set in ice for 2 min to allow the tubules to settle and the supernatant containing interstitial cells is carefully removed with a pipette. 5. 10 ml of HBSS is then added and the tube is swirled gently to wash the tubules. The tube is then placed in ice for 2 min to again allow tubules to settle followed by removing the supernatant. This procedure is repeated two more times. At the end, a majority of interstitial cells are removed leaving behind individual seminiferous tubules. 6. To disassociate tubules, a 10 ml solution of trypsin/EDTA containing 10 mg hyaluronidase and 1 ml of DNase I (7 mg/ ml) is added to the digestion tube. Tubules are gently drawn into and out of a 5 ml serological pipette several times and the tube is then incubated in a 37  C water bath for 5 min.

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7. The cell suspension is then pipetted several times and the digestion tube is returned to the water bath for another 5 min. A single-cell suspension is generated by pipetting with a P1000. 8. FBS is added to the digestion solution at a volume of 10 % to stop the enzyme actions and the single-cell suspension is passed through a 40 μm filter that has been prewashed with 1 ml HBSS into a new 50 ml conical tube. The strainer is washed twice with 5 ml of HBSS. 9. Cells are pelleted by centrifugation at 600  g for 7 min at 4  C. At this point, the cell pellet can be suspended in mSFM at a concentration of 1–10  106 cells/ml and transplanted as an unselected population or subjected to enrichment protocols described below. 3.4 Strategies for Enrichment of SSCs

The concentration of SSCs or even spermatogonia in single-cell suspensions generated from testes is low because of the multitude of other germ cell and somatic cell types. Thus, in many experimental scenarios enrichment for SSCs is of benefit. Here, we describe methodologies that are variations of a continuous Percoll gradient and magnetic activated cell sorting (MACS) for cell surface proteins to enrich for spermatogonia and SSCs that were originally reported by Kubota and Brinster [25]. The single-cell suspensions can also be processed for fluorescence-activated cell sorting (FACS) using antibody staining or based on expression of fluorescent protein reporter transgenes.

3.5 Enrichment of Spermatogonia with a Continuous Percoll Gradient

Fractionation of a testis single-cell suspension using a continuous 30 % Percoll gradient yields a bottom pellet that is enriched for SSC content. For adult testes, 12 colonies are generated in recipient testes per 105 cells transplanted which represent 5-fold enrichment compared to the un-fractionated cell population [25]. 1. The cell pellets prepared from adult or pup donor testes described above are suspended in 10 ml of HBSS. 2. 5 ml of the cell suspension is carefully layered on top of 2 ml of 30 % Percoll solution in a 15 ml conical tube. It is essential that the cell suspension is not mixed with the Percoll solution. To aid this, the conical tube can be inverted at a 45 angle and the cell suspension pipetted slowly. Two distinct layers of solution should be visible after dispensing the cell suspension and two tubes are utilized for the 10 ml of donor cell suspension. 3. Tubes are then centrifuged at 600  g for 8 min at 4  C. 4. The supernatant is removed from each tube and the pelleted cells are suspended in 1 ml of DPBS-S and transferred to a new 15 ml conical tube. Cell number is then determined with a hemocytometer and the suspension is centrifuged at 600  g

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for 7 min at 4  C. The cell pellet can then be prepared for transplantation by suspension in mSFM at 1–10  106 cells/ ml or processed for additional fractionation by MACS or FACS. 3.6 Selection of Thy1+ Cells Using MACS

MACS can be employed to fractionate subsets of germ cells based on expression of specific proteins that are localized at the cell surface. For example, MACS selection of testis cells expressing Thy1 produces a population that is enriched for SSCs compared to the unfractionated cell populations of neonatal, pup, and adult testes. However, it is important to note that the Thy1+ testis cell fraction is not pure SSCs nor is it even pure germ cells. The Thy1+ testis cell fraction from pup donors generates 124–230 colonies per 105 cells transplanted which represents an enrichment of 9- to 16fold compared to unsorted pup cells [25–27]. As discussed in Subheading 3.13, this level of colonization equates to only 1 in 22 cells of the Thy1+ fraction being an actual SSC. Thus, while Thy1 fractionation can be used to enrich for SSCs it does not yield a pure population. The following protocol is a modified version of the procedure reported by Kubota et al. [25]. 1. The pelleted cells from Percoll selection are suspended in DPBS-S and Thy1 magnetic beads are added at a dilution of 1:10. The suspension is then incubated at 4  C for 20 min with mixing after 10 min. 2. 1 ml of DPBS-S is added to the incubation solution and the cells are pelleted by centrifugation at 600  g for 7 min at 4  C. 3. The cell pellet is then suspended in 1 ml of DPBS-S. 4. A MACS column is assembled per the manufacturer’s instructions and rinsed with 0.5 ml DPBS-S. 5. The cell suspension is then loaded into the column after which the column is rinsed three times with 0.5 ml DPBS-S each. 6. Adherent cells are then eluted through the column by adding 1 ml of mSFM, total cell number is determined using a hemocytometer, and cells are then pelleted by centrifugation at 600  g for 7 min at 4  C. 7. The cell pellet is washed by suspending in mSFM and again pelleted by centrifugation. 8. Lastly, the cell pellet is suspended in mSFM at a concentration of 1–10  106 cells/ml in preparation for transplantation.

3.7 Preparation of Cells from Primary Cultures of Undifferentiated Spermatogonia

For several mammalian species including mice, rats, rabbits, and hamsters, primary cultures of undifferentiated spermatogonia can be maintained for long periods of time [28–31]. Although the cultures are comprised purely of undifferentiated spermatogonia, only a minor portion of the population is SSCs. For mouse cultures,

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the SSC content ranges from 10 to 20 % [29, 32–35]. A powerful aspect of the primary cultures is that experimental manipulations can be conducted such as treatment with pharmacological agents, RNAi to reduce the expression of genes of interest, overexpression of certain genes, and supplementation of media with soluble growth factors. Combining these treatments with transplantation analyses allows for exploring alterations in maintenance of the SSC pool [27, 36–43]. 1. mSFM is removed with a serological pipette and each well is washed with 1 ml of HBSS. The wash is collected into a 15 ml conical tube. 2. For a 24-well plate format, 0.25 ml of trypsin/EDTA solution is added per well and the plate is incubated at 37  C for 5 min. 3. FBS is then added directly to the well at 10 % of the volume and cells are pipetted gently to generate a single-cell suspension. 4. The cell suspension is then added to the 15 ml tube containing the initial wash. 1 ml of HBSS is added to the well, collected, and then combined in the collection tube. 5. The collection tube is centrifuged at 600  g for 7 min at 4  C. The supernatant is removed and the cells are suspended in 1 ml mSFM. Cell concentration is determined with a hemocytometer and the tube is again centrifuged at 600  g for 7 min at 4  C. 6. The cell pellet is then suspended in mSFM at a concentration of 1  106 cells/ml in preparation for transplantation. 3.8 Microinjection of Donor Cells into Recipient Seminiferous Tubules

While many parameters of donor cell preparation, including enrichment techniques and other treatments prior to transplantation, depend on the goal of the study at hand, one factor that must always be considered is concentration of donor cell suspension. A high concentration of cells can result in clogging of the microinjection needle or too many colonies in recipient testes to accurately determine the boundaries of each, thus making quantification difficult. Also, diluting the cell suspension too much can result in no or very low colony formation in recipient testes. In our hands, an ideal concentration of donor cells is 1  106 cells/ml for enriched cell suspensions. 1. The recipient animal is anesthetized and placed on a platform for visualization through a dissecting microscope (e.g., Olympus SZX7). A midline abdominal incision is then made through the skin and abdominal muscle. 2. A testis is pulled through the abdominal incision and immobilized outside of the body cavity. A sterile cardstock platform placed at the incision site is helpful for immobilizing the testis. Positioning of the testis is important to orient the efferent duct for access by a micropipette needle.

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3. Donor cell suspension is then loaded into the micropipette. In our hands, borosilicate glass with an internal diameter of 0.75 mm and external diameter of 1 mm is effective for creating micropipettes with a puller. The volume loaded into the micropipette for injection depends on the recipient model being used. For busulfan-treated adult recipients, 10 μl can be injected per testis. For adult W/WV recipients, 6 μl of volume can be injected. These volumes will result in filling of 70–80 % of the seminiferous tubules. After filling with cell suspension, the micropipette is inserted into the efferent duct and advanced to the rete testis. Care should be taken to avoid destroying the efferent duct. Also, it is helpful to make a small hole in the efferent duct with a 30-gauge needle through which the micropipette can be inserted (see Note 3). 4. Pressure is then supplied to the micropipette through silastic tubing that is attached to an injector, syringe, or mouthpiece. To avoid damage to the tubules or injection into the interstitial space, the flow rate should be relatively slow, dispensing the volume over a 3–5-min period (see Note 4). 5. After both testes have been filled with donor cell suspension and replaced into the body cavity, the abdominal muscle is closed with braided silk suture and skin closed with wound clips. Care should be taken to avoid catching the fat pad of either testis in the suture. If this were to occur, a cryptorchid condition would ensue and regeneration of donor-derived spermatogenesis would be limited. 3.9 Considerations for Technical and Biological Replication

When utilizing SSC transplantation as a quantitative assay, it is important to design experiments with proper replication. Biological replication (i.e., n values) is applied to the aspect of an experiment being subjected to a treatment. Typically, the donor cell suspension is the subject of treatments or conditions, for example, selection of donor cells for specific attributes (e.g., surface proteins or expression of fluorescent reporters), treatment with siRNAs, treatment with pharmacological agents, or isolation from testes of mutant models. In these scenarios, the donor cell suspension is the biological replicate and should be considered the n-value. The donor cells should be transplanted into the testes of multiple recipients that were subjected to a standard preparation (i.e., busulfan), which would provide technical replication. In some experiments, treatments may be applied to the recipient, for example, testing new methods for depleting the endogenous germline, alteration of somatic cell components, or mutant animals with fertility defects. In these scenarios, the recipient would be the biological replicate (n-value) and a standard donor cell suspension at the identical concentration should be transplanted into each recipient testis. For experiments in which the donor cells are the subject of

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treatments or conditions, we recommend that at least three donor cell suspensions isolated from different animals be used for biological replication and at least four recipients for each biological replicate be used to achieve proper technical replication (i.e., n ¼ 3 and 12 recipient testes for each treatment). 3.10 Analysis of Recipient Testes for Colonies of Donor-Derived Spermatogenesis

Following transplantation recipient animals should be aged at least 8 weeks to encompass multiple rounds of spermatogenesis. This timeframe is essential for assessment of spermatogenic colonies derived from SSCs. In theory, non-stem cell spermatogonia could reform transient colonies of spermatogenesis but these would not persist beyond one round of spermatogenesis. The most useful parameters from transplantation analyses are colony number and colony length. Because each colony is clonally derived from an individual SSC [16, 17], colony number is a retrospective measure of the relative number of SSCs in the microinjected donor cell suspension. Colony length provides a measure of SSC proliferation and regeneration following initial colonization [44]. The validity of collecting these measures depends on the donor cells. Accurate determination of both colony number and length depends on the ability to discern the boundaries of each colony. For this reason, donor cells must be marked by constitutive expression of a reporter transgene such as GFP or LacZ. Analyzing cross sections of recipient testes is not a valid approach for assessing either the number or the length of donor-derived colonies because defining the boundaries is nearly impossible unless strict serial sectioning of the entire testis and 3D reconstruction are utilized. In our hands, donor cells marked by expression of a LacZ transgene that is integrated into the Rosa26 locus are ideal for determining colony number and length.

3.11 LacZ Staining Recipient Testes

1. Testes are collected from euthanized recipients, the tunica albuginea is gently removed, and the tissue is immersed in 4 % paraformaldehyde for 60 min at 4  C. 2. Following fixation, testes are rinsed in LacZ rinse buffer three times for 30 min each at 4  C on a rocking platform. 3. Testes are then incubated in X-Gal staining solution overnight at 35  C on a rocking platform. 4. LacZ rinse buffer is used to wash the testes several times before post-fixation in 10 % formalin to clear the tissue.

3.12 Qualitative and Quantitative Assessment of Donor-Derived Spermatogenic Colonies

Although donor-derived colonies of spermatogenesis in recipient testes can be seen with the naked eye following X-Gal staining (if LacZ-marked cells were used) or under a UV light (if fluorescent protein reporter cells were used), visualizing the boundaries of each colony requires the use of a dissecting microscope. The seminiferous tubules of each recipient testis should be carefully teased apart with fine forceps or hypodermic 30-gauge needles to reveal all

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colonies. Care must be taken not to break individual colonies. For X-Gal-stained recipient testes, each donor-derived colony will be observable as a dark blue region and the length will be variable [15]. The length of each colony can be determined from digital images using software programs such as NIH Image J [44]. Although cross sections of testes from genetically sterile recipient mice (e.g., W/Wv) are useful for qualitative assessment of whether SSCs were present in an experimental cell population, quantitative determination of relative SSC number is challenging because of the difficulty in determining the boundaries of individual colonies. Raw numbers of donor-derived colonies can be normalized to 1  105 cells transplanted for calculating a relative number of SSCs in an experimental cell population. These values can be compared among treatments or conditions to determine changes in SSC content. However, it is sometimes necessary to apply a correction factor for differences in recovered cell number from a donor testis or culture well because a standard number of cells should be transplanted into each recipient testis to reduce variability in colonization efficiency. For example, if 1  105 total cells are isolated from the testes of a control male and 0.5  105 total cells are isolated from the testes of a mutant male and a standard number of cells is transplanted into each recipient testis, it is possible that the number of donor-derived colonies that are generated will be similar. In this scenario, the actual SSC content in mutant testes is less than control because the total number of cells prior to transplant is reduced by 50 %. In contrast, if the mutant cell suspension produces twice as many colonies compared to control, the actual SSC content is no different than control. 3.13 Determination of SSC Purity in Fractionated Testis Cell Suspensions

The stem cell purity of experimental populations or fractions isolated from a tissue is often difficult to determine. When conducted in a quantitative manner, transplantation analyses can provide an assessment of stem cell purity within a cell population. For the mammalian male germline, cell fractions isolated from testes have been reported to produce anywhere from 228 (Thy1+ cells) to 1 (unselected testis populations) colony of donor-derived spermatogenesis per 105 cells transplanted [15, 40, 43, 44]. The colonization efficiency of SSCs when transplanted into busulfan-treated adult recipient testes has been calculated at 5–12 % [11, 20, 40, 42]. Using the formula: [# of cells transplanted/(# of colonies/% colony efficiency)], a relative purity of SSC content in a transplanted cell population can be calculated. To our knowledge, the greatest number of colonies generated from a cell population isolated from donor testes directly that has been reported to date is for the Thy1+ fraction [25, 26]. Using the formula with a 5 % colonization efficiency, the SSC content in the Thy1+ fraction is 1 in 22 cells [100,000 cell transplanted/(228 colonies/5 % colonization efficiency)].

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Another approach for determining stem cell purity in a cell population is to perform transplantation on a limiting dilution scale. For example, an experimental cell population could be transplanted dilution series of 10,000; 1000; 100; 10; and 1 cells. For assessment, one would determine the fewest number of cells that could be transplanted to generate a single colony of donor-derived spermatogenesis. Employing the formula described above would provide a measure of SSC purity. For example, if one colony were produced from as few as ten cells transplanted, a claim of SSC purity could be made if a colonization efficiency of 10 % were employed, [10 cells transplanted/(1 colony/10 % colonization efficiency)] ¼ 1 in 1 or 100 %. At present, definitive evidence that any cell population isolated from the testes of any mammalian species is pure SSCs has not been reported. However, we have recently observed the formation of two colonies of donor-derived spermatogenesis following transplantation of 50 Id4-GfpBright cells that were isolated by FACS from testes of an Id4-Gfp transgenic donor mouse line (A.R. Helsel and J.M. Oatley, unpublished data). Based on a 5 % colonization efficiency, we can calculate the SSC content of the Id4-GfpBright fraction as 1 in 1.25 cells [50 cells transplanted/(2 colonies/5 % colonization efficiency)] which is by far the greatest enrichment reported to date and the population may actually be pure stem cells.

4

Notes 1. Both DPBS-S and mSFM should be sterile filtered. 2. In a hood, heat 1PBS to 60  C on a hotplate and then add 4 % w/v PFA. Solution will be cloudy. Add 10 N NaOH dropwise until the solution becomes clear. Cool and adjust the pH to 7.4. 3. Proper positioning of the micropipette is critical. If it is not advanced into the rete testis, the donor suspension will not enter the seminiferous tubules. On the other hand, advancing the micropipette too far into the rete testis can result in injection into the interstitial space of the testis rather than into the seminiferous tubules. Both cases will result in poor or no colonization. 4. Adding sterile trypan blue solution (0.25 %) to the donor cell suspension at a volume of 5 % allows for visual confirmation that the suspension has been injected into the seminiferous tubules of a recipient testis. When properly injected the cell suspension will be seen coursing throughout the tubules. Injection into the interstitial space between tubules will appear as a cloud of blue underneath the tunica albuginea.

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References 1. Oatley JM, Brinster RL (2012) The germline stem cell niche unit in mammalian testes. Physiol Rev 92(2):577–595. doi:10.1152/ physrev.00025.2011 2. Oatley JM, Brinster RL (2008) Regulation of spermatogonial stem cell self-renewal in mammals. Annu Rev Cell Dev Biol 24:263–286. doi:10.1146/annurev.cellbio.24.110707. 175355 3. Drumond AL, Meistrich ML, Chiarini-Garcia H (2011) Spermatogonial morphology and kinetics during testis development in mice: a high-resolution light microscopy approach. Reproduction 142(1):145–155. doi:10. 1530/REP-10-0431 4. Chiarini-Garcia H, Russell LD (2001) Highresolution light microscopic characterization of mouse spermatogonia. Biol Reprod 65 (4):1170–1178 5. Huckins C (1971) The spermatogonial stem cell population in adult rats. I. Their morphology, proliferation and maturation. Anat Rec 169(3):533–557. doi:10.1002/ar.10916 90306 6. Huckins C, Oakberg EF (1978) Morphological and quantitative analysis of spermatogonia in mouse testes using whole mounted seminiferous tubules, I. The normal testes. Anat Rec 192(4):519–528. doi:10.1002/ar.1091920 406 7. de Rooij DG, Russell LD (2000) All you wanted to know about spermatogonia but were afraid to ask. J Androl 21(6):776–798 8. Tegelenbosch RA, de Rooij DG (1993) A quantitative study of spermatogonial multiplication and stem cell renewal in the C3H/101 F1 hybrid mouse. Mutat Res 290(2):193–200 9. De Rooij DG, Janssen JM (1987) Regulation of the density of spermatogonia in the seminiferous epithelium of the Chinese hamster: I. Undifferentiated spermatogonia. Anat Rec 217(2):124–130. doi:10.1002/ar.109217 0203 10. Kluin PM, de Rooij DG (1981) A comparison between the morphology and cell kinetics of gonocytes and adult type undifferentiated spermatogonia in the mouse. Int J Androl 4 (4):475–493 11. Oakberg EF (1971) Spermatogonial stem-cell renewal in the mouse. Anat Rec 169 (3):515–531. doi:10.1002/ar.1091690305 12. Chan F, Oatley MJ, Kaucher AV, Yang QE, Bieberich CJ, Shashikant CS, Oatley JM (2014) Functional and molecular features of the Id4+ germline stem cell population in

mouse testes. Genes Dev 28(12):1351–1362. doi:10.1101/gad.240465.114 13. Nagano MC (2003) Homing efficiency and proliferation kinetics of male germ line stem cells following transplantation in mice. Biol Reprod 69(2):701–707. doi:10.1095/ biolreprod.103.016352 14. Brinster RL, Avarbock MR (1994) Germline transmission of donor haplotype following spermatogonial transplantation. Proc Natl Acad Sci U S A 91(24):11303–11307 15. Brinster RL, Zimmermann JW (1994) Spermatogenesis following male germ-cell transplantation. Proc Natl Acad Sci U S A 91 (24):11298–11302 16. Kanatsu-Shinohara M, Inoue K, Miki H, Ogonuki N, Takehashi M, Morimoto T, Ogura A, Shinohara T (2006) Clonal origin of germ cell colonies after spermatogonial transplantation in mice. Biol Reprod 75(1):68–74. doi:10. 1095/biolreprod.106.051193 17. Nagano M, Avarbock MR, Brinster RL (1999) Pattern and kinetics of mouse donor spermatogonial stem cell colonization in recipient testes. Biol Reprod 60(6):1429–1436 18. Ogawa T, Dobrinski I, Avarbock MR, Brinster RL (2000) Transplantation of male germ line stem cells restores fertility in infertile mice. Nat Med 6(1):29–34. doi:10.1038/71496 19. Ogawa T, Dobrinski I, Brinster RL (1999) Recipient preparation is critical for spermatogonial transplantation in the rat. Tissue Cell 31 (5):461–472. doi:10.1054/tice.1999.0060 20. Nagano M, Brinster RL (1998) Spermatogonial transplantation and reconstitution of donor cell spermatogenesis in recipient mice. APMIS 106(1):47–55, discussion 56–47 21. Zhang Z, Shao S, Meistrich ML (2006) Irradiated mouse testes efficiently support spermatogenesis derived from donor germ cells of mice and rats. J Androl 27(3):365–375. doi:10.2164/jandrol.05179 22. Mintz B, Russell ES (1957) Gene-induced embryological modifications of primordial germ cells in the mouse. J Exp Zool 134 (2):207–237 23. Ogawa T, Arechaga JM, Avarbock MR, Brinster RL (1997) Transplantation of testis germinal cells into mouse seminiferous tubules. Int J Dev Biol 41(1):111–122 24. Shinohara T, Orwig KE, Avarbock MR, Brinster RL (2001) Remodeling of the postnatal mouse testis is accompanied by dramatic changes in stem cell number and niche accessibility. Proc Natl Acad Sci U S A

Transplantation as a Quantitative Assay to Study Mammalian Male Germline Stem Cells 98(11):6186–6191. doi:10.1073/pnas.11115 8198 25. Kubota H, Avarbock MR, Brinster RL (2004) Culture conditions and single growth factors affect fate determination of mouse spermatogonial stem cells. Biol Reprod 71(3):722–731. doi:10.1095/biolreprod.104.029207 26. Kubota H, Avarbock MR, Brinster RL (2003) Spermatogonial stem cells share some, but not all, phenotypic and functional characteristics with other stem cells. Proc Natl Acad Sci U S A 100(11):6487–6492. doi:10.1073/pnas. 0631767100 27. Oatley JM, Oatley MJ, Avarbock MR, Tobias JW, Brinster RL (2009) Colony stimulating factor 1 is an extrinsic stimulator of mouse spermatogonial stem cell self-renewal. Development 136(7):1191–1199. doi:10.1242/ dev.032243 28. Ryu BY, Kubota H, Avarbock MR, Brinster RL (2005) Conservation of spermatogonial stem cell self-renewal signaling between mouse and rat. Proc Natl Acad Sci U S A 102(40): 14302–14307. doi:10.1073/pnas.0506970 102 29. Kubota H, Avarbock MR, Brinster RL (2004) Growth factors essential for self-renewal and expansion of mouse spermatogonial stem cells. Proc Natl Acad Sci U S A 101(47): 16489–16494. doi:10.1073/pnas.0407063 101 30. Kubota H, Wu X, Goodyear SM, Avarbock MR, Brinster RL (2011) Glial cell line-derived neurotrophic factor and endothelial cells promote self-renewal of rabbit germ cells with spermatogonial stem cell properties. FASEB J 25(8):2604–2614. doi:10.1096/fj.10-175 802 31. Kanatsu-Shinohara M, Muneto T, Lee J, Takenaka M, Chuma S, Nakatsuji N, Horiuchi T, Shinohara T (2008) Long-term culture of male germline stem cells from hamster testes. Biol Reprod 78(4):611–617. doi:10.1095/ biolreprod.107.065615 32. Kanatsu-Shinohara M, Inoue K, Ogonuki N, Miki H, Yoshida S, Toyokuni S, Lee J, Ogura A, Shinohara T (2007) Leukemia inhibitory factor enhances formation of germ cell colonies in neonatal mouse testis culture. Biol Reprod 76(1):55–62. doi:10.1095/biolreprod.106. 055863 33. Kanatsu-Shinohara M, Ogonuki N, Iwano T, Lee J, Kazuki Y, Inoue K, Miki H, Takehashi M, Toyokuni S, Shinkai Y, Oshimura M, Ishino F, Ogura A, Shinohara T (2005) Genetic and epigenetic properties of mouse male germline stem cells during long-term culture.

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genes important for spermatogonial stem cell self-renewal and survival. Proc Natl Acad Sci U S A 103(25):9524–9529. doi:10.1073/ pnas.0603332103 44. Dobrinski I, Ogawa T, Avarbock MR, Brinster RL (1999) Computer assisted image

analysis to assess colonization of recipient seminiferous tubules by spermatogonial stem cells from transgenic donor mice. Mol Reprod Dev 53(2):142–148. doi:10.1002/(sici)109 8-2795(199906)53:2 3.0. co;2-o

Chapter 13 Mouse Fetal Germ Cell Isolation and Culture Techniques Cassy M. Spiller, Guillaume Burnet, and Josephine Bowles Abstract The fetal gonad contains a great variety of differentiating cell populations, of which germ cells make up a small percentage. In order to study germ cell-specific gene and protein expression, as well as determine direct effects of signaling molecules, it is necessary to prepare enriched populations of germ cells and maintain them in culture for several hours to multiple days. The protocols in this chapter are designed to provide a guide for the isolation or enrichment of mouse primordial germ cells (from 9.5 days postcoitum (dpc) to 18.5 dpc) by flow cytometry (Subheading 3.1) or magnetic sorting (Subheading 3.2), followed by primary germ cell culture (Subheading 3.3). Key words Germ cells, FACS, Magnetic sorting, MACS, Serum-free

1

Introduction Mouse fetal gonads, from the time of their elaboration at about 10.5 days postcoitum (dpc), are composed of both somatic and germ cells. In the remaining period before birth, which occurs in mice in the range of 18–22 dpc, appropriate development and sexual fate determination of the germ cells are critical for fertility. In humans inappropriate prenatal germ cell development is relevant not only to potential infertility but is also believed to underlie the formation of germ cell-derived tumors during adult life. Although, as a particular type of stem cell, the germ cells have some intrinsically programmed developmental features it is well established that germ cells also receive and respond to instructive signals that emanate from the surrounding somatic cells. Importantly, it is believed that such signals are entirely responsible for ensuring sex-specific development of germ cells during fetal life. In order to thoroughly understand fetal germ cell development it is often desirable to obtain pure, or at least enriched, germ cell populations. Such cell pools can then be characterized in terms of their transcriptome, epigenome, and protein expression. Isolated germ cells can also be cultured, for short periods of time, so that the

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effects of exposure to factors known to be produced by gonadal somatic cells (candidate instructive agents) can be assessed directly. Here we present protocols for obtaining pure (Subheading 3.1) or enriched (Subheading 3.2) populations of germ cells from mouse fetal gonads and for culturing them for short periods of time in the absence of feeder cells (Subheading 3.3).

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Materials

2.1 Isolation of PGCs by FluorescenceActivated Cell Sorting (FACS)

In the transgenic mouse line Oct4ΔPE:eGFP (OG2) enhanced GFP (eGFP) expression is controlled by the Oct4 (Pou5f1) promoter and distal enhancer [1]; this line is very useful for isolating migratory germ cells (9.5–10.5 dpc) and gonadal germ cells (11.5–18.5 dpc). eGFP expression in this line does not recapitulate Oct4 expression faithfully in that the protein is detectable for a longer period of time, perhaps because the eGFP protein is more stable than the Oct4 protein. Nevertheless, intensity of eGFP expression declines over time and at a faster rate in female germ cells than in male germ cells as is the case for endogenous Oct4. The protocol described in Subheading 3.1 outlines isolation of gonadal germ cells (11.5–18.5 dpc) only. For isolation of migratory germ cells (9.5–10.5 dpc) please see [2] for tissue dissection. The remaining protocol(s) can then be followed for isolation and culture. It is assumed that the user has sufficient understanding and experience with flow cytometry and fluorescent activated cell sorting (or access to a facility that provides this service) for the isolation of germ cells using this protocol. 1. 11.5–18.5 dpc pregnant CD1 females previously mated with OG2 homozygous males (see Note 1). 2. 1 Phosphate-buffered saline (PBS), sterile, for dissecting. 3. 10 cm Culture dishes for dissection of fetuses and gonads. 4. Forceps with sharp tips (watchmaker’s #5 or #55) for dissection, sterile. 5. Dissecting stereomicroscope. 6. 0.05 % Trypsin/EDTA. 7. 18- and 23-gauge needles and 1 ml syringes, sterile, for tissue dissociation. 8. 1 Hanks’ Balanced Salt Solution (HBSS). 9. HBSS + 1 % BSA. 10. 5 ml FACS tubes compatible with flow cytometer. 11. Sorting flow cytometer.

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Enrichment of germ cell populations can also be achieved using germ cell-specific primary antibodies and magnetically-conjugated secondary antibodies that allow for on-column purification. Due to germ cell differentiation as development progresses, different antibodies are used to identify and isolate germ cells at various stages. SSEA-1 is useful for migratory germ cells and early gonadal germ cells (9.5–12.5) [3, 4] while E-cadherin is useful for later gonadal germ cell isolations (13.5–15.5 dpc) [5]. Using this method (outlined in Subheading 3.2), germ cells can be isolated from any wildtype or transgenic line of interest. 1. 11.5–15.5 dpc pregnant females of desired genetic background (see Note 1). 2. 1 PBS, sterile, for dissecting. 3. 10 cm culture dishes for dissection of fetuses and gonads. 4. Forceps with sharp tips (watchmaker’s #5 or #55) for dissection, sterile. 5. 0.05 % Trypsin/EDTA (Invitrogen). 6. 18- and 23-gauge needles and 1 ml syringes, sterile, for tissue dissociation. 7. MACS buffer: 1 PBS, 2 mM EDTA, and 0.5 % BSA (Sigma Aldrich). 8. MACS MS separation columns (Miltenyi Biotech). 9. MACS pre-separation filters (Miltenyi Biotech). 10. Anti-SSEA-1 (CD115) microbeads, mouse (Miltenyi Biotech)— for 9.5–12.5 dpc. 11. Anti-E-Cadherin, mouse (BD Biosciences)—for 13.5–15.5 dpc. 12. Anti-mouse IgG 13.5–15.5 dpc.

microbeads

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13. MACS MultiStand (Miltenyi Biotech). 14. MS Colum Adapter (Miltenyi Biotech). 15. 10 ml Collection tubes. 2.3 Primary Fetal Germ Cell Culture

Fetal germ cells do not survive well in culture. A number of protocols have been developed for the culture of germ cells isolated from fetal mouse tissues, but to our knowledge these invariably rely on the presence of serum, feeder cell layers, or active signaling molecules (examples include refs. 6, 7). Although survival is certainly aided by the presence of serum, feeder layers, and “survival factors,” we aimed to develop chemically defined media useful for studies in which the effect of various signaling molecules can be tested. For such studies (see Subheading 3.3), we find that short-term culture (less than or up to 24 h) is informative and that this can be supported in “serum-free” medium as detailed below. As it is well

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known that the combination of kit ligand (also known as stem cell factor or Steel factor), basic fibroblast growth factor, and leukemia inhibitory factor (KL, bFGF, LIF) stimulates germ cells to form embryonic germ cell colonies [8, 9] we prefer not to include any of these compounds in our cultures. For cultures longer than 24 h we use “serum-containing” medium. 1. 10 ml Serum-free media (see Note 2): 1.5 ml of KnockOut Serum Replacement (Invitrogen), 8 ml of KnockOut Dulbecco’s modified Eagle’s medium (KO DMEM) (Invitrogen), 50 μl 25 μg/ml penicillin/streptomycin (Invitrogen) (see Note 3), 100 μl 2 mM Glutamax-1 (Invitrogen), 100 μl 0.1 mM MEM nonessential amino acids (Invitrogen), 100 μl 1 insulin‐transferrin‐sodium selenite media supplement (ITS) (Sigma Aldrich), 100 μl 1 N2 supplement (Invitrogen), 100 μl 1 mg/ml N‐acetyl‐L‐cysteine (NAC) (Sigma Aldrich) (see Note 4), and 1.8 μl of a 1/25 dilution of 0.1 mM β‐mercaptoethanol (Invitrogen). See Note 5. 2. 10 ml Serum-containing media (see Note 2): 1.5 ml of 15 % fetal bovine serum (FBS, AusGenx) (see Note 6), 8 ml of DMEM (Invitrogen), 50 μl 25 μg/ml penicillin/streptomycin (Invitrogen) (see Note 3), 100 μl 2 mM Glutamax-1 (Invitrogen), 100 μl 0.1 mM MEM nonessential amino acids (Invitrogen), 100 μl 1 insulin‐transferrin‐sodium selenite media supplement (ITS) (Sigma Aldrich), 100 μl 1 N2 supplement (Invitrogen), 100 μl 1 mg/ml N‐acetyl‐L‐cysteine (NAC) (Sigma Aldrich) (see Note 4), and 1.8 μl of a 1/25 dilution of 0.1 mM β‐mercaptoethanol (Invitrogen). See Note 5. 3. 12-, 24-, or 48-well culture plates, sterile. 4. Humidified 37  C, 5 % CO2 incubator.

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Methods

3.1 Isolation of PGCs by FluorescenceActivated Cell Sorting (FACS)

1. Euthanize 11.5–18.5-day pregnant mice (see Note 7). Harvest embryos from the uterus, dissect embryos away from extraembryonic membranes, and place in culture dish containing 1 PBS. Cut the head off each embryo using forceps, then cut along the ventral midline, and remove internal organs. 2. Remove the genital ridges (with mesonephros attached) from the dorsal wall of the embryo (11.5–15.5 dpc), dorsal wall beneath kidneys (ovaries, 16.5–18.5 dpc), or ventral posterior region (testes; 16.5–18.5 dpc). Using a needle cut away the mesonephros (Fig. 1) and collect the gonads (pooled or sexed; see Note 8) into a 1.5 ml microcentrifuge tube (see Note 9).

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Fig. 1 Location and morphology of male and female gonads. (a) Schematic of genital ridge localisation within a 12.5 dpc embryo. (b) Schematic of testis and ovaries attached to mesonephros at 14.5 and 18.5 dpc

3. Add 1 ml of 0.05 % trypsin/EDTA to each tube and incubate with gentle rocking at 37  C for 5 min (11.5–13.5 dpc) or up to 10 min (14.5–18.5 dpc). Using an 18-gauge needle gently syringe five times to break up any larger tissue that has not dissociated, followed by a 23-gauge needle to achieve a singlecell suspension. Add 500 μl of HBSS + 1 % BSA to dilute the trypsin (see Note 10). Centrifuge at 500  g for 10 min at 4  C. Remove supernatant and wash once with 1 ml HBSS. Resuspend the pellet in 0.5 ml HBSS and ensure a single-cell suspension by passing through a 23-gauge needle once more. Place in 5 ml FACS tube on ice and proceed to flow cytometer. 4. Viable cells are gated based on forward scatter (FSC) vs. side scatter (SSC) profile (see Note 11). eGFP vs. FSC will delineate a highly eGFP-positive population of cells that are the germ cells (Fig. 2). Exclude doublets (see Note 12). Gate this population for collection as germ cells and the remaining eGFPnegative cells as somatic cells. Due to the brightness of the eGFP it is not necessary to provide a GFP-negative control, although dissection of another tissue (such as kidney or limb), processed using the above method, could serve this purpose. 5. Sorted germ cell (eGFP+) and somatic cell (eGFP ) populations can be immediately processed for RNA extraction, DNA extraction, immunocytochemistry (following cytospinning), or primary cell culturing (see Protocol 3.3). The germ cell population isolated by this method is highly pure, although there will be some germ cell contamination of the somatic population (Fig. 3).

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Fig. 3 Assessment of germ cell purity from OG2 isolations. (a) Expression of germ cell marker Mvh in GFP+ve (germ cell) and GFP ve (somatic cell) populations collected from male and female gonads at 13.5 dpc. (b) Expression of somatic cell markers Sox9 (male) and Follistatin (Fst; female) in GFP+ve (germ cell) and GFP ve (somatic cell) populations collected from male and female gonads at 13.5 dpc 3.2 Isolation of PGCs by MACS

1. Euthanize 11.5–15.5-day pregnant mice (see Note 7). Harvest embryos from the uterus and dissect embryos away from extraembryonic membranes and place in culture dish containing PBS. Cut the head off each embryo using forceps, then cut along the ventral midline, and remove internal organs. 2. Remove the genital ridges (with mesonephros attached) from the dorsal wall of the embryo (11.5–15.5 dpc). Using a needle cut away the mesonephros (Fig. 1; see Note 13) and collect the gonads (pooled or sexed; see Note 9) into a 1.5 ml microcentrifuge tube (see Note 14). 3. Add 1 ml of 0.05 % trypsin/EDTA to each tube and incubate with gentle rocking at 37  C for 5 min (11.5–13.5 dpc) or up to 10 min (14.5–15.5 dpc) (see Note 10). Using an 18-gauge needle gently syringe five times to break up any larger tissue that has not dissociated, followed by a 23-gauge needle to achieve a single-cell suspension. Add 1 ml of MACS buffer to dilute the trypsin. Centrifuge at 500  g for 10 min at 4  C. Remove supernatant and wash once with 1 ml MACS buffer followed by centrifugation. 4. Resuspend the pellet in 80 μl MACS buffer and 20 μl of antiSSEA-1 (CD115) microbeads (9.5–12.5 dpc) OR in 94 μl MACS buffer and 4 μl of anti-E-cadherin (13.5–15.5 dpc) and incubate with agitation at 4  C for 45 min. Add 1 ml of MACS buffer and centrifuge at 500  g for 10 min at 4  C. Remove supernatant and wash once with 1 ml MACS buffer followed by centrifugation. 5. If using anti-SSEA-1 (CD115) microbeads (9.5–12.5 dpc) proceed to step 6. If using anti-E-cadherin (13.5–15.5 dpc),

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resuspend the pellet in 80 μl MACS buffer and 20 μl of antimouse IgG microbeads and incubate with agitation at 4  C for 15 min. Add 1 ml of MACS buffer and centrifuge at 500  g for 10 min at 4  C. Remove supernatant and wash once with 1 ml MACS buffer followed by centrifugation. 6. Resuspend cells in 1.5 ml MACS buffer and apply to a presoaked MiniMACS column on magnetic backing. Somatic cells are eluted with 3 ml of MACS buffer into 5 ml collection tube below column (see Note 15). Remove the column from the magnetic backing, place into a 2 ml microfuge tube, and elute germ cells in 2 ml MACS buffer (see Note 16). 7. Sorted germ cell and somatic cell populations can be immediately processed for RNA extraction, DNA extraction, immunocytochemistry (following cytospinning), or primary cell culturing (see Subheading 3.3). This method provides an enriched germ cell population that will carry some somatic cell contamination (see Note 17). There will also be some germ cell contamination of the somatic population. 3.3 Primary Fetal Germ Cell Culture

1. Pellet germ cells by centrifugation at 500  g for 10 min at 4  C. Remove supernatant and resuspend in an appropriate volume of serum-free or serum-containing media. The volume used for resuspension should be slightly more than half the total volume of media required for all wells. Divide suspended germ cells into culture plate cells adding half the final planned volume per well (see Note 18). 2. Prepare treatments (dilute active molecules/chemical inhibitors in media) in microfuge tubes at twice the final concentration required. After mixing gently add treated media to each culture plate well to make up to the final volume required (add the same volume of media as the volume of cells in media already dispensed). We generally use 0.5 ml (48-well), 1 ml (24-well), or 2 ml (12-well plates) (see Note 19). 3. Incubate plates in a humidified 37  C, 5 % CO2, incubator. Serum-free germ cell culturing can be maintained for up to 24 h. Serum-containing germ cell culturing can be maintained for up to 48 h. 4. To harvest cells pipette media up and down and transfer to a microfuge tube. Wash each well at least once with PBS and further pipetting to ensure good cell recovery (see Note 20). Centrifuge at 500  g for 10 min to pellet cells and proceed to RNA extraction, DNA extraction, or immunocytochemistry (following cytospinning).

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Notes 1. All protocols involving live animals must be reviewed and approved by an animal ethics committee and must conform to government regulations for the care and use of laboratory animals. The OG2 line was originally generated on a CBA/ CaJ X C57BL/6J background and we maintain it on a C57BL/ 6 background, but in order to ensure large litter sizes and large numbers of germ cells per gonad we mate homozygous OG2 studs with wild-type CD1 outbred females. Usually a minimum of two or more litters is pooled for this protocol. Noon on the day of plug is counted as 0.5 dpc. 2. Media is prepared under sterile conditions in a Class II Biological Hazard laminar flow hood. 3. Germ cell survival is substantially improved when penicillin and streptomycin concentrations are lower than is normal for cell culture. 4. NAC is an antioxidant that substantially improves germ cell survival [6] 5. Make media fresh prior to culturing. 6. Use FBS that has been cell culture tested. 7. Additional reagents and equipment for euthanasia of the mice are described in [10]. 8. Sexing gonads at 11.5 dpc is only possible by PCR and therefore males and females are generally pooled at this time point. If sexing is required, each embryo is dissected into a 24-well plate containing warm DMEM + 10 % FBS, and a tail tip is taken for PCR sexing as per protocol [11]. Once results are determined (2–3 h total time) embryos are separated based on sex and gonads dissected as per this standard protocol. 9. It is possible to isolate germ cells from single gonad pairs, although germ cell recovery is very low due to loss of cells during the dissociation process. Usually two or more sexed, pooled litters are harvested at a time to ensure good germ cell yield. 10. To minimize cell damage and death keep trypsin incubation time to a minimum and syringe gently for the least number of times to achieve a single-cell suspension. 11. The germ cells cluster with a high FSC and low SSC due to their large size and low cellular complexity, and are usually identifiable by these features alone. 12. Excluding doublets, along with ensuring a single-cell suspension is achieved, will ensure highest purity of germ cells.

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13. When isolating populations using E-cadherin (13.5–15.5 dpc) it is imperative that all mesonephric tissue is cleanly removed due to the presence of E-cadherin-positive cells within this tissue. 14. It is possible to isolate germ cells from single gonad pairs, although germ cell recovery is very low due to loss of cells during the dissociation/washing process. Usually two or more pregnant females are harvested at a time to ensure good germ cell yield. 15. Flow-thru containing somatic cells can be passed back through the column to maximally increase germ cell yield. Following this, pass another 1 ml MACS buffer through column before eluting germ cells. 16. Once the MACS buffer has flowed through the column via gravity, gently insert the pusher into the top of the column to force out the last few microliters of buffer and cells. 17. Minimize somatic cell contamination of the germ cell population by ensuring a single-cell suspension after dissociation; somatic cells that are bound to germ cells will be isolated as “germ cells” using this technique. Germ cell purity, determined by alkaline phosphatase staining [12], is about 90 % using this protocol. 18. Because purified germ cells are quite adherent we avoid loss by adding them straight to cell culture plates rather than preparing the various treatments in tubes. 19. Germ cells are cultured sparsely, seeded at approximately 20,000 cells per well (24 well plate) or as low as 5000 cells per well (48-well plate). 20. Germ cells become quite adherent to the culture dish during culturing. Be sure to thoroughly suspend the cells by pipetting upon harvesting.

Acknowledgments This work was supported by the Australian Research Council [DP140104059] and Cancer Council Queensland [APP1012325]. C.S. is a University of Queensland Postdoctoral Fellow. References 1. Szabo PE, Hubner K, Scholer H, Mann JR (2002) Allele-specific expression of imprinted genes in mouse migratory primordial germ cells. Mech Dev 115(1–2):157–160, doi: S0925477302000874 [pii]

2. Durcova-Hills G, Surani A (2008) Reprogramming primordial germ cells (PGC) to embryonic germ (EG) cells. Curr Protoc Stem Cell Biol Chapter 1:Unit1A 3. doi:10.1002/ 9780470151808.sc01a03s5

PGC Isolation and Culture 3. Solter D, Knowles BB (1978) Monoclonal antibody defining a stage-specific mouse embryonic antigen (SSEA-1). Proc Natl Acad Sci U S A 75(11):5565–5569 4. Donovan PJ, Stott D, Godin I, Heasman J, Wylie CC (1987) Studies on the migration of mouse germ cells. J Cell Sci Suppl 8:359–367 5. Mackay S, Nicholson CL, Lewis SP, Brittan M (1999) E-cadherin in the developing mouse gonad. Anat Embryol (Berl) 200(1):91–102 6. Farini D, Scaldaferri ML, Iona S, La Sala G, De Felici M (2005) Growth factors sustain primordial germ cell survival, proliferation and entering into meiosis in the absence of somatic cells. Dev Biol 285(1):49–56. doi:10.1016/j.ydbio. 2005.06.036 7. De Miguel MP, Donovan PJ (2000) Isolation and culture of mouse germ cells. Methods Mol Biol 137:403–408. doi:10.1385/1-59259066-7:403

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8. Resnick JL, Bixler LS, Cheng L, Donovan PJ (1992) Long-term proliferation of mouse primordial germ cells in culture. Nature 359 (6395):550–551. doi:10.1038/359550a0 9. Matsui Y, Zsebo K, Hogan BL (1992) Derivation of pluripotential embryonic stem cells from murine primordial germ cells in culture. Cell 70 (5):841–847, doi:0092-8674(92)90317-6 [pii] 10. Donovan J, Brown P (2006) Euthanasia. Curr Protoc Immunol Chapter 1:Unit 1 8. doi:10. 1002/0471142735.im0108s73 11. McFarlane L, Truong V, Palmer JS, Wilhelm D (2013) Novel PCR assay for determining the genetic sex of mice. Sex Dev 7(4):207–211. doi:10.1159/000348677 12. Lawson KA, Dunn NR, Roelen BA, Zeinstra LM, Davis AM, Wright CV, Korving JP, Hogan BL (1999) Bmp4 is required for the generation of primordial germ cells in the mouse embryo. Genes Dev 13(4):424–436

Chapter 14 Rattus norvegicus Spermatogenesis Colony-Forming Assays F. Kent Hamra Abstract Knowledge gaps persist on signaling pathways and metabolic states in germ cells sufficient to support spermatogenesis independent of a somatic environment. Consequently, methods to culture mammalian stem cells through spermatogenesis in defined systems have not been established. Lack of success at culturing mammalian stem cells through spermatogenesis in defined systems reflects an inability to experimentally recapitulate biochemical events that develop in germ cells during a seminiferous epithelial cycle. Complex germ and somatic cell associations that develop each seminiferous epithelial cycle support such a hypothesis, conceivably explaining why highly pure mammalian spermatogonia have not developed into meiosis, much less through meiosis without somatic cells. Here, we outline an in vitro spermatogenesis colony-forming assay to study how differentiating spermatogonial syncytia develop from rat spermatogonial stem cell lines. Robust spermatogonial differentiation under defined culture conditions will facilitate molecular biology studies on pre-meiotic steps in gamete development, and provide a soma-free bioassay to identify spermatogenic factors that promote meiotic progression in vitro. Key words Spermatogenesis, Spermatogonial stem cell, Germline stem cell, Self-renewal, Proliferation, Differentiation, In vitro

1

Introduction Spermatozoa are produced within the testes’ seminiferous epithelium by consecutive developmental processes termed spermatogenesis and spermiogenesis [1]. Spermatogonial stem cells maintain spermatogenesis through their unique abilities to self-renew or produce spermatogonial syncytia that differentiate through meiosis to form haploid gametes termed round spermatids [2]. Round spermatids can then undergo the male germline-specific postmeiotic process, spermiogenesis, to differentiate into fully elongated spermatozoa [1]. In mammals, spermatogonial stem cells reside within a population of “A-single” (As) spermatogonia [3–5]. As spermatogonia divide to renew germline stem cells but can also produce syncytia

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of early “undifferentiated” spermatogenic progenitors termed A-paired (Apr) and A-aligned (Aal) spermatogonia [5]. In rodents, undifferentiated spermatogonia mitotically arrest during seminiferous epithelial cycle stages VI–VIII, and then transform into “differentiating” type A1 spermatogonia under control of retinoic acid (active vitamin A derivative) and KIT ligand (KITL) [6, 7]. Type A1 spermatogonia re-enter the mitotic cell cycle and give rise to subsequent generations of differentiating spermatogonia (types A2 > A3 > A4 > Int > B) [8], by which time germ cell numbers/ syncytium can be amplified >100-fold prior to entering meiosis to form spermatocytes [9]. Rodent spermatogonial stem cells can be maintained for a long term in culture [10], but can only be cultured through meiosis in recipient testes [2, 11], or in organ culture within seminiferous tubules [12, 13]. Chemically defined culture systems that support soma-free spermatocyte maturation and spermatid development from mammalian spermatogonial stem cell lines remain to be established, but will provide controllable methodology to investigate molecular mechanisms that govern pre-meiotic, meiotic, and post-meiotic processes in germ cell development. Glial cell line-derived neurotrophic factor (Gdnf ) and fibroblast growth factor (Fgf ) genes encode polypeptides that support proliferation of mouse [14, 15], rat [16, 17], and hamster [18] spermatogonial stem cells in culture, and are essential components in serum-free media that maintain mouse spermatogonial stem cell proliferation for a long-term without somatic cells [19]. In contrast to highly defined culture systems that maintain rodent spermatogonial stem cell proliferation, prominent knowledge gaps exist on signaling pathways in germ cells needed to support additional premeiotic, meiotic, or post-meiotic spermatogenic differentiation steps independent of somatic cells. Recently, two polypeptides, neuregulin-1 (NRG1) and KITL, were reported to robustly support syncytial growth of differentiating spermatogonia without somatic cells by signaling germ cell survival in response to all-trans-retinoic acid (ATRA) [20]. By supplementing a serum-free rat spermatogonial culture medium (SG medium) with ATRA and NRG1 (or KITL), an effective serum-free rat spermatogonial differentiation medium was formulated (SD medium) [20]. This chapter outlines how rat spermatogonial stem cell lines derived and propagated in SG medium are stimulated to undergo pre-meiotic spermatogenic differentiation by culturing in SD medium. Spermatogonial lines sub-cultured in GDNF-containing medium commonly grow in colonies that form “clumps,” which contain stem spermatogonia based on their ability to regenerate spermatogenesis in recipient testes [14–18, 21]. Spermatogonial lines cultured in GDNF-containing medium also contain progenitor-like “undifferentiated” type A spermatogonia based

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on ineffectiveness at regenerating spermatogenesis in recipient testes [14, 22, 23]. Because undifferentiated spermatogonia are prone to syncytia formation when propagated in GDNF-containing medium (Fig. 1a), we find that TEX14+ syncytial clones containing two or more cells are commonly passaged during subculture, which more rapidly develop into larger syncytia upon differentiation (Fig. 1b). To avoid harsher dissociation treatments needed to generate single spermatogonial suspensions from spermatogonial lines, after passaging, spermatogonia are pre-incubated in a modified SG medium termed As-SG medium for 5–8 days to select against clump and syncytia formation [20]. Pre-incubation in As-SG medium biologically enriches spermatogonial cultures with As spermatogonia prior to initiating differentiation in SD medium. Unlike SG medium, As-SG medium does not contain GDNF (Fig. 2), which reduces spermatogonial syncytial size and heterogeneity within cultures prior to initiating differentiation in SD medium. Shorter pre-incubation times in As-SG medium prior to initiating differentiation in SD medium increase heterogeneity in spermatogonial syncytia size and differentiation state within cultures. Conversely, longer incubation times in SD medium increase mean nuclei number/syncytium/culture and effectively deplete spermatogonia of their ability to regenerate spermatogenesis in recipient testes [20].

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Subculture Undifferentiated Rat Spermatogonial Line on MEFs in SG Medium

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Molecular Biology Assays Fig. 2 Spermatogonial culture media for in vitro spermatogenesis colony-forming assays. Undifferentiated rat spermatogonial lines are derived and maintained on DR4 MEFs in SG medium (contains GDNF and FGF2). Undifferentiated spermatogonial colonies grow in nonuniform clumps (top image) [14, 17]. Spermatogonia are passaged onto new DR4 MEF feeder layers (shown), or laminin-coated dishes, and cultured in As-SG medium (contains FGF2 and lapatinib) to enrich cultures with As spermatogonia (middle image). Spermatogenic differentiation is induced by culturing spermatogonia in SD medium (contains FGF2, GDNF, ATRA, NRG1, and/or KITL). *GDNF is not an essential component of SD medium, but increases SD medium effectiveness on laminin [20]. **NRG1 and KITL are interchangeable, yet essential SD medium components required for differentiating spermatogonial syncytia to survive on laminin. DR4 MEFs produce NRG1/KITL-like factors, and thus, neither NRG1 nor KITL is required in SD medium to support spermatogonial differentiation on MEFs [20]. Green ¼ tgGCS-EGFP germline-specific transgene [27]. Scale, 100 μm

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Rattus norvegicus spermatogenesis colony assays provide flexible, scalable, and highly defined experimental platforms to investigate molecular mechanisms regulating spermatogonial proliferation, differentiation, and degeneration. CRISPR/Cas9mediated gene targeting in rat spermatogonial lines enables recessive traits controlling spermatogonial fate to be analyzed using in vitro and in vivo spermatogenesis colony-forming assays [24]. Herein, we tailor a spermatogenesis colony-forming assay to study clonal development of differentiating spermatogonial syncytia from cultures enriched with undifferentiated, rat type As spermatogonia (Fig. 2).

2

Materials The following in vitro spermatogenesis colony-forming assay was established with undifferentiated, rat spermatogonial lines derived on DR4 MEFs (Fig. 3a) [17, 21, 25]. Rat spermatogonial lines are endowed with spermatogonial stem cells that regenerate and maintain spermatogenesis for a long term in recipient rat testes (Fig. 3b, c) [24, 26]. Based on spermatogenic colonies formed/ recipient testis/number of germ cells transplanted (~1/65), rat spermatogonial lines represent a most potent source of germline stem cells [17, 20, 21]. Functionally validated rat spermatogonial lines from wild-type rats, or transgenic rats that express germlinespecific EGFP [27] and dtTOMATO markers, can be requested from the Hamra lab. Fully functional, genetically modifiable rat spermatogonial lines can also be derived from 23- to 24-day-old Sprague-Dawley rats as detailed in Chapter 12 (Rat Spermatogonial Stem Cell Mediated Gene Transfer), Springer Press edition: “Advanced Protocols for Animal Transgenics” edited by Shirley Pease and Thomas L. Saunders (see Note 1) [25].

2.1 Spermatogenesis Colony-Forming Assays

1. Laboratory should be equipped with staff and basic equipment to conduct eukaryotic cell culture under sterile conditions: Eppendorf pipets and tips, serological pipets, 0.2 μm filters, swinging bucket table-top centrifuge, media water bath at 34–35  C, hemocytometer, fluorescence/phase-contrast microscopes, CO2-supplied cell culture incubators set at 36–37  C, and dedicated refrigerators and freezers for storing reagents. 2. Certified biosafety cabinet in clean, low-traffic environment dedicated to conducting eukaryotic cell culture under sterile conditions. 3. Rat spermatogonial stem cell line: request frozen stocks of functionally validated spermatogonial lines from F.K. Hamra. Alternatively, isolate undifferentiated rat As and Apr

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spermatogonia fresh, or derive primary rat spermatogonial lines as outlined (see Note 1) [25]. 4. Costar Clear TC-Treated Microplates, Individually Wrapped, Sterile 6 well plate, 12 well plate, 24 well plate, 48 well plate or 96 well plate (Corning, Inc.) (see Note 2). 5. Dulbecco’s modified Eagle’s medium: Ham’s F12 medium 1:1 (DHF12): (Sigma, Inc.). 6. Antibiotic–antimycotic solution (100): 10,000 U/mL Penicillin G sodium (U/v), 10,000 μg/mL streptomycin sulfate (w/v), and 25 μg/mL amphotericin B (w/v) (Life Technologies Inc.). 7. DHF12 solution, Dulbecco’s modified Eagle’s medium: Ham’s F12 medium 1:1, 1 % antibiotic–antimycotic solution (v/v). 8. 1 mg/ml Laminin solution (Sigma, Inc.). 9. Parafilm M (Bemis, Inc.). 10. Heat-inactivated fetal bovine serum (FBS) (Tissue Culture Biologicals, Inc.). 11. Dulbecco’s modified Eagle’s medium-high glucose (DMEMhigh glucose) (Sigma, Inc.). 12. MEF medium: Dulbecco’s modified Eagle’s medium, 1.5 g/l sodium bicarbonate (v/v), and 15 % heat-inactivated FBS (v/v). 13. DR4 Mouse Embryonic Fibroblasts (MEFs) (ATCC). 14. Irradiator source to mitotically arrest DR4 MEFs (optional: see Note 3). 15. Recovery Cell Culture Freezing Medium (Life Technologies, Inc.). 16. T225 Flasks Angled Neck (Corning, Inc.). 17. Gelatin from Porcine Skin-Type A (Sigma, Inc.). 18. 0.1 % Gelatin solution: Dissolve gelatin in ultrapure laboratory-grade water (1 g gelatin/L) and autoclave on liquid cycle. Store stock solution at 4  C up to 2 months; filter-sterile before each use. 19. Dulbecco’s phosphate-buffered saline (PBS) (Sigma, Inc.). ä Fig. 3 (continued) after transplanting tgGCS-EGFP+ rat spermatogonia. Scale, 200 μm. (Right) Relationship between number of tgGCS-EGFP+ spermatogonia transplanted and donor-derived spermatogenic colonies formed/testis scored d27 after transplantation (SEM, n ¼ 5 rats, right testis transplanted). Scale, 200 μm. (c) Donor-derived spermatogenesis in rat seminiferous tubules d178 after transplanting tgGCS-EGFP+ rat spermatogonia (green) clonally expanded from an individually picked colony in SG Medium. Scale, 200 μm

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20. B-27 Supplement minus vitamin A (50) (Life Technologies, Inc.). 21. L-Glutamine (Life Technologies, Inc.). 22. Recombinant FGF2 (Sigma, Inc.). 23. Recombinant GDNF (R&D Systems, Inc). 24. Recombinant NRG1β1, T176-K246 (R&D Systems, Inc.). 25. Recombinant KITL (R&D Systems, Inc.). 26. All-trans-retinoic acid (Sigma, Inc.). 27. Lapatinib (LC Laboratories, Inc.). 28. 2-Mercaptoethanol. 29. Protease-, nuclease-, and fatty acid-free bovine serum albumin (Calbiochem, Inc.). 30. 0.1 M Sodium phosphate buffer, pH 7.2. 31. 4 % Paraformaldehyde solution, pH 7.2: 10 ml 16 % Paraformaldehyde (Thermo Scientific, Inc.), 30 ml sodium phosphate solution, pH 7.2 (v/v). 32. Methanol (Pharmco-AAPER, Inc.). 33. Germ cell marker antibodies: Rabbit anti-rat DAZL IgG [28]; rabbit anti-rat TEX14 IgG [29] (request primary antibody stocks to DAZL and TEX14 from F.K. Hamra). 34. Spermatogonial marker antibodies: Mouse anti-ZBTB16 IgG (Active Motif, Inc) and/or mouse anti-SALL4 IgG (clone 6E3; Abnova). 35. Secondary antibodies: Highly cross-absorbed AlexaFluor 595conjugated, goat anti-mouse IgG (Life Technologies), AlexaFluor 488-conjugated, goat anti-rabbit IgG (Life Technologies). 36. Hoechst 33342 dye. 37. Roche Western Blocking Reagent (Roche Applied Science; distributed by Sigma, Inc.). 38. Triton X-100.

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Methods Steps are outlined to conduct in vitro spermatogenesis colonyforming assays initiated by seeding rat spermatogonial stem cell lines onto laminin matrix (Subheading 3.1) or DR4 MEF feeder layers (Subheading 3.2) in As-SG medium for a pre-incubation period prior to conditionally driving spermatogenic differentiation using SD medium (Fig. 2). Spermatogenesis colony-forming assays conducted on laminin in SD medium provide controllable

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Fig. 4 Analyzing spermatogonial syncytia development in culture. (a) Spermatogonial syncytia at the 2-cell, 4cell, and 8-cell steps in development after culturing for 6 days on laminin in SD medium containing 5 nM KITL [20]. Spermatogonia were co-labeled with an antibody to rat TEX14 (green; top) and the nuclear dye, Hoechst 33342 (blue; bottom). The rat TEX14 IgG selectively labels germ cell cytoplasm and forms concentrated foci at ring canals marking cytoplasmic bridges within a spermatogonial syncytium. (b) Spermatogonial syncytia containing 4–16 cells scored/well (0.96 cm2) by counting relative numbers of TEX14+ foci and Hoechst 33342+ nuclei/syncytium (see Note 19). Cultures were scored after 6 days in SD media supplemented with respective ligands at 5 nM in place of KITL and NRG1 (SEM, triplicate wells/condition). BMP bone morphogenetic protein, PDGF platelet-derived growth factor. Scale, 200 μm

conditions to study molecular mechanisms critical for spermatogonial differentiation (Fig. 4a). Optionally, MEFs produce paracrine factors that positively impact the spermatogenesis colony-forming assays outlined in Subheading 3.6. In vitro spermatogenesis colony-forming assays on DR4 MEFs or other feeder cell types provide a workable platform to discover/investigate factors that regulate spermatogonial biology. Spermatogenic factor effects can then be probed in more detail without somatic cells on an extracellular matrix, such as a laminin matrix (Fig. 4b) [20, 29, 30]. CRISPR/Cas9-mediated gene editing technology can be combined with in vitro spermatogenesis colony-forming assays to study how genes regulate stem, progenitor, and differentiating spermatogonia fate (Fig. 5) [24].

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Fig. 5 Colony-forming assays to study genetic effects on spermatogonial fate. Clonal dominance assays in SG medium and differentiation assays in SD medium can be used to identify and measure gene mutation effects on spermatogonial fate. *Biallelic mutation frequency/cell/culture catalyzed by CRISPR/Cas9 depends on targeting construct and gene delivery efficiencies

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Prepare laminin-coated dishes the day before plating spermatogonia to initiate colony-forming assays (Subheading 3.6). 1. Prepare frozen stocks of laminin to avoid multiple freeze–thaws. Vials containing 1 mg/ml laminin solution are received frozen from the manufacturer. To make frozen stocks, thaw one vial of mouse laminin on ice (requires 1–2 h). Once thawed, make ~six 150 μl laminin solution aliquots in sterile microfuge tubes on ice. Store the laminin stocks at 80  C for up to 1 year. 2. The day prior to isolating or passaging spermatogonia for colony-forming assays, coat wells in a sterile 48-well culture dish with laminin (~5.9 μg/cm2). To prepare, thaw one 150 μl aliquot from the 1 mg/ml laminin stocks on ice (requires ~30 min) and dilute the entire aliquot volume into a sterile 50 ml tube containing 8 ml DHF12 solution. Slowly swirl or gently rock the tube by hand to mix the contents. 3. Add 0.3 ml of the diluted laminin solution/well into the center 24 wells of a 48-well plastic culture dish (0.95 cm2 wells). 4. To reduce evaporation from laminin-coated wells, add 0.4 ml PBS to the remaining 24 uncoated, “outside” wells. Additionally, wrap the dish with a Parafilm strip and store overnight (16–24 h) at 4  C. 5. The next day, equilibrate the dish to room temperature (22–24  C) within a biosafety cabinet, discard laminin solution from the center 24 wells, wash each laminin-coated well once with 0.4 ml DHF12 solution, and immediately proceed to steps in Subheading 3.6. Coating 24 wells with laminin will be sufficient to conduct colony-forming assays for six or eight test conditions at n ¼ 3 or n ¼ 4 wells/condition, respectively. DR4 MEFs are used to maintain rat spermatogonial stem cell lines [25] and can be used in colony-forming assays [20]. MEF feeder layers require ~2 days to prepare before plating spermatogonia to initiate colony-forming assays. 1. Primary stocks of DR4 mouse embryonic fibroblasts (MEFs) are purchased from ATCC, and expanded according to the manufacturer’s protocol after plating into MEF medium at ä

3.2 Preparing Mouse Embryonic Fibroblast Feeder Layers

Fig. 5 (continued) [24, 25]. **Phenotypes that disrupt spermatogonial viability or proliferation in SG medium preclude the ability to clonally enrich for targeted germline mutations. (Bottom images) Clonally expanded wild-type and Erbb3 knockout spermatogonial lines on laminin after culturing 7 days in As-SG medium and 6 days in SD medium. ***Mutant germline analyses on DR MEFs and/or in recipient testes can also be conducted to investigate spermatogonial phenotypes identified on laminin [20]. Scale, 100 μm

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37  C/5 % CO2. MEFs are sub-cultured up to four passages following their thawing and initial plating from the manufacturer’s vial (see Note 3). 2. Following expansion into T225 flasks, secondary stocks of MEFs are harvested according to the manufacturer’s protocol, irradiated (100 Gy) in MEF medium, and then cryo-preserved in liquid nitrogen for future use by freezing in Recovery Cell Culture Freezing Medium according to the manufacturer’s protocol (see Note 4). 3. Pre-coat tissue culture dishes with a solution of filter-sterilized 0.1 % gelatin for 1 h at room temperature (22–24  C). Rinse 1 with sterile PBS before plating MEFs. 4. Prior to culturing spermatogonia, thaw and plate irradiated DR4 MEFs into gelatin-coated dishes (4.5  105 cells/cm2) in MEF medium (0.2–0.3 ml/cm2) for 16–48 h. Rinse 1 with PBS and pre-incubate in SG medium (0.25–0.3 ml/cm2) for an additional 16–48 h. Discard pre-incubation medium and plate spermatogonia onto the MEFs in fresh SG medium (0.3–0.4 ml/cm2), as described in step 1, Subheading 3.6. 3.3 Formulating Spermatogonial Culture Media

1. Spermatogonial culture medium (SG medium) is prepared by supplementing Dulbecco’s modified Eagle’s medium:Ham’s F12 medium 1:1 in the following order (see Notes 5–7): (a) 1 Concentration antibiotic–antimycotic solution (v/v). (b) 4 mM L-Glutamine (final concentration ¼ 6 mM). (c) 100 μM 2-Mercaptoethanol. (d) 1 Concentration of B27 Supplement Minus Vitamin A (v/v). (e) 6 ng/ml GDNF. (f) 6 ng/ml FGF2. 2. Equilibrate to 34–35  C and filter sterilize.

3.4 Formulating Rat A-Single Spermatogonia Culture Media (As-SG Medium)

1. Rat A-Single Spermatogonia Culture Medium (As-SG medium) is prepared identical to that of SG medium, with the exception that As-SG medium does not contain GDNF (see Notes 8 and 9). Optionally, 1 μM lapatinib (ERBB1, 2, 4 inhibitor) can be supplemented into As-SG medium to help reduce syncytia formation.

3.5 Formulating Spermatogonial Differentiation Medium (SD Medium)

1. Spermatogonial differentiation medium (SD medium) is prepared identical to that of SG medium, except that the concentration of GDNF is reduced to 2 ng/ml, and SD medium is further supplemented with 3 μM ATRA, 40 ng/ ml NRG1β1, and/or 100 ng/ml KITL (see Notes 10 and 11).

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1. Day 1 (d1), plate ~1  103 to 1.5  103 rat spermatogonia/ well (0.95 cm2) in 0.4 ml As-SG medium/well of a 48-well culture dish (0.95 cm2 wells) coated with laminin (Subheading 3.1) or containing irradiated DR4 MEFs (Subheading 3.2) (see Note 12). 2. Pre-incubate spermatogonia in As-SG medium at ~36.5  C, 5 % CO2, for ~6 days (d1–d7) to enrich cultures with As spermatogonia. Change As-SG medium (0.4 ml/well) every 2 days during the pre-incubation period using fresh As-SG medium (i.e., change media on d3, d5, d7) (see Note 13). 3. On d7, change out culture medium by feeding spermatogonia with SD medium (0.4 ml/well) and incubate for 5–8 days at ~36.5  C, 5 % CO2, to produce differentiating spermatogonia (see Note 14). 4. Feed cultures with fresh SD medium every 2 days during the ~1-week differentiation period. Six to eight days in SD medium promotes robust development of spermatogonial syncytia for analysis in colony-forming assays. 5. After 6–8 days in SD medium (~d14), remove medium and fix cells directly in 0.4 ml/well ice-cold 4 % paraformaldehyde and 0.1 M sodium phosphate buffer, pH 7.2, for 20–30 min on ice (see Notes 15–17). 6. Post-fix cells in 0.4 ml/well 20  C methanol for 5 min on ice. 7. Wash fixed cultures 2 with 0.4 ml PBS/well/wash at room temperature (22–24  C) (see Note 18). 8. Label spermatogonial cultures to detect desired molecular markers (see Subheading 3.7). 9. Score spermatogenic units/well (n ¼ 3–4 wells/test condition/experiment, S.E.M.) using a microscope to count spermatogonia/well exhibiting single, paired, or longer syncytial morphologies co-labeled with reagents to detect desired molecular markers (i.e., co-labeling with anti-TEX14 IgG and Hoechst 33342 dye allow individual spermatogonial syncytia to be clearly classified) (Fig. 4) (see Notes 19 and 20).

3.7 Immunofluorescence Labeling on Spermatogonial Cultures

1. Prepare 1 stock of Roche Western Blocking Reagent in 0.1 M phosphate buffer pH 7.2, 0.01 % Triton X-100 (v/v) (Blocking Reagent). 2. Pre-incubate fixed cultures in 0.4 ml Blocking Reagent/well for 1–2 h at room temperature (22–24  C). 3. During the pre-incubation step in Blocking Reagent (step 2), prepare primary antibody solutions by diluting respective antibodies raised to germ cell and/or spermatogonial markers into a required volume of fresh Blocking Reagent: mouse antiZBTB14 IgG at 0.2 μg/ml, mouse anti-SALL4 at 0.2 μg/ml,

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rabbit anti-DAZL IgG at ~5 nM (1/800 dilution) [28], and/ or rabbit anti-TEX14 IgG at ~5 nM (1/800 dilution) [30]. 4. Remove Blocking Reagent from fixed cultures and then add 0.4 ml of respective primary antibody solution/well and incubate overnight (14–24 h) at room temperature (22–24  C). 5. Discard primary antibody solutions and wash cultures three times for 10–30 min per wash with 0.4 ml PBS/well/wash (see Note 18). 6. During the PBS wash step (step 5) prepare respective AlexaFluor 488-conjugated (green fluorophore) and/or AlexaFluor 594-conjugated (red fluorophore) secondary antibody solutions by diluting to ~5 μg/ml (1/400 dilution from 2 mg/ml manufacture stock) when required for indirect labeling (see Note 21). To directly label nuclei with a blue fluorophore, dilute Hoechst 33342 dye (1/2000) directly into the secondary antibody solution. 7. Remove final PBS wash and incubate fixed cultures in 0.4 ml/ secondary antibody solution/well for 60–90 min at 22–24  C. 8. Discard primary antibody solutions and wash fixed cultures three times for 10–30 min/wash with 0.4 ml PBS/well/wash. 9. Add 0.4 ml/PBS to fixed cultures for viewing and storage. Wrap culture plates with a thin strip of Parafilm to minimize evaporation during storage. 10. View labeled cultures and score spermatogenic units as described in step 9 of Subheading 3.6.

4

Notes 1. If one chooses to derive his or her own rat spermatogonial lines, or utilize freshly isolated laminin-binding As/Apr spermatogonia to initiate colony-forming assays, it should be noted that the dispase solution composition previously utilized in published protocols to digest 22–24-day-old rat seminiferous tubules [21, 25, 28, 31–33] was modified by the manufacturer in 2015, and is significantly less effective. We recommend the former dispase digest be replaced by a single-step collagenase digest for 15 min, at 34  C in Dulbecco’s modified Eagle’s medium:Ham’s F12 medium 1:1 (Sigma, Inc.), and 1.2 mg/ ml Clostridium histolyticum Collagenase (Sigma; 2.1 units/mg FLGPA) [25]. Filter sterilize. 2. The described in vitro spermatogenesis colony-forming assay (Subheading 3.6) is routinely conducted using Costar 48-well plates.

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3. Irradiating MEFs prevents them from dividing and allows them to form a more stable feeder layer for spermatogonia to grow on. If an irradiator source is not available, or irradiating one’s own MEFs is not desirable, several companies provide preirradiated DR4 MEFs, but at an elevated price compared to non-irradiated MEFs. As an alternative to irradiating MEFs, MEFs can be mitotically inactivated before use as feeder layers by a simple pretreatment with mitomycin-C [17, 30]. 4. It should also be stressed that primary DR4 MEF lots vary in quality between different companies, and lots from the same company can vary. We find ATCC’s DR4 MEFs consistently reliable for culturing rat spermatogonial lines with potent sperm-forming potential [24]. 5. Prepare 100 stock of 2-mercaptoethanol (¼10 mM) fresh by diluting 7.8 μl 2-mercaptoethanol from the manufacturer into 10 ml Dulbecco’s modified Eagle’s medium:Ham’s F12 medium 1:1. Upon regular use, bottles containing 2mercaptoethanol provided by the manufacturer are replaced with new bottles approximately every 4 months. 6. Polypeptide growth factors (GDNF, FGF2) should be reconstituted as instructed by the manufacturer, but using filtersterilized 0.01 % protease-, nuclease-, and fatty acid-free bovine serum albumin (Calbiochem, Inc.) in PBS (w/v). FGF2 is essential for undifferentiated spermatogonia and DR4 MEF viability in SG medium. GDNF is essential for spermatogonial colony/clump formation on DR4 MEFs in SG medium. 7. Higher GDNF and/or FGF2 concentrations can be used to increase the growth rate of some rat spermatogonial lines when cultured on some lots of DR4 MEFs in SG medium. For example, the original SG medium formulation consisted of 20 ng/ml GDNF and 25 ng/ml FGF2 [21]. 8. As-SG medium is identical to the recently reported SGFmedium [20]. Omitting GDNF from SG medium selects for spermatogonia in the single (As) cell state, and thus selects against syncytial growth of undifferentiated spermatogonial clones/clumps that can complicate colony counts and phenotyping. 9. It should be emphasized that after enriching for As spermatogonia by selection in As-SG medium, cultures will still contain a relatively low percent of Apr and Aal spermatogonia. 10. SD medium is a modified formulation of the serum-free, SG medium [21]. KITL can be supplemented into SD medium at 100 ng/ml in place of NRG1β1, or in combination with up to 40 ng/ml NRG1β1 [20].

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11. NRG1β1 and KITL stimulate respective lapatinib-sensitive and lapatinib-insensitive survival pathways in differentiating spermatogonia [20]. 12. Plating ~103 undifferentiated rat spermatogonia/well in a 48well plate format (0.95 cm2/well) from a given rat spermatogonial line typically yields 150–300 syncytial colonies containing 4–16 nuclei/syncytium/well after 6 days in SD medium (steps 3 and 4, Subheading 3.6). Plating ~103 spermatogonia/ 0.95 cm2 yields well-separated syncytia. 13. Pre-incubation time in As-SG medium is optional, and can be shortened or omitted prior to steps 3 and 4, Subheading 3.6, to expedite studies on spermatogonial differentiation. Spermatogonial numbers/culture and surface area/culture in SG, As-SG, and/or SD medium can also be scaled to best meet scientific aims requiring other types of cellular and molecular analyses. 14. Colony-forming assays in SD medium conducted at 32.5  C, 34.5  C, or 36.5  C yield similar results with respect to differentiating spermatogonial syncytia (ZBTB16 , SALL4 , DAZL+, TEX14+) development. Spermatogonial syncytia develop slower at reduced temperatures. 15. Post-fixation in MeOH promotes adherence to plates and minimizes colony loss during wash steps following primary and secondary antibody labeling. 16. Post-fixation in MeOH permeabilizes paraformaldehyde-fixed spermatogonial membranes, which facilitates antibody delivery into fixed spermatogonia to label intracellular antigens. All the antibodies listed in Subheading 2.1 are compatible with postfixation in MeOH. Post-fixation in MeOH is optional if not compatible with antibody binding to a particular antigen under study because Blocking Reagent contains 0.01 % Triton X-100 detergent. Triton X-100 also permeabilizes 4% paraformaldehyde-fixed spermatogonial membranes. 17. It is not necessary to rinse cultures with PBS before fixation with 4 % paraformaldehyde, or before fixation with methanol. 18. PBS washes should be conducted relatively gently, taking care not to add or remove each PBS wash too vigorously, which can wash colonies off the plate and increase experimental error. Minimize time between PBS washes so as not to allow fixed spermatogonial cultures to air-dry. Air-drying increases background fluorescence. 19. Scoring various-length syncytia is most conveniently done by co-labeling cytoplasmic bridges and nuclei with the TEX14 IgG and Hoechst 33342 dye, respectively (Fig. 1). Syncytia that have advanced to the 2-, 4-, 8-, 16-, and 32-cell steps in

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development will be recognized by TEX14+ foci [34] localized within or adjacent to each cytoplasmic bridge separating nuclei comprising a syncytium (Fig. 1b). It should also be noted that irregular-length syncytia are also observed based on the TEX14 labeling profile, primarily due to a lag in some cells within a given syncytium completing metaphase/anaphase at the time of fixation, which is revealed by nuclear labeling with Hoechst 33342 dye. 20. As, Apr, and Aal spermatogonia represent “undifferentiated” stem and progenitor spermatogonia, originally classified in the rat [5]. Prior to the discovery of proteins that selectively mark distinct spermatogonial types, undifferentiated spermatogonia were distinguished by less heterochromatin, shorter syncytial length, lower abundance, and unique cell division kinetics during an epithelial cycle compared to differentiating spermatogonia (A1–A4, Int, B) [5]. Antibodies generated to protein markers can now be used to distinguish between undifferentiated and more differentiated spermatogonial types [35]. In vivo, As and Apr spermatogonia are more refractory to differentiating into type A1 spermatogonia each epithelial cycle compared to Aal spermatogonia [5]. However, smaller populations of As and Apr spermatogonial fractions are subject to differentiation more directly into A1 spermatogonia each epithelial cycle [5, 22], which has been verified by loss of molecular markers for undifferentiated spermatogonia in vivo as they progress through rodent spermatogenic stages IV–VIII [36, 37]. Similarly, retinoic acid in SD medium effectively drives differentiation in As, Apr, and Aal spermatogonia in vitro, as monitored by loss of molecular markers for undifferentiated spermatogonia (ZBTB16, SALL4) (Fig. 5b) [20], and the inability of rat spermatogonial lines to regenerate spermatogenesis in recipient rat testes following culture in SD medium [20]. Thus, As, Apr, and Aal spermatogenic units are experimentally defined relative to molecular markers used to distinguish between undifferentiated and differentiating spermatogonia. 21. Labeling cultures with molecular probes prior to fixation (i.e., EdU incorporation, fluorescent transgenes/tags) is compatible with procedures described in Subheadings 3.6 and 3.7.

Acknowledgements This work was supported by National Institutes of Health grants from the Eunice Kennedy Shriver National Institute of Child Health and Human Development: R01HD053889 and R01HD061575, and the Office of the Director: R24OD011108.

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References 1. Clermont Y (1972) Kinetics of spermatogenesis in mammals: seminiferous epithelium cycle and spermatogonial renewal. Physiol Rev 52:198–236 2. Brinster RL, Avarbock MR (1994) Germline transmission of donor haplotype following spermatogonial transplantation. Proc Natl Acad Sci U S A 91:11303–11307 3. Huckins C (1971) The spermatogonial stem cell population in adult rats. 3. Evidence for a long-cycling population. Cell Tissue Kinet 4:335–349 4. Huckins C (1971) The spermatogonial stem cell population in adult rats. II. A radioautographic analysis of their cell cycle properties. Cell Tissue Kinet 4:313–334 5. Huckins C (1971) The spermatogonial stem cell population in adult rats. I. Their morphology, proliferation and maturation. Anat Rec 169:533–557 6. Nishimune Y, Haneji T, Kitamura Y (1980) The effects of steel mutation on testicular germ cell differentiation. J Cell Physiol 105:137–141 7. Van Pelt AM, De Rooij DG (1990) The origin of the synchronization of the seminiferous epithelium in vitamin A-deficient rats after vitamin A replacement. Biol Reprod 42:677–682 8. Monesi V (1962) Autoradiographic study of DNA synthesis and the cell cycle in spermatogonia and spermatocytes of mouse testis using tritiated thymidine. J Cell Biol 14:1–18 9. Dym M, Fawcett DW (1971) Further observations on the numbers of spermatogonia, spermatocytes, and spermatids connected by intercellular bridges in the mammalian testis. Biol Reprod 4:195–215 10. Nagano M, Avarbock MR, Leonida EB et al (1998) Culture of mouse spermatogonial stem cells. Tissue Cell 30:389–397 11. Brinster RL, Zimmermann JW (1994) Spermatogenesis following male germ-cell transplantation. Proc Natl Acad Sci U S A 91:11298–11302 12. Sato T, Katagiri K, Gohbara A et al (2011) In vitro production of functional sperm in cultured neonatal mouse testes. Nature 471:504–507 13. Sato T, Katagiri K, Kubota Y et al (2013) In vitro sperm production from mouse spermatogonial stem cell lines using an organ culture method. Nat Protoc 8:2098–2104 14. Kanatsu-Shinohara M, Ogonuki N, Inoue K et al (2003) Long-term proliferation in

culture and germline transmission of mouse male germline stem cells. Biol Reprod 69:612–616 15. Kubota H, Avarbock MR, Brinster RL (2004) Growth factors essential for self-renewal and expansion of mouse spermatogonial stem cells. Proc Natl Acad Sci U S A 101:16489–16494 16. Ryu BY, Kubota H, Avarbock MR et al (2005) Conservation of spermatogonial stem cell selfrenewal signaling between mouse and rat. Proc Natl Acad Sci U S A 102:14302–14307 17. Hamra FK, Chapman KM, Nguyen DM et al (2005) Self renewal, expansion, and transfection of rat spermatogonial stem cells in culture. Proc Natl Acad Sci U S A 102:17430–17435 18. Kanatsu-Shinohara M, Muneto T, Lee J et al (2008) Long-term culture of male germline stem cells from hamster testes. Biol Reprod 78:611–617 19. Kanatsu-Shinohara M, Miki H, Inoue K et al (2005) Long-term culture of mouse male germline stem cells under serum-or feederfree conditions. Biol Reprod 72:985–991 20. Chapman KM, Medrano G, Chaudhary J et al (2015) NRG1 and KITL signal downstream of retinoic acid in the germline to support somafree syncytial growth of differentiating spermatogonia. Cell Death Discov 1:15018 21. Wu Z, Falciatori I, Molyneux LA et al (2009) Spermatogonial culture medium: an effective and efficient nutrient mixture for culturing rat spermatogonial stem cells. Biol Reprod 81:77–86 22. Chan F, Oatley MJ, Kaucher AV et al (2014) Functional and molecular features of the Id4+ germline stem cell population in mouse testes. Genes Dev 28:1351–1362 23. Yeh JR, Zhang X, Nagano MC (2012) Indirect effects of Wnt3a/beta-catenin signalling support mouse spermatogonial stem cells in vitro. PLoS One 7, e40002 24. Chapman KM, Medrano GA, Jaichander P et al (2015) Targeted germline modifications in rats using CRISPR/Cas9 and spermatogonial stem cells. Cell Rep 10:1828–1835 25. Chapman KM, Saidley-Alsaadi D, Syvyk AE et al (2011) Spermatogonial stem cell mediated gene transfer. In: Pease S, Saunders T (eds) Advanced Protocols for Animal Transgenesis: An ISTT Manual, Vol. 1. Springer Press, pp 237–266 26. Izsvak Z, Frohlich J, Grabundzija I et al (2010) Generating knockout rats by transposon

Rattus norvegicus Spermatogenesis Colony-Forming Assays mutagenesis in spermatogonial stem cells. Nat Methods 7:443–445 27. Cronkhite JT, Norlander C, Furth JK et al (2005) Male and female germline specific expression of an EGFP reporter gene in a unique strain of transgenic rats. Dev Biol 284:171–183 28. Hamra FK, Gatlin J, Chapman KM et al (2002) Production of transgenic rats by lentiviral transduction of male germ-line stem cells. Proc Natl Acad Sci U S A 99:14931–14936 29. Hamra FK, Chapman KM, Nguyen D et al (2007) Identification of neuregulin as a factor required for formation of aligned spermatogonia. J Biol Chem 282:721–730 30. Hamra FK, Schultz N, Chapman KM et al (2004) Defining the spermatogonial stem cell. Dev Biol 269:393–410 31. Hamra FK, Chapman KM, Wu Z et al (2008) Isolating highly pure rat spermatogonial stem cells in culture. Methods Mol Biol 450:163–179 32. Ivics Z, Izsvak Z, Chapman KM et al (2011) Sleeping Beauty transposon mutagenesis of the

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rat genome in spermatogonial stem cells. Methods 53:356–365 33. Ivics Z, Izsvak Z, Medrano G et al (2011) Sleeping Beauty transposon mutagenesis in rat spermatogonial stem cells. Nat Protoc 6:1521–1535 34. Greenbaum MP, Yan W, Wu MH et al (2006) TEX14 is essential for intercellular bridges and fertility in male mice. Proc Natl Acad Sci U S A 103:4982–4987 35. Phillips BT, Gassei K, Orwig KE (2010) Spermatogonial stem cell regulation and spermatogenesis. Philos Trans R Soc Lond B Biol Sci 365:1663–1678 36. Beumer TL, Roepers-Gajadien HL, Gademan IS et al (2000) Involvement of the D-type cyclins in germ cell proliferation and differentiation in the mouse. Biol Reprod 63:1893–1898 37. Ikami K, Tokue M, Sugimoto R et al (2015) Hierarchical differentiation competence in response to retinoic acid ensures stem cell maintenance during mouse spermatogenesis. Development 142:1582–1592

Chapter 15 Identification of Mouse piRNA Pathway Components Using Anti-MIWI2 Antibodies Takamasa Hirano, Hidetoshi Hasuwa, and Haruhiko Siomi Abstract The mouse testis has served as a popular model system to study a wide range of biological processes, including germ cell development, meiosis, epigenetic changes of chromatin, transposon silencing, and small RNA-mediated epigenetic modifications. PIWI-interacting RNAs (piRNAs) are a class of small RNAs that are almost exclusively expressed in animal gonads. They repress transposons by forming effector complexes with PIWI proteins to maintain genome integrity of the germline. Here we describe detailed procedures of how to produce monoclonal antibodies against a mouse nuclear PIWI protein, MIWI2, which functions in de novo DNA methylation of target transposon loci. We then describe how to use the antibodies to isolate associated complexes and to detect MIWI2 immunohistochemically. Key words piRNA, MIWI2, Monoclonal antibody, Inclusion body, Mouse embryonic testes

1

Introduction Transposable elements (TEs) are major structural components of mammalian genomes [1]. Nearly half of human and mouse genomes are comprised of TEs, mostly retrotransposons. Although recent studies suggest that TEs enhance evolvability by facilitating the rewiring of regulatory networks through their mutagenic character, for the same reason, they pose a major threat to genome integrity [1, 2]. Thus, host species have evolved control mechanisms to restrict the activity of TEs. One such mechanism involves small silencing RNA pathways that act as defense systems in almost all eukaryotes to protect the host genome against the activity of TEs [3]. In animal gonads, the PIWI-interacting RNA (piRNA) pathway mediates TE silencing [4, 5]. piRNAs are produced from single-stranded long transcripts that are transcribed from discrete genomic loci, termed piRNA clusters that are packed with a large number and various types of TEs [6]. Maturation of piRNAs occurs in the cytoplasm, where they are loaded onto germline-specific Argonaute proteins or PIWI proteins to yield effector complexes

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named piRISCs. piRNAs in the complexes direct cleavage of cytoplasmic target TE transcripts by means of base-pairing or induce specific chromatin modifications including DNA and histone methylation on target TE loci to transcriptionally repress them [4, 5]. The mouse genome encodes three distinct PIWI proteins: PIWIL1 (MIWI), PIWIL2 (MILI) and PIWIL4 (MIWI2) [4, 5, 7]. In contrast to many other animals, in which PIWI proteins are expressed in both male and female gonads, the expression of mouse PIWI proteins is rather restricted to testes. Consistent with this, knockout (KO) mice for each PIWI gene show TE de-repression in the testis, defects in spermatogenesis, and male sterility but no discernible phenotype in female mice. Each mouse PIWI protein shows a specific expression pattern during spermatogenesis: MIWI2 is present from 14.5 dpc (days post coitum) to 3 days after birth (P3), and MIWI is expressed from the appearance of pachytene meiotic cells to elongated spermatids, while MILI is detected from 12.4 dpc in embryonic germ cells to the time of postnatal haploid round spermatid appearance [8–11]. Both MIWI and MILI are cytoplasmic and exhibit small RNA-guided endonuclease or Slicer activity to cleave target TE transcripts [12, 13]. In contrast, once loaded with piRNAs, MIWI2 is transported into the nucleus and induces de novo DNA methylation on target TE chromatin loci to transcriptionally silence TEs [11, 14]. Thus, the MIWI2-piRNA pathway is a useful system to study mechanisms that direct DNA methylation on specific chromatin loci in germ cells. In this chapter, we describe the production of anti-MIWI2 monoclonal antibodies for characterizing MIWI2 localization and MIWI2 nuclear complexes. These antibodies can be used to study many aspects of MIWI2-mediated epigenetic modifications, as well as the expression of genes regulated by MIWI2-piRISC complexes.

2

Materials

2.1 Recombinant MIWI2 Protein Isolation

1. pGEX-5  1 vector (GE Healthcare). 2. pET-28a vector (Novagen/Millipore). 3. Miwi2_1nt-F_EcoRI primer: GACGGGCCCGTG-30 . 4. Miwi2_600nt-R_XhoI primer: GAACTGAATGCACACAG-30 .

50 -ATGAATTCATGAGTG50 -ATCTCGAGTTAGAA-

5. Escherichia coli (E. coli): BL21(DE3). 6. Terrific broth (Difco/BD). 7. Isopropyl β-D-1-thiogalactopyranoside (IPTG) (Wako Pure Chemical Industries). 8. E. coli suspension buffer: 1 mM KH2PO4, 1 mM EDTA, and adjust to pH 7.0 with 1 N KOH.

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9. Inclusion body purification buffer: 4 % Triton X-100 in E. coli suspension buffer. 10. TiterMax Gold adjuvant (TiterMax). 11. Digital homogenizer (Branson, Sonifier 250D). 12. Coomassie brilliant blue (CBB) solution: 0.25 % (w/v) CBB R, 50 % (v/v) methanol, and 20 % (v/v) acetic acid. 13. Mice: BALB/cA (CLEA Japan). 2.2 Isolation of MIWI2-Associated piRNAs

1. Dynabeads protein G (Life Technologies/Thermo). 2. DynaMag™-2 Magnet (Life Technologies/Thermo). 3. RIPA Buffer: 50 mM Tris–HCl pH 8.0, 0.1 % sodium dodecyl sulfate (SDS), 1 % (v/v) Triton X-100, 1 mM dithiothreitol (DTT), 150 mM NaCl, 0.5 % (w/v) sodium deoxycholate (see Note 1), 2 μg/ml pepstatin, 2 μg/ml leupeptin and 0.5 % aprotinin. 4. Disposable Pellet Pestle (Kimble Kontes). 5. Nuclease-free Water (Ambion/Thermo). 6. UltraPure Buffer-Saturated Phenol (Ambion/Thermo). 7. Chloroform (Wako Pure Chemical). 8. Pellet Paint NF Co-precipitant (Novagen/Millipore). 9. Alkaline phosphatase, calf intestinal (CIP) (10,000 units/ml) (New England Biolabs (NEB)). 10. NEBuffer 3.1 (NEB). 11. T4 polynucleotide kinase (T4 PNK) (10,000 units/ml) (NEB). 12. T4 PNK buffer (NEB). 13. [γ-32P]ATP (6000 Ci/mmol) (Perkin Elmer). 14. MicroBio Spin-30 columns (BioRad). 15. Gel loading buffer II (Ambion/Thermo). 16. Gel dryer (ATTO). 17. Decade marker (Ambion/Thermo). 18. Image Plates (Fujifilm). 19. BAS2500 BioImage Analyzer (Fujifilm).

2.3 MIWI2 Immunohistochemistry

1. Paraformaldehyde (PFA) (Wako Pure Chemical). 2. Sucrose solutions (15 and 30 % [w/v] in PBS). 3. SCEM (Section-Lab). 4. Cryostat. 5. Target Retrieval Solution (Dako). 6. Block Ace (DS Pharma Biomedical). 7. Alexa Fluor Anti-mouse IgG series (Molecular Probes/ Thermo).

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8. 40 , 6-Diamidino-2-phenylindole (DAPI) solution (Dojindo). 9. PermaFluor aqueous mounting medium (Thermo). 10. Confocal laser microscope.

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Methods

3.1 Preparation of Recombinant MIWI2 Expression Vectors

3.2 Inclusion Body Purification

To construct recombinant protein expression vectors that express GST- and 6  His-tagged N terminal fragments of MIWI2 (1–200 amino acids) (see Note 2), PCR was performed using a Miwi2 cDNA and the Miwi2_1nt-F_EcoRI and Miwi2_600nt-R_XhoI primers (see Subheading 2.1). The amplified fragment was then inserted into EcoRI/XhoI sites of pGEX-5  1 (for a GST-tagged protein) and pET-28a (for a 6  His-tagged protein). These were named pGEX-5  1-Miwi2 (1–200aa) and pET28a-Miwi2 (1–200aa), respectively. 1. Inoculate 3 ml of Terrific broth containing antibiotics (100 μg/ ml ampicillin or 30 μg/ml kanamycin sulfate) with BL21(DE3) containing pGEX-5  1-Miwi2 (1–200aa) or pET28a-Miwi2 (1–200aa). 2. Incubate at 37  C overnight. 3. Add each culture to 50 ml of fresh Terrific broth with appropriate antibiotics. 4. Incubate at 37  C for 6–8 h. 5. Measure the OD600. 6. Dilute each culture into 500 ml of fresh Terrific broth (final OD600 0.2). 7. Incubate at 37  C to OD600 0.8–1.0. 8. Add 500 μl of 300 mM IPTG (final IPTG concentration 0.3 mM). 9. Incubate at 16  C overnight or for 2–4 h at 37  C. 10. Centrifuge at 3500  g for 20 min. 11. Discard medium and resuspend the pellet with 20 ml of E. coli suspension buffer. Sonicate cells on ice with the conditions; 30 % output, total of 15 cycles (on 1 s to off 4 s). 12. Add 20 % Triton X-100 (final concentration 4 %) and rotate for 15 min at 4  C. 13. Centrifuge at 20,000  g for 20 min. 14. Discard supernatant. *Inclusion bodies cannot dissolve in this solution; therefore, the majority of the recombinant MIWI2 protein precipitates as the pellet (see Note 3).

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15. Resuspend the pellet in 20 ml of inclusion body purification buffer by vigorous pipetting. 16. Rotate at room temperature (RT) for 20 min. 17. Centrifuge at 12,000  g for 20 min. 18. Repeat steps 16 and 17 until the color of pellet becomes white or light gray. *Generally these steps 16–18 need to be repeated three to four times. 19. Resuspend the pellet in 20 ml of ddH2O. 20. Rotate at RT for 10 min and centrifuge at 20,000  g for 20 min. 21. Remove supernatant. 22. Repeat step 19 to 21 twice (see Notes 4 and 5). 23. Store the pellet at –80  C until use. 3.3 Immunize Mice with Recombinant MIWI2 Inclusion Bodies

To generate anti-MIWI2 antibodies, immunize mice with the GSTfused N-terminal fragment of MIWI2. The 6  His-fused Nterminal region of MIWI2 is used in the positive selection screening experiments. 1. Combine 1 mg of inclusion bodies containing recombinant MIWI2, 150 μl of PBS, and 150 μl of TiterMax Gold adjuvant. 2. Mix to an emulsion using a syringe. 3. Inject 100 μl of the emulsion into the peritoneal cavity of an 8-week-old female BALB/cA mouse (see Notes 6 and 7). 4. Inject 100 μl of the emulsion into the peritoneal cavity every 2 weeks. 5. When production of specific antibodies is observed (judged by western blot analysis with serum), inject 100 μl of the emulsion at 7 and 3 days before cell fusion (see below).

3.4 Fusion of Myeloma Cells with Immunized Mouse Lymphocytes

1. Dissect the lymph nodes and spleen from the immunized mouse. 2. Fuse lymphocytes with the P3U1 myeloma cells using a standard protocol [15, 16]. 3. Perform the ELISA screening using the 6  His-taggedMIWI2 and conditioned medium (see Note 8). 4. Perform second and third screens (for example, western blotting, immunoprecipitation, and immunostaining) (Fig. 1).

3.5 Immunoprecipitation of MIWI2 from Mouse Embryonic Testes

1. Collect 40 testes from E14 to P3 mice. *Testes can be stored at –80  C until use (see Note 9). 2. Transfer 30 μl of Dynabeads protein G to a fresh 1.5 ml sterile tube.

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kDa 175 -

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58 46 -

30 anti-β-TUBULIN

Fig. 1 5A2-E5 hybridoma clone producing antibody against MIWI2. Cell lysates from E17.5 mouse testes and NIH/3T3 cells (used as negative control) were analyzed by western blotting using anti-MIWI2 (5A2-E5) and anti-beta-TUBULIN antibodies. The endogenous MIWI2 protein was observed as an 80 kDa band by anti-MIWI2 (5A2-E5) antibody

3. Wash the Dynabeads with 1 ml RIPA buffer. 4. Add 500 μl of RIPA buffer containing 5 μg of mouse antiMIWI2 (5A2-E5) antibody and rotate at 4  C for 30 min. 5. Add 50 μl of ice-cold RIPA buffer to the testes and homogenize on ice using a pestle. 6. Add RIPA buffer up to 500 μl. 7. Pass the lysate through a syringe five times using a G-18 needle. Repeat using a G-25 needle. 8. Centrifuge at 20,000  g at 4  C for 30 min. 9. Transfer the supernatant to a new tube and add RIPA buffer up to 1 ml. 10. Wash beads with immobilized antibody five times in 1 ml RIPA buffer. 11. Add testis lysate to the beads. 12. Rotate at 4  C for 3 h. 13. Wash beads four times with 1 ml of RIPA buffer. 14. Resuspend beads in 1 ml of RIPA buffer and transfer to a new 1.5 ml tube. *To check the immunoprecipitation efficiency, retain an aliquot of beads and subject to western blot analysis. *MIWI2-bound beads can be kept at –80  C after removal of supernatant (see Note 10).

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3.6 Visualization of MIWI2-Associated piRNAs

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1. Remove supernatant from MIWI2-bound beads. 2. Add 150 μl of Nuclease-free Water and 150 μl of buffersaturated phenol (see Note 11). 3. Mix the solution by vortexing for 10 s. 4. Centrifuge at 20,000  g at RT for 5 min. 5. Transfer the upper layer (aqueous fraction) to a fresh tube. 6. Add 150 μl of neutralized phenol solution (see Note 12). 7. Mix by vortexing thoroughly for 10 s. 8. Centrifuge at 20,000  g at RT for 5 min. 9. Transfer the upper layer solution to a fresh tube. 10. Add 500 μl of chloroform and shake the tube vigorously. 11. Transfer the upper layer solution to a fresh tube. 12. Add 15 μl of 3 M NaOAc (pH 5.2), 1 μl of Pellet Paint NF (diluted 1:5), and 600 μl of ice-cold 100 % ethanol (see Note 13). 13. Mix by vortexing thoroughly for 10 s. 14. Centrifuge at 20,000  g at 4  C for at least 30 min. 15. Remove supernatant and add 1 ml of ice-cold 80 % (v/v) ethanol (see Note 14). 16. Centrifuge at 20,000  g at 4  C for 5 min. 17. Carefully remove supernatant. 18. Resuspend the pellet in 5 μl of Nuclease-free Water. 19. Create the following reaction mix to remove the monophosphate group at 50 end of piRNAs. 2 μl RNA solution (from step 17), 5 μl NEBuffer 3.1, 42.5 μl Nuclease-free Water, and 0.5 μl CIP. 20. Incubate at 37  C for 30 min. 21. Perform phenol/chloroform isolation and ethanol precipitation (steps 6–17) and dissolve pellet in 5 μl of Nuclease-free Water. 22. Make the following reaction mix to radioactively label piRNAs: 2 μl CIPed RNA solution (from step 21), 3 μl T4 PNK buffer, 0.5 μl [γ-32P]ATP, 24 μl Nuclease-free Water, and 0.5 μl T4 PNK. 23. Incubate at 37  C for 30 min. 24. Purify the radio-labeled piRNAs using a Micro-Bio Spin column according to the manufacturer’s procedure. 25. Perform phenol/chloroform extraction and ethanol precipitation (step 6–17). 26. Dissolve in 5 μl of Nuclease-free Water. 27. Add 5 μl of gel loading buffer II.

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anti-MIWI2

Non-immune

nt 150 -

MW

IP

100 90 80 70 60 50 40 -

30 -

MIWI2-piRNA

20 -

Fig. 2 MIWI2-associated piRNAs. MIWI2-associated piRNAs were isolated from MIWI2 immunoprecipitants and visualized using 32P radioisotope. Small RNAs from anti-MIWI2 and non-immune mouse IgG immunoprecipitants were resolved on a 12 % acrylamide/6 M urea denaturing gel. piRNAs of approximately 28 nucleotides are observed in the MIWI2 immunoprecipitant lane. The left lane shows molecular weight marker (MW)

28. Heat at 95  C for 1 min. 29. Keep on ice. 30. Separate the RNAs by electrophoresis using a 12 % (w/v) acrylamide/6 M urea-denaturing gel (see Note 15). *Use constant voltage of 250 V. 31. Dry the gel using a gel dryer. 32. Expose the dried gel to an imaging plate and visualize the signals using a BAS2500 image analyzer (Fig. 2). 3.7 MIW2 Immunohistochemistry in Embryonic Testis

1. Fix testes in 4 % (w/v) PFA / PBS at 4  C overnight. 2. Wash in PBS for 5 min three times. 3. Transfer testes to 15 % sucrose solution and incubate at 4  C overnight. 4. Transfer testes to 30 % sucrose solution and incubate at 4  C overnight. 5. Imbed the testes in SCEM compound.

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6. Section the embedded testes at 6 μm using a cryostat and mount on glass slides. 7. Air-dry sections. 8. Wash with PBS at RT for 5 min three times. 9. Boil in the Target Retrieval Solution or 10 mM sodium acetate buffer pH 6.0 using an autoclave (heat to 105  C for 5 min and allow to cool over 1 h) (see Note 16). 10. Wash with PBS three times. 11. Incubate with 2 % Block Ace at 4  C for 1 h. 12. Incubate with first antibody in 0.1 % Triton X-100 / 2 % Block Ace at 4  C overnight. 13. Wash with PBS at RT for 5 min. 14. Repeat wash five times. 15. Incubate with secondary antibody in 0.1 % Triton X-100 / 2 % Block Ace at 4  C for 1 h (see Note 17). 16. Wash with PBS at RT for 5 min. 17. Repeat wash five times. 18. Mount with cover slips and PermaFluor aqueous mounting medium. 19. Observe using fluorescence microscopy (Fig. 3).

a

b

MIWI2

MIWI2 / DAPI

Fig. 3 Immunostaining of embryonic testis with anti-MIWI2 antibody (5A2-E5). (a) MIWI2 protein in gonocytes of the E17.5 testis. White and red indicate MIWI2 and blue indicates DAPI. (b) MIWI2 localizes to the nucleus and to cytoplasmic granules. High-magnification image of (a) is shown. Previous studies have localized MIWI2 to the nucleus and cytoplasmic granules, called piP-bodies [11, 19]. Immunostaining with anti-MIWI2 (5A2E5) is found in both the nucleus and piP-bodies (yellow arrowhead )

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Notes 1. Sodium deoxycholate solution should be used immediately after the powder is dissolved in water. 2. Members of the Argonaute/Piwi family are defined by the presence of PAZ and PIWI domains and they normally consist of one variable N-terminal domain and conserved C-terminal PAZ, MID, and PIWI domains [17, 18]. Thus the variable Nterminal domain should be used to immunize mice to obtain specific monoclonal antibodies that recognize individual PIWI proteins. 3. Please check whether your recombinant protein is solubilized. If your protein is solubilized, tag purification should be used. For proteins that are not solubilized, use this method. 4. Your pellet may be yellowish at first but after repeated rounds of this step, your pellet will become white or grey. 5. To check the purity of recombinant protein, dissolve an inclusion body aliquot in SDS sample buffer and subject it to SDSPAGE followed by CBB staining. 6. Check the reactivity of immunized mice by western blot analysis of serum after the third injection. We recommend aborting the immunization if the titer is not raised after the tenth injection. 7. The 6  His-fused N-terminal fragment of MIWI2 also forms inclusion bodies in E. coli. The inclusion body dissolves in 12 M urea in PBS. After centrifugation at 20,000  g at 4  C for 30 min, dilute the supernatant 1:4 in PBS and use to coat ELISA plates for ELISA screening. 8. Injection of antigen should follow the animal care guidelines of your institute. 9. Testes should be flash-frozen in liquid nitrogen and then stored at –80  C. 10. For piRNA isolation, we recommend phenol/chloroform extraction before storing to avoid RNase contamination. 11. We recommend the use of buffer-saturated phenol. This is because Dynabeads tend to accumulate at the interface between organic and aqueous phases when you use unsaturated phenol. When buffer-saturated phenol is used, Dynabeads tend to be present in the inorganic phase and thus contamination of Dynabeads-bound RNAs can be avoided. 12. We recommend the use of neutralized phenol because the use of acidic phenol can result in failure to purify small RNAs.

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13. After this step, the RNA can be stored at –80  C for several days. 14. Small RNAs with low-GC contents are sometime washed out by 70 % (v/v) ethanol. We recommend the use of 80 % (v/v) ethanol for small RNAs. 15. We recommend pre-electrophoresis of the gel for ~30 min to reduce the gel “smiling effect.” 16. The epitope for the anti-MIWI2 monoclonal antibody is inactivated or masked by formalin fixation. Retrieval steps are required for some antibodies. 17. DAPI can be added to the secondary antibody reaction solution, if co-staining of the nuclei is required.

Acknowledgement The authors would like to thank the members of the Siomi laboratory for their comments and for critical reading of the manuscript. This work was supported by grants from MEXT to H.S. References 1. Bie´mont C, Vieira C (2006) Genetics: junk DNA as an evolutionary force. Nature 443:521–524 2. Lynch M (2007) The origins of genome architecture. Sinauer Associates, Sunderland, MA 3. Slotkin RK, Martienssen R (2007) Transposable elements and the epigenetic regulation of the genome. Nat Rev Genet 8:272–285 4. Siomi MC, Sato K, Pezic D, Aravin AA (2011) PIWI-interacting small RNAs: the vanguard of genome defence. Nat Rev Mol Cell Biol 12:246–258 5. Iwasaki YW, Siomi MC, Siomi H (2015) PIWIinteracting RNA: its biogenesis and functions. Annu Rev Biochem 84:405–433 6. Yamanaka S, Siomi MC, Siomi H (2014) piRNA clusters and open chromatin structure. Mobile DNA 5:22 7. Pillai RS, Chuma S (2012) piRNAs and their involvement in male germline development in mice. Dev Growth Differ 54:78–92 8. Deng W, Lin H (2002) miwi, a murine homolog of piwi, encodes a cytoplasmic protein essential for spermatogenesis. Dev Cell 2:819–830 9. Kuramochi-Miyagawa S, Kimura T, Ijiri TW et al (2004) Mili, a mammalian member of piwi family gene, is essential for spermatogenesis. Development 131:839–849

10. Carmell MA, Girard A, van de Kant HJG et al (2007) MIWI2 is essential for spermatogenesis and repression of transposons in the mouse male germline. Dev Cell 12:503–514 11. Aravin AA, Sachidanandam R, Bourc’his D et al (2008) A piRNA pathway primed by individual transposons is linked to de novo DNA methylation in mice. Mol Cell 31:785–799 12. De Fazio S, Bartonicek N, Di Giacomo M et al (2011) The endonuclease activity of Mili fuels piRNA amplification that silences LINE1 elements. Nature 480:259–263 13. Reuter M, Berninger P, Chuma S et al (2011) Miwi catalysis is required for piRNA amplification-independent LINE1 transposon silencing. Nature 480:264–267 14. Kuramochi-Miyagawa S, Watanabe T, Gotoh K et al (2008) DNA methylation of retrotransposon genes is regulated by Piwi family members MILI and MIWI2 in murine fetal testes. Genes Dev 22:908–917 15. Ishizuka A, Siomi MC, Siomi H (2002) A Drosophila fragile X protein interacts with components of RNAi and ribosomal proteins. Genes Dev 16:2497–2508 16. Hirano T, Iwasaki YW, Lin ZY-C et al (2014) Small RNA profiling and characterization of piRNA clusters in the adult testes of the

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common marmoset, a model primate. RNA 20:1223–1237 17. Carmell MA, Xuan Z, Zhang MQ, Hannon GJ (2002) The Argonaute family: tentacles that reach into RNAi, developmental control, stem cell maintenance, and tumorigenesis. Genes Dev 16:2733–2742

18. Swarts DC, Makarova K, Wang Y et al (2014) The evolutionary journey of Argonaute proteins. Nat Struct Mol Biol 21:743–753 ˜ eda J 19. Aravin AA, van der Heijden GW, Castan et al (2009) Cytoplasmic compartmentalization of the fetal piRNA pathway in mice. PLoS Genet 5:e1000764

Chapter 16 Efficient Induction and Isolation of Human Primordial Germ Cell-Like Cells from Competent Human Pluripotent Stem Cells Naoko Irie and M. Azim Surani Abstract We recently reported a robust and defined culture system for the specification of human primordial germ cell-like cells (hPGCLCs) from human pluripotent stem cells (hPSCs), both embryonic stem cells (ESCs) and induced pluripotent stem cells (iPSCs) in vitro (Irie et al. Cell 160: 253–268, 2015). Similar attempts previously produced hPGCLCs from hPSCs at a very low efficiency, and the resulting cells were not fully characterized. A key step, which facilitated efficient hPGCLC specification from hPSCs, was the induction of a “competent” state for PGC fate via the medium containing a cocktail of four inhibitors. The competency of hPSCs can be maintained indefinitely and interchangeably with the conventional/lowcompetent hPSCs. Specification of hPGCLC occurs following sequential expression of key germ cell fate regulators, notably SOX17 and BLIMP1, as well as initiation of epigenetic resetting over 5 days. The hPGCLCs can be isolated using specific cell surface markers without the need for generating germ cellspecific reporter hPSC lines. This powerful method for the induction and isolation of hPGCLCs can be applied to both hESCs and iPSCs, which can be used for advances in human germ line biology. Key words Human primordial germ cells, Human pluripotent stem cells, Embryonic stem cells, Induced pluripotent stem cells, Primordial germ cell-like cells, Germ cell specification, Epiblasts, Epigenetic resetting

1

Introduction The germ cell lineage is determined very early in embryonic development. Human primordial germ cells (PGCs) are specified in early postimplantation embryos around 2 weeks after fertilization, which makes them inaccessible for research [1]. It is however possible to use human pluripotent stem cells (hPSCs), which are human embryonic stem cells (hESCs) from inner cell mass of blastocysts, and human induced pluripotent stem cells (hiPSCs) from somatic cells for this purpose. These pluripotent cells can be derived and maintained indefinitely in vitro [2, 3], and can potentially provide an in vitro model to study the specification of primordial germ

Michael Buszczak (ed.), Germline Stem Cells, Methods in Molecular Biology, vol. 1463, DOI 10.1007/978-1-4939-4017-2_16, © Springer Science+Business Media New York 2017

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cell-like cells (PGCLCs). Most studies on mammalian early germ cell development have thus far been performed on the mouse model [4, 5]. Analyses of genetically modified mutant mice have revealed that bone morphogenetic protein (BMP) 4 is a critical signaling factor that is necessary to induce nascent PGCs in the epiblast at approximately embryonic day (E) 6.25. Three transcription factors, Prdm1 (encoding BLIMP1), Prdm14, and Tfap2c (encoding AP2γ), are necessary and sufficient for mouse PGC fate [4–7]. An efficient in vitro method for differentiating PGCLCs from mouse PSCs via an epiblast-like cells (EpiLCs) was reported in 2011 [8, 9]. EpiLCs, which are competent for mouse germ line differentiation, are induced in the presence of ActivinA and basic fibroblast growth factor (bFGF) from ground-state mouse PSCs cultured in 2i with leukemia inhibitory factor (LIF) [10]. A significant proportion of day 2 EpiLCs aggregates respond to BMP4, BMP8b, LIF, stem cell factor (SCF), and epidermal growth factor (EGF) and differentiate into PGCLCs after 4 days in culture [8]. Evidence suggests that human PSCs do not respond similarly to mouse PSCs with respect to PGCLC specification. This is because of the differences between mouse and human PSCs in terms of their requirements for propagating the pluripotent state in culture [1, 11, 12]. While mouse PSCs require LIF for supporting pluripotency and acquire dome-shaped colonies, human PSCs require bFGF and have flattened epithelial morphology. There are also differences between mouse and human early postimplantation development at the time of PGC specification. Mouse epiblasts form cup-shaped egg cylinder with extraembryonic ectoderm (ExE) positioned on top. ExE secretes BMP4 to induce PGC precursors in a subpopulation of the epiblast cells. By contrast, human epiblasts are bilaminar flat disc shaped that do not have an equivalent structure to mouse ExE on top of the epiblast [1, 11, 12]. This likely contributes to the differences in the mechanism of PGC specification in mouse and human. Here we describe a newly established, defined, and efficient method to induce human PGCLCs from human PSCs [13]. Based on previous reports, hPSCs cultured in conventional conditions in the presence of bFGF exhibit a low potential for differentiation into PGCLCs [14, 15]. Strikingly, we found that hPSCs cultured in medium containing four inhibitors (4i) for GSK3β, MEK, p38, and JNK display high competency for hPGCLC fate [13, 16]. After our initial attempts with “pre-induction,” we found that hPSCs cultured in 4i medium are already highly competent for hPGCLC fate (hereinafter called “competent hPSCs”) (Fig. 1). The initial studies were carried out on hESCs carrying a germ line-specific reporter NANOS3-mCherry that enabled us to isolate hPGCLCs [13]. Approximately 10–50 % of the NANOS3mCherry-positive hPGCLCs cells are detected amongst the 3D-structured embryoids following a 4–5-day induction with

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low cell attachment U-bottomed plates

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bFGF GSK3βi TGFβ MEKi p38i LIF JNKi Cytokines

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Fig. 1 Overview of hPGCLC induction from hPSCs. Conventional hPSCs are maintained in the presence of bFGF; can be converted to competent condition in a medium which contains bFGF, TGFβ, LIF, GSK3ß inhibitor (i), MEKi, p38i, and JNKi. Seed 2000–4000 competent hPSCs in low cell attachment U-bottomed plates to induce hPGCLC differentiation by adding BMP2 or BMP4, LIF, SCF, EGF, and ROCK inhibitor. A subpopulation of the cells in embryoids express human germ cell specifiers, including SOX17, BLIMP1, and TFAP2C, during specification of hPGCLCs (red circles). Day 5 hPGCLCs show initiation of global epigenetic resetting, NANOS3, and cell surface markers, CD38 and TNAP

BMP2 or BMP4, LIF, SCF, EGF, and ROCK inhibitor (Fig. 1). The hPGCLCs show the global gene expression profile of nascent PGCs, which match with more advanced in vivo hPGCs, and a seminoma cell line, which has key characteristics of early hPGCs. Moreover, hPGCLCs exhibit the expression of key epigenetic regulators and initiation of global epigenetic resetting around days 4–5 of induction [13, 17]. Notably, the hPGCLC induction system led to the identification of SOX17 as a novel key regulator of hPGCLC specification, which is best known for the induction of definitive endoderm fate. Downstream of SOX17 is BLIMP1, and these two genes are critical for the human PGCLC fate. We also found CD38 as a new human germ line-specific cell surface marker, which is expressed on NANOS3-positive hPGCLCs, in vivo hPGCs, and seminoma. Although tissue-nonspecific alkaline phosphatase (TNAP) activity is a conserved marker for PGCs, we were able to detect it on the cell surface and use it to specifically isolate hPGCLCs. Using the combination of CD38 and TNAP, we are able to isolate hPGCLCs, in vivo hPGCs, and seminoma cells without the need for any reporters. Alternative protocols to induce hPGCLCs were published subsequently [18, 19]. Sugawa et al. performed pre-induction with BMP4, ActivinA, and bFGF from conventional hPSCs followed by hPGCLC induction in the presence of BMP4 and LIF. They used cell surface markers KIT and TRA-1-81 to isolate hPGCLC

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populations. Sasaki et al. obtained hPGCLCs induced from hPSCs cultured in a pre-formulated commercial medium [20] and isolated with cell surface markers, EpCAM and INTEGRINα6. The efficiency of hPGCLCs increases from approximately 20–60 % when the hPSCs in the commercial medium are pre-induced into incipient mesoderm-like cells (iMeLCs) with ActivinA and WNT signaling agonist CHIR99021. The iMeLCs however retain competency for hPGCLCs only transiently [19]. The hPGCLCs obtained using our method and that described by Sasaki et al. are transcriptionally similar, while the procedure by Sugawa et al. shows some differentially expressed genes in their hPGCLCs [19]. Our germ lineagecompetent hPSCs maintain their competency, potentially indefinitely, while they undergo self-renewal [13]. Here we describe our procedure for hPGCLC differentiation and isolation from competent hPSCs as well as for the induction and maintenance of competent hPSCs. We believe that this method will greatly facilitate knowledge of human germ line biology and properties, and the causes of infertility and germ cell tumors.

2

Materials

2.1 Reagents and Equipment

1. Feeder cell medium: DMEM high glucose supplemented with 10 % (vol/vol) fetal bovine serum (FBS), 100 units/ml penicillin, 100 μg/ml streptomycin, and 2 mM L-glutamine. Filtersterilize the medium by using a membrane filter (0.22 μm pore size). 2. Conventional hPSC medium: DMEM/F12 þ GlutaMAX or DMEM/F12 with 2 mM L-glutamine added, supplemented with 20 % (vol/vol) knockout serum replacement (KSR) (see Note 1), 0.1 mM MEM non-essential amino acid solution, 0.1 mM 2-mercaptoethanol, and 10–20 ng/ml of bFGF (see Note 2). bFGF should be added freshly prior to use. Filter-sterilize the medium by using a membrane filter (0.22 μm pore size). 3. Conventional hPSC dissociation medium: Dissolve 10 mg/ml of dispase II (Thermo Fisher Scientific) in phosphate-buffered saline (PBS), filter through 0.22 μm membrane, and store at 20  C. Final working concentration is approximately 0.6–2.4 units/ml with 1:10 dilution. 4. 10 mM ROCK inhibitor (Y-27632 dihydrochloride) (TOCRIS bioscience) stock solution in sterile water. Store at 20  C. Final working concentration is 10 μM. 5. Competent hPSC medium: Knockout DMEM supplemented with 20 % (vol/vol) KSR (see Note 1), 100 units/ml penicillin, 100 μg/ml streptomycin, 2 mM L-glutamine, 0.1 mM nonessential amino acids, 0.1 mM 2-mercaptoethanol, 20 ng/ml human LIF, 8 ng/ml bFGF, 1 ng/ml TGF-β1, 3 μM

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CHIR99021, 1 μM PD0325901, 5 μM SB203580, and 5 μM SP600125. After adding all the components, filter the medium through 0.22 μm membrane bottle-top filters (see Note 2). 6. hPGCLC medium: Glasgow’s MEM (GMEM) supplemented with 100 U/ml penicillin, 100 μg/ml streptomycin, 2 mM Lglutamine, 0.1 mM nonessential amino acids, 0.1 mM 2mercaptoethanol, and 1 mM sodium pyruvate and store at 4  C. Freshly add 15 % (vol/vol) KSR (membrane filtered, pore size 0.22 μm) and the following cytokines: 500 ng/ml BMP4 or BMP2 (batch tested for hPGCLC differentiation, see Note 3), 1 μg/ml human LIF, 100 ng/ml SCF, 50 ng/ml EGF, and 10 μM ROCK inhibitor. 7. 0.1 % Gelatin in sterile water. 8. 6-Well Nunclon Delta surface plates. 9. Irradiated mouse embryonic fibroblasts (MEFs) (GlobalStem). 10. 15 ml Conical centrifuge tubes. 11. TrypLE Express (Thermo Fisher Scientific). 12. CellTrics filters (Sysmex). 13. Ultra-low cell attachment U-bottom 96-well plates (Corning). 14. Anti-TNAP antibody (BD Pharmingen) or anti-CD38 antibody (BioLegend) conjugated with fluorochrome of choice. 15. FACS solution: 3 % (v/v) FBS in PBS. 16. 40 ,6-Diamidino-2-phenylindole (DAPI): 5 mg/ml DAPI stock solution in sterile water. Final working concentration is 1 μg/ ml with 1:5000 dilutions. 17. Flow cytometer (e.g., BD LSRFortessa (BD Biosciences) or CyAn ADP analyzer (Beckman Coulter)).

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Methods

3.1 Maintenance of Conventional and Competent hPSCs

1. Preparation of feeder cells for conventional and competent hPSCs culture: Coat 6-well Nunclon Delta surface plates with 1 ml of 0.1 % gelatin into each well for 10–15 min at room temperature. Plate 4–5 million irradiated mouse embryonic fibroblasts (MEFs) (GlobalStem) in feeder cell media into 12–18 wells of 6-well plates, and incubate at 37  C, 5 % CO2, for at least 5 h. If the feeder cells cannot be used immediately, change the medium after 24 h and repeat every other day. Feeder cells can be used up to 4 days after thawing. 2. Maintenance of conventional hPSCs: Seed conventional hPSCs on the feeder cells in 2 ml of conventional hPSC medium with 10 μM ROCK inhibitor. Replace conventional media every day and passage the cells every 4–6 days when the colonies become 70–80 % confluent. Passage conventional hPSCs by adding

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1:10 volume of the conventional hPSC dissociation medium in the cell culture medium (200 μl in 2 ml) and incubate at 37  C, 5 % CO2, for approximately 20–30 min till the colonies have detached. Observe the colonies under the microscope every 5 min and swirl the plates during the incubation to help the colonies detach if necessary. Collect and wash the colonies in 15 ml conical centrifuge tubes with PBS twice with centrifugation at 200  g for 5 min. Pipet gently with P1000 pipetman to break the colonies into 0.5–1 mm in PBS. Plate between 1/5 and 1/10 of the cells using fresh conventional hPSC medium with the ROCK inhibitor onto feeder cells. 3. Maintenance of competent hPSCs: Culture competent hPSCs on feeder cells in 2 ml of competent medium in 6-well plates. Replace competent media every day and passage the cells every 4–6 days when the colonies become 80–90 % confluent. Wash competent hPSCs with PBS once and dissociate with TrypLE Express for 1–2 min or longer at 37  C in 5 % CO2. Wash the cells with feeder cell medium in 15 ml conical centrifuge tubes with centrifugation at 200  g for 5 min and plate 1/5–1/10 of the cells (see Note 4) onto feeder cells in fresh competent medium with 10 μM of ROCK inhibitor. 3.2 Conversion of Conventional and Competent hPSCs

Conventional and competent hPSC states are reversible, which requires at least 2 weeks and for 2–3 passages for the cells to adapt to their new state. The conventional and competent cells have distinct morphologies (Fig. 2) and growth rates [16]. 1. To induce competent hPSCs from conventional hPSCs, replace the conventional medium with the competent medium, at approximately 24 h after passaging conventional hPSCs with conventional hPSC dissociation medium. The competent media should be replaced every day. When colonies become

Fig. 2 Differential interference contrast images of conventional and competent hESCs. WIS2 hESC line was cultured for more than 14 days in conventional (a) and competent (b) hPSC conditions

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70–80 % confluent after approximately 3–4 days, passage the colonies with TrypLE Express, and seed 1/6–1/10 of the cells in fresh competent medium with 10 μM ROCK inhibitor onto feeder cells. Repeat the process for at least 2 weeks and for 2–3 passages. These competent hPSCs can be used thereafter for hPGCLC induction. 2. To prepare conventional hPSCs from competent hPSCs, replace the competent medium with conventional medium at approximately 24 h after passaging competent hPSCs. Replace with fresh conventional medium every day and passage the colonies with conventional hPSC dissociation medium until the colonies achieve 70–80 % confluency. Plate 1/6–1/10 of these cells onto feeder cells, using fresh conventional medium containing 10 μM ROCK inhibitor. Continue the process for at least 2 weeks and passage the cells 2–3 times to obtain conventional hPSCs. 3.3 Induction and Isolation of hPGCLCs

1. Dissociate 80–90 % confluent competent hPSCs with TrypLE Express for 1 min at 37  C, 5 % CO2. Inactivate and dilute TrypLE Express with feeder cell medium and filter dissociated cells with 50 μm sterile single-pack CellTrics filters in 15 ml conical centrifuge tubes. Centrifuge at 200  g for 5 min and resuspend the cells in hPGCLC medium. Count the number of cells (see Note 5). Plate 2000–4000 cells in 100–200 μl hPGCLC medium per well into ultra-low cell attachment Ubottom 96-well plates. Centrifuge the plate at 200  g for 2 min to ensure that the embryoids are well formed. Alternatively, low cell attachment V-bottomed 96-well plates (MoBiTec GmbH) can be used to promote the formation of embryoids. The embryoids can be allowed to develop without the necessity for changing the medium for 5 days. 2. Collect day 4 or day 5 embryoids into 15 ml conical centrifuge tubes using wide-bore pipette tips (see Note 6). To dissociate day 4 embryoids, wash them once with PBS in a centrifuge at 200  g for 1 min. Dissociate with TrypLE Express for 8–20 min at 37  C by pipetting every 5 min. For day 5 embryoids, pretreat with 0.38 mg/ml EGTA and 10 mg/ml PVA in PBS on ice for 10 min and wash with PVA-PBS before dissociation with TrypLE Express (see Note 7). Alternatively, use trypsin-EDTA (0.25 %) for dissociating day 1–5 embryoids. Dilute the dissociated cells in feeder cell medium and centrifuge at 400  g, 5–10 min. Resuspend the dissociated cells in 50 μl of FACS solution consisting of 3 % (v/v) FBS in PBS. 3. Incubate the cells on ice for 30–60 min with 5 μl of anti-TNAP antibody (BD Pharmingen) and/or 5 μl of anti-CD38 antibody (BioLegend) conjugated with fluorochrome of choice depending on the fluorescence indicator in the cell lines if present, and in accordance with the detection setup of the flow cytometer (see Notes 8 and 9). After the incubation, wash and centrifuge the

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stained cells in FACS solution at 400  g for 8–12 min. Resuspend the cells in 350–500 μl FACS solution with 1 μg/ml DAPI. Analyze by flow cytometers such as BD LSRFortessa (BD Biosciences) and CyAn ADP analyzer (Beckman Coulter) or sort by cell sorter, such as MoFlo (Beckman Coulter) and S3 Cell Sorter (Bio-Rad). Gating strategies to detect and analyze the hPGCLC population are shown in Fig. 3. a

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Fig. 3 Gating strategy for FACS analysis to detect hPGCLC population. Side scatter and forward scatter plot was used for detecting the intact cell population, and the population was expanded in the side scatter area and side scatter width plot to select single cells, followed by elimination of dead cells showing high DAPI intensity. The live cell populations were analyzed for NANOS3-mCherry, TNAP and CD38 for NANOS3-mCherry reporter hPSCs (a), and TNAP and/or CD38 for wild-type hPSCs (b) to determine the percentage of induced hPGCLCs

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Notes 1. KSR stocks: Frozen stocks are thawed at 4  C but not for longer than 24 h. Make aliquots for single use only, and keep them at 20  C. Do not warm KSR to 37  C. 2. It is recommended to prepare aliquots for use for approximately 1 week. These can be kept at 4  C. Warm up the media to room temperature before use; avoid warming the media repeatedly. 3. The efficiency of hPGCLC induction differs when using commercially available BMP2 and BMP4. These variations are dependent on the batch and source of the cytokines. It is recommended to perform batch tests for BMP2 and BMP4. 4. Competent hPSCs are affected by and sensitive to cell density. We recommend plating cells at different densities to obtain the best condition for competent hPSCs. 5. One well of a 6-well plate of 80–90 % confluent competent hPSCs usually has approximately 1–2 million cells. These are sufficient for plating into 250–500 wells with 4000 cells each of ultra-low cell attachment U-bottom 96-well plates. Since there is significant cell death at the beginning of hPGCLC induction, the numbers of cells in individual day 4 embryoids are on average between 4000 and 5000 cells. The hPGCLC number on day 4 is around 800–1000, assuming an induction efficiency of around 20 %. 6. FACS analysis during each stage of differentiation can be performed using the following number of embryoids: 18 embryoids for day 1, 15 embryoids for day 2, 12 embryoids for day 3, 9 embryoids for day 4, and 6 embryoids for day 5. These are usually enough to collect the data for 10,000 live single-cell populations by FACS analyzers. 7. Embryoids around day 5 of the hPGCLC induction display a rich extracellular matrix structure. As a result, they are very difficult to dissociate using TrypLE Express. Dissociation of embryoids is even harder beyond 5 days, even with the pretreatment with EGTA-PVA in PBS. We have observed that the hPGCLCs do not show any further differentiation with continued culture after 5–6 days. 8. Expression of cell surface TNAP can be detected in competent hPSCs (Fig. 3). Most of the cells in day 1–2 embryoids still express cell surface TNAP; however it starts to decrease in the non-hPGCLC population around day 3 [13]. On the other hand CD38, which is not expressed in competent hPSCs, starts to be expressed in a subset of day 4 NANOS3-positive hPGCLCs, and the detection increases to a greater number of day 5 hPGCLCs. Therefore, the combination of cell surface markers TNAP and CD38 is suitable for the detection and isolation of late NANOS3-positive hPGCLCs [13].

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9. Since our reporter hPSC line has NANOS3-mCherry and GFP transgenes, we used PerCP-Cy5.5 and Alexa Fluor 647conjugated antibodies and DAPI for eliminating dead cells.

Acknowledgement This work was supported by a Wellcome Trust Investigator Award, and by a BIRAX Grant. We thank Dr. Toshihiro Kobayashi, Dr. Roopsha Sengupta, and Dr. Carlos le Sage for critical reading of the manuscript. The institute is funded by a core grant from the Wellcome Trust (092096) and Cancer Research UK (C6946/A14492). References 1. De Felici M (2013) Origin, migration, and proliferation of human primordial germ cells. In: Coticchio G, Albertini DF, De Santis L (eds) Oogenesis. Springer, London, pp 19–37 2. Thomson JA, Itskovitz-Eldor J, Shapiro SS et al (1998) Embryonic stem cell lines derived from human blastocysts. Science 282:1145–1147 3. Takahashi K, Tanabe K, Ohnuki M et al (2007) Induction of pluripotent stem cells from adult human fibroblasts by defined factors. Cell 131:861–872 4. Hayashi K, de Sousa Lopes SMC, Surani MA (2007) Germ cell specification in mice. Science 316:394–396 5. Leitch HG, Tang WW, Surani MA (2013) Primordial germ-cell development and epigenetic reprogramming in mammals. Curr Top Dev Biol 104:149–187 6. Nakaki F, Hayashi K, Ohta H et al (2013) Induction of mouse germ-cell fate by transcription factors in vitro. Nature 501:222–226 7. Magnu´sdo´ttir E, Dietmann S, Murakami K et al (2013) A tripartite transcription factor network regulates primordial germ cell specification in mice. Nat Cell Biol 15:905–915 8. Hayashi K, Ohta H, Kurimoto K et al (2011) Reconstitution of the mouse germ cell specification pathway in culture by pluripotent stem cells. Cell 146:519–532 9. Hayashi K, Ogushi S, Kurimoto K et al (2012) Offspring from oocytes derived from in vitro primordial germ cell-like cells in mice. Science 338:971–975 10. Ying Q-L, Wray J, Nichols J et al (2008) The ground state of embryonic stem cell selfrenewal. Nature 453:519–523 11. Irie N, Tang WWC, Surani MA (2014) Germ cell specification and pluripotency in mammals:

a perspective from early embryogenesis. Reprod Med Biol 13:203–215 12. Nichols J, Smith A (2012) Pluripotency in the embryo and in culture. Cold Spring Harb Perspect Biol 4:a008128 13. Irie N, Weinberger L, Tang WWC et al (2015) SOX17 is a critical specifier of human primordial germ cell fate. Cell 160:253–268 14. Gkountela S, Li Z, Vincent JJ et al (2013) The ontogeny of cKITþ human primordial germ cells proves to be a resource for human germ line reprogramming, imprint erasure and in vitro differentiation. Nat Cell Biol 15:113–122 15. Kee K, Angeles VT, Flores M et al (2009) Human DAZL, DAZ and BOULE genes modulate primordial germ-cell and haploid gamete formation. Nature 462:222–225 16. Gafni O, Weinberger L, Mansour AA et al (2013) Derivation of novel human ground state naive pluripotent stem cells. Nature 504:282–286 17. Tang WWC, Dietmann S, Irie N et al (2015) A unique gene regulatory network resets the human germline epigenome for development. Cell 161:1453–1467 18. Sugawa F, Arau´zo-Bravo MJ, Yoon J et al (2015) Human primordial germ cell commitment in vitro associates with a unique PRDM14 expression profile. EMBO J 34:1009–1024 19. Sasaki K, Yokobayashi S, Nakamura T et al (2015) Robust in vitro induction of human germ cell fate from pluripotent stem cells. Cell Stem Cell 17:178–194 20. Nakagawa M, Taniguchi Y, Senda S et al (2014) A novel efficient feeder-free culture system for the derivation of human induced pluripotent stem cells. Sci Rep 4:3594

INDEX A Anesthesia ............................................54, 65, 67, 97, 165 Antibody ........................... 5, 8, 9, 15, 25, 26, 40, 43, 45, 51, 53, 54, 57, 58, 105, 129, 130, 132, 135, 136, 141, 143, 144, 147, 149–152, 163, 187, 192, 193, 197, 198, 200, 210, 213, 215, 221, 223 Antibody staining..........15, 25, 106, 109, 110, 130, 163 A single-cell .............. 161, 163, 165, 177, 179, 181, 182 Asymmetric cell division (ACD) ........50, 51, 57, 59, 139

Dissection ................................................................ 16, 25, 53, 54, 57, 58, 65, 67–69, 72, 77, 78, 80, 86–90, 95–97, 100, 106, 108, 110, 119, 122, 129, 132, 134, 142, 161, 165, 167, 174–177, 179, 181, 209 Dissociation .................................................. 81, 122, 140, 174, 175, 181, 182, 187, 220, 222, 223, 225 Distal tip cell (DTC)...................................2–5, 8–11, 13, 14, 23, 25, 26 Drosophila melanogaster.................................................. 75

E B Bone morphogenetic protein (BMP) .........................193, 218, 219, 221, 225 BrdU ................................................................................ 15

C Caenorhabditis elegans (C. elegans)..................... 2–26, 38 CCR4-NOT deadenylation complex .......................93, 94 Cell culture .........................................175, 176, 180–182, 189, 191, 194, 196, 222 Cell cycle..........................................................3, 4, 11–13, 15–17, 19, 20, 22, 26, 53, 54, 58, 72, 186 Cell isolation................ 58, 115–124, 173–177, 179–182 Cell transplantation.............................140, 155–169, 189 Centrosome orientation .................................... 53, 54, 59 Centrosome orientation checkpoint (COC) ...........54, 56 Clonal analysis ............................................................... 134 COC. See Centrosome orientation checkpoint (COC) Confocal microscopy .............................................. 50, 53, 58, 60, 66, 67, 85–90, 106, 111, 120, 121, 130, 147, 148, 151, 152 Cre. See Cre/lox Cre/loxp.......................................................................... 20 CreER .......................................................... 125, 126, 133 Cystoblasts........................................................ 94–97, 100

D Danio rerio. See Zebrafish Deadenylation ...........................................................93, 94 Differentiation....................................................... 2–5, 11, 20, 22, 38, 42, 59, 85, 94, 115, 126, 128, 130, 131, 175, 186–189, 191, 194, 196, 197, 200, 201, 218–221, 225

EdU ........................................... 5, 13, 15–19, 26, 37–39, 41, 42, 45, 201 Embryo ..................................................... 3, 93, 115–124, 176, 177, 179, 181, 217 Embryonic stem cells (ESCs) .....................................3, 51 Epiblasts......................................................................... 218 Epigenetic resetting ...................................................... 219 Erbb3 ........................................................... 140, 146, 195

F Fibroblast growth factor (Fgf) ................... 176, 186, 188 Fixation .........................................................................8, 9, 12, 13, 16, 24, 25, 39, 41, 59, 60, 104, 106, 108, 132, 137, 143, 147–149, 151, 167, 200, 201, 215 Flp. See Flp/Frt Flp/Frt ............................................................... 20, 57, 58 Fluorescence-activated cell sorting (FACS)................149, 163, 164, 169, 174, 176–178, 221, 223–225 Frog. See Xenopus Frozen sections....................................130, 134, 135, 149 Fusion protein ............................................................... 209

G Gal4............................................................. 56–58, 60, 89, 90, 96, 97, 100 β-Galactosidase ..................................................... 147, 148 GBs. See Gonialblasts (GBs) Germ cell .............................................. 1, 2, 9–12, 18, 22, 26, 37, 38, 52, 57, 59, 60, 103, 116, 127, 128, 130, 133, 136, 140, 144, 148, 150, 152, 159, 164, 173–175, 177, 178, 180–182, 186, 189, 217, 219, 220

Michael Buszczak (ed.), Germline Stem Cells, Methods in Molecular Biology, vol. 1463, DOI 10.1007/978-1-4939-4017-2, © Springer Science+Business Media New York 2017

227

GERMLINE STEM CELLS 228 Index Germline stem cells (GSCs) ............................... 2–26, 38, 42, 49, 51–54, 57–60, 63, 64, 70, 71, 75, 85, 86, 89, 90, 93–101, 103–112, 127, 128, 130, 155–169, 185, 189 Germ plasm .........................................116, 118–120, 122 GFP5, 11–14, 20, 25, 26, 56–60, 70, 72, 73, 86, 89, 90, 129, 149, 167, 177, 178, 226 Glial cell line-derived neurotrophic factor (GDNF) .......................... 186–188, 192, 196, 199 Gonad adult ..................................................... 1, 10, 106, 108 fetal ....................................... 127, 136, 137, 173, 174 larval........................................................................... 13 Gonialblast (GB) ............................ 49, 51, 54, 56, 57, 64

H Human.................................................... 3, 35, 37–45, 51, 87, 151, 173, 205, 217–225

I 4i hPSC medium ..........................................218–223, 225 Immunofluorescence ................................. 51, 53, 54, 57, 58, 104–106, 140, 144, 147, 152, 197, 198 Inclusion body............................................. 208, 209, 214 Induced pluripotent stem cells (iPS) human ...................................................................... 217 In situ hybridization .................................. 11, 23, 38–43, 45, 46, 104–109

Monoclonal antibody............................... 5, 26, 141, 151, 206, 214, 215 Mouse .......................................... 5, 15, 50, 52, 125–128, 131, 133, 134, 137, 140, 141, 143, 145–151, 156, 157, 159, 160, 164, 169, 173–175, 180, 186, 187, 191, 192, 195, 197, 205–207, 209, 210, 212, 218, 221 Mouse embryonic fibroblast (MEF) feeder cells................................................. 195, 196 mRNA.......................................23, 46, 93–101, 111, 117 mRNA translation .....................................................93, 94 Mutation............................................................... 159, 194

N Nanos........................................38, 89, 94, 103, 116, 118 Nanos2.................................................103–107, 110, 111 Nanos3.................................................218, 219, 224, 225 Nematode. See Caenorhabditis elegans (C. elegans) Niche................................................ 3, 13, 20, 22, 49, 57, 63–67, 69, 71–73, 75, 76, 85, 100 Notch ...................................................3, 4, 11–13, 22, 23

O Oocyte ...................................................... 2–5, 11, 13, 24, 26, 37, 38, 85, 93, 106, 110, 111, 127, 128, 133–135 Ovary .................................................................37, 38, 64, 75, 85, 103, 131, 133, 134, 137

J

P

JAK-STAT........................................................................ 57

Parasite................................................................ 35, 37–45 Pax7+ ..................................................................... 139–152 Percoll gradient .................................................... 163, 164 Phenotype...................................20, 22, 26, 46, 195, 206 PIWI ............................................................ 205, 206, 214 PIWI-interacting RNA (piRNA)...................76, 205–215 Pluripotency .................................................................. 218 Pluripotent stem cells human ............................................................. 217–225 Poly(A) tail ..............................................................93–101 Poly(A) test (PAT) assays..........................................94, 96 Polyadenylation ............................................................... 93 Polymerase chain reaction (PCR) ................................. 39, 41–43, 45, 76, 82, 95, 99, 105, 106, 112, 129, 181, 208 Prdm1 (encoding BLIMP1)......................................... 218 Prdm14 .......................................................................... 218 Primordial germ cell-like cells (PGCLCs) .......... 217–225 Primordial germ cells (PGCs) ....................................... 75, 115–124, 128, 131, 136, 159, 174–180, 217–219 Progenitor cells .............................. 3, 12, 23, 24, 50, 126 Pumilio ............................................................................ 94

L Larvae .................................................... 35, 76–78, 80, 88 Lineage tracing....................................3, 63, 72, 139–152 Live-cell imaging .......................................................85–90

M Magnetic activated cell sorting (MACS) ....................161, 163, 164, 174–175, 177–180, 182 Magnetic sorting ........................................ 161, 163, 164, 174–175, 177–180, 182 Maternal-to-zygotic transition (MZT) ........................ 119 Meiosis ............................................... 2, 4, 11, 13, 20, 22, 23, 64, 185, 186 Microinjection .....................................158, 161, 165–167 Migration..............................................38, 46, 63, 80, 82, 116, 119, 139, 159, 174, 175 miRNA.................................................................... 94, 119 Mitosis ................................. 4, 19, 20, 23, 52, 56, 57, 59 MIWI2.................................................................. 205–215

GERMLINE STEM CELLS Index 229 R Rat................................ 52, 140, 164, 186–193, 195–201 Rattus norvegicus. See Rat Recombinant protein ...................................206–208, 214 Reproduction .................................................37, 115, 155 Riboprobe.................................................... 39, 42, 43, 45 RNA isolation............................................................75–82 RNAi ...................................... 20, 23, 43, 44, 57, 94, 165 Rosa 26 reporter ........................................................... 148

S Schistosoma mansoni ......................................... 35, 37–45 Seminiferous tubules.................................. 139, 140, 144, 147, 150, 155, 157, 158, 160, 162, 165–167, 169, 186, 190, 191, 198 Sex maturation ..........................................................37, 38 Somatic cyst stem cells (CySCs) ...............................64, 71 Specification.............................................. 38, 49, 50, 103, 115, 116, 118, 217–219 Sperm .................................................... 2–5, 7, 13, 26, 37, 38, 64, 104, 106, 155, 199 Spermatogenesis .................................................. 139, 140, 148, 155–161, 166, 167, 169, 186–193, 195–201, 206 Spermatogenesis colony forming assay .............. 186–193, 195–201 Spermatogonia ........................................... 3, 56, 64, 104, 139, 140, 144, 146, 148, 150–152, 155–157, 161–164, 167, 185–190, 193, 195–201 Spermatogonial stem cells (SSCs) ...................... 139–152, 156–169, 177, 181, 185, 186, 189–191, 195 Spindle orientation.......................................................... 54

Stem cells ............................................1–4, 20, 75, 85, 94, 103, 115, 126–128, 131, 133, 140, 148, 156, 157, 159, 169, 173, 176, 185, 186, 189, 217

T Tamoxifen (Tmx) .............. 125–133, 135, 136, 146–148 TdTomato ............................................................ 147–149 Testis ....................................... 49, 50, 54, 56, 58, 59, 63, 103, 139, 142, 144, 146, 147, 150, 152, 157, 158, 160, 161, 163–169, 177, 189, 191, 206, 210, 213 Tfap2c (encoding AP2g) ..................................... 218, 219 Time-lapse live-imaging. See Live cell imagining Tmx. See Tamoxifen (Tmx) Transcription .............................................. 12, 13, 22, 23, 39, 41–43, 45, 93–95, 107, 112, 116–118, 122, 159, 205, 206, 218 Tudor ...................................................................... 38, 116

U Unpaired (Upd) .............................................................. 49

V Vasa .................................................38, 50, 52–54, 57–59, 86, 104–106, 111, 116, 118, 133, 150

X Xenopus ................................................................. 115–124

Z Zebrafish ............................................................... 103–112