Plant Embryogenesis: Methods and Protocols (Methods in Molecular Biology, 2122) 1071603418, 9781071603413

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Methods in Molecular Biology 2122

Martin Bayer Editor

Plant Embryogenesis Methods and Protocols

METHODS

IN

MOLECULAR BIOLOGY

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

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

For over 35 years, biological scientists have come to rely on the research protocols and methodologies in the critically acclaimed Methods in Molecular Biology series. The series was the first to introduce the step-by-step protocols approach that has become the standard in all biomedical protocol publishing. Each protocol is provided in readily-reproducible step-bystep fashion, opening with an introductory overview, a list of the materials and reagents needed to complete the experiment, and followed by a detailed procedure that is supported with a helpful notes section offering tips and tricks of the trade as well as troubleshooting advice. These hallmark features were introduced by series editor Dr. John Walker and constitute the key ingredient in each and every volume of the Methods in Molecular Biology series. Tested and trusted, comprehensive and reliable, all protocols from the series are indexed in PubMed.

Plant Embryogenesis Methods and Protocols

Edited by

Martin Bayer Department of Cell Biology, Max Planck Institute for Developmental Biology, Tübingen, Germany

Editor Martin Bayer Department of Cell Biology Max Planck Institute for Developmental Biology Tu¨bingen, Germany

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-0341-3 ISBN 978-1-0716-0342-0 (eBook) https://doi.org/10.1007/978-1-0716-0342-0 © Springer Science+Business Media, LLC, part of Springer Nature 2020, corrected publication 2021 Chapter 7 is licensed under the terms of the Creative Commons Attribution 4.0 International License (http:// creativecommons.org/licenses/by/4.0/). For further details see license information in the chapters. 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, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Humana imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature. The registered company address is: 233 Spring Street, New York, NY 10013, U.S.A.

Preface Plants are sessile organisms and adjust their growth to changing environmental situations. Therefore, most of the body architecture is developed post-embryonically and only a miniature version of the plant seedling is established during embryogenesis. In this process, many different tissue types are generated from a single, undifferentiated cell. This includes the establishment of the stem cell niches that orchestrate post-embryonic growth. Unraveling the molecular mechanisms that drive the embryonic patterning process is therefore crucial for our understanding of plant development in general. The study of early events in plant embryogenesis, however, is experimentally difficult in seed plants as it takes place deeply buried in maternal seed and fruit tissue. Isolation and imaging of embryos is therefore a major technical challenge. Furthermore, all essential tissue types are established at early stages of embryogenesis and in many plant species only very limited amount of tissue is available experimentally. In recent years, the study of plant embryogenesis has greatly benefited from new and improved methods that have been developed specifically to overcome these limitations. In this book, we are presenting an overview of state-of-the-art methods for the study of plant embryogenesis in the model organism Arabidopsis thaliana as well as other model and non-model species. The selection of organisms focuses on seed plants and if possible, several independent methods are included to address a specific question. The first part focuses on genetic screens and mutagenesis in Arabidopsis and maize with general experimental considerations and detailed protocols to obtain embryonic mutants. The second part includes protocols for advanced phenotypic analysis and live imaging. Transcriptional profiling is addressed in the third part of the book and includes a variety of recently developed protocols that should enable the reader to address a wide range of scientific questions. The tiny amount of embryonic tissue that is available experimentally seriously impairs biochemical approaches. Protein–protein interactions are therefore visualized by a FRET-FLIM approach in part four. Part five includes methods that introduce other model and non-model species beyond Arabidopsis thaliana. These protocols can also be used as a starting point for the study of other, so far understudied plant species. The last part of the book finally introduces experimental systems that allow to culture or produce zygotic and somatic embryos in vitro. The emphasis lies on protocols that allow the study of the earliest events of plant embryogenesis rather than on protocols that are developed to produce somatic embryos for plant propagation and transformation. The selection of methods in this book is intended to serve not only as a reference for the work in Arabidopsis thaliana and other model species but also as inspiration for new approaches. I hope that this book will be helpful for the study of embryogenesis in seed plants and can serve as a basis for a lot of exciting reports in the future. ¨ bingen, Germany Tu

Martin Bayer

The original version of this book was revised. The correction to this book is available at https://doi.org/ 10.1007/978-1-0716-0342-0_21

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

PART I

v ix

MUTAGENESIS AND GENETIC SCREENS

1 Genetic Screens to Target Embryo and Endosperm Pathways in Arabidopsis and Maize . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Stewart Gillmor, A. Mark Settles, and Wolfgang Lukowitz 2 EMS Mutagenesis of Arabidopsis Seeds . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Stewart Gillmor and Wolfgang Lukowitz 3 EMS Mutagenesis of Maize Pollen . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Mark Settles

3 15 25

PART II MICROSCOPIC IMAGING AND PHENOTYPIC ANALYSIS OF ARABIDOPSIS EMBRYOS 4 Live-Cell Imaging of Zygotic Intracellular Structures and Early Embryo Pattern Formation in Arabidopsis thaliana . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Minako Ueda, Yusuke Kimata, and Daisuke Kurihara 5 Analyzing Subcellular Reorganization During Early Arabidopsis Embryogenesis Using Fluorescent Markers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Che-Yang Liao and Dolf Weijers 6 From Stained Plant Tissues to Quantitative Cell Segmentation Analysis with MorphoGraphX . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Merijn Kerstens, Soeren Strauss, Richard Smith, and Viola Willemsen

PART III

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49

63

EXPRESSION ANALYSIS AND TRANSCRIPTOMICS

7 Small RNA In Situ Hybridizations on Sections of Arabidopsis Embryos. . . . . . . . Katalin Pa´ldi, Magdalena Mosiolek, and Michael D. Nodine 8 Manual Isolation of Living Early Embryos from Tobacco Seeds. . . . . . . . . . . . . . . Peng Zhao, Xuemei Zhou, Ce Shi, and Meng-xiang Sun 9 Profiling Transcriptomes of Manually Dissected Arabidopsis Embryos . . . . . . . . . Ping Kao and Michael D. Nodine 10 Laser-Assisted Microdissection of Plant Embryos for Transcriptional Profiling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ana Marcela Florez-Rueda, Lucas Waser, and Ueli Grossniklaus 11 Cell Type-Specific Transcriptomics in the Plant Embryo Using an Adapted INTACT Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Joakim Palovaara and Dolf Weijers

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87 101 113

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Contents

Transcriptomic Profiling of the Arabidopsis Embryonic Epidermis Using FANS in Combination with RNAseq . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 151 Daniel Slane, Kenneth W. Berendzen, Janika Wittho¨ft, ¨ rgens and Gerd Ju

PART IV 13

Visualizing Protein Associations in Living Arabidopsis Embryo . . . . . . . . . . . . . . . 167 Yuchen Long, Yvonne Stahl, Stefanie Weidtkamp-Peters, and Ikram Blilou

PART V 14

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PROTEIN INTERACTION STUDIES

EMBRYOGENESIS IN OTHER MODEL AND NON-MODEL SPECIES

Imaging of Embryo Sac and Early Seed Development in Maize after Feulgen Staining . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 191 Kamila Kalinowska, Junyi Chen, and Thomas Dresselhaus Using Giant Scarlet Runner Bean (Phaseolus coccineus) Embryos to Dissect the Early Events in Plant Embryogenesis . . . . . . . . . . . . . . . . . . . . . . . . . 205 Min Chen, Anhthu Q. Bui, and Robert B. Goldberg Microscopical Detection of Cell Death Processes During Scots Pine Zygotic Embryogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 223 Jaana Vuosku and Suvi Sutela

PART VI

IN VITRO SYSTEMS TO STUDY EMBRYOGENESIS

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Regulation of Somatic Embryo Development in Norway Spruce . . . . . . . . . . . . . Sara von Arnold, Tianqing Zhu, Emma Larsson, Daniel Uddenberg, and David Clapham 18 In Vitro Production of Zygotes by Electrofusion of Rice Gametes . . . . . . . . . . . . Md Hassanur Rahman, Erika Toda, and Takashi Okamoto 19 Isolated Microspore Culture in Brassica napus. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Patricia Corral-Martı´nez, Carolina Camacho-Ferna´ndez, and Jose M. Seguı´-Simarro 20 Anther Culture in Eggplant (Solanum melongena L.). . . . . . . . . . . . . . . . . . . . . . . . Antonio Calabuig-Serna, Rosa Porcel, Patricia Corral-Martı´nez, and Jose M. Seguı´-Simarro Correction to: Small RNA In Situ Hybridizations on Sections of Arabidopsis Embryos . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

241

257 269

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C1 295

Contributors KENNETH W. BERENDZEN • Center for Plant Molecular Biology, University of Tu¨bingen, Tu¨bingen, Germany IKRAM BLILOU • King Abdullah University of Science and Technology (KAUST), Biological and Environmental Sciences and Engineering (BESE), Thuwal, Saudi Arabia ANHTHU Q. BUI • Department of Molecular, Cell, and Developmental Biology, University of California, Los Angeles, CA, USA; Inviata Ltd., Research Triangle Park, Morrisville, NC, USA ANTONIO CALABUIG-SERNA • Cell Biology Group—COMAV Institute, Universitat Polite`cnica de Vale`ncia, Valencia, Spain CAROLINA CAMACHO-FERNA´NDEZ • Cell Biology Group—COMAV Institute, Universitat Polite`cnica de Vale`ncia, Valencia, Spain JUNYI CHEN • Cell Biology and Plant Biochemistry, University of Regensburg, Regensburg, Germany; Hubei Key Laboratory of Genetic Regulation and Integrative Biology, School of Life Sciences, Central China Normal University, Wuhan, China MIN CHEN • Department of Molecular, Cell, and Developmental Biology, University of California, Los Angeles, CA, USA DAVID CLAPHAM • Department of Plant Biology and Linnean Center for Plant Biology, Uppsala BioCenter, Swedish University of Agricultural Sciences, Uppsala, Sweden PATRICIA CORRAL-MARTI´NEZ • Cell Biology Group—COMAV Institute, Universitat Polite`cnica de Vale`ncia, Valencia, Spain THOMAS DRESSELHAUS • Cell Biology and Plant Biochemistry, University of Regensburg, Regensburg, Germany ANA MARCELA FLOREZ-RUEDA • Department of Plant and Microbial Biology and Zurich–Basel Plant Science Center, University of Zurich, Zurich, Switzerland C. STEWART GILLMOR • Laboratorio Nacional de Gen
omica para la Biodiversidad (Langebio), Unidad de Gen
omica Avanzada, Centro de Investigaci
on y de Estudios Avanzados del Instituto Polite´cnico Nacional (CINVESTAV-IPN), Irapuato, Mexico ROBERT B. GOLDBERG • Department of Molecular, Cell, and Developmental Biology, University of California, Los Angeles, CA, USA UELI GROSSNIKLAUS • Department of Plant and Microbial Biology and Zurich–Basel Plant Science Center, University of Zurich, Zurich, Switzerland GERD JU¨RGENS • Department of Cell Biology, Max Planck Institute for Developmental Biology, Tu¨bingen, Germany KAMILA KALINOWSKA • Cell Biology and Plant Biochemistry, University of Regensburg, Regensburg, Germany PING KAO • Gregor Mendel Institute (GMI), Austrian Academy of Sciences, Vienna Biocenter (VBC), Vienna, Austria MERIJN KERSTENS • Department of Plant Sciences, Wageningen University and Research, Wageningen, The Netherlands YUSUKE KIMATA • Division of Biological Science, Graduate School of Science, Nagoya University, Nagoya, Japan DAISUKE KURIHARA • Institute of Transformative Bio-Molecules (ITbM), Nagoya University, Nagoya, Japan; JST, PRESTO, Nagoya, Japan

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Contributors

EMMA LARSSON • Department of Plant Biology and Linnean Center for Plant Biology, Uppsala BioCenter, Swedish University of Agricultural Sciences, Uppsala, Sweden CHE-YANG LIAO • Laboratory of Biochemistry, Wageningen University, Wageningen, The Netherlands YUCHEN LONG • Laboratoire Reproduction et De´veloppement des Plantes (RDP), Univ Lyon, ENS de Lyon, UCB Lyon 1, CNRS, INRA, Lyon, France WOLFGANG LUKOWITZ • Department of Plant Biology, University of Georgia, Athens, GA, USA MAGDALENA MOSIOLEK • Gregor Mendel Institute (GMI), Austrian Academy of Sciences, Vienna Biocenter (VBC), Vienna, Austria MICHAEL D. NODINE • Gregor Mendel Institute (GMI), Austrian Academy of Sciences, Vienna Biocenter (VBC), Vienna, Austria TAKASHI OKAMOTO • Department of Biological Sciences, Tokyo Metropolitan University, Tokyo, Japan KATALIN PA´LDI • Gregor Mendel Institute (GMI), Austrian Academy of Sciences, Vienna Biocenter (VBC), Vienna, Austria JOAKIM PALOVAARA • Molecular Genetics, University of Bremen, Bremen, Germany ROSA PORCEL • Cell Biology Group—COMAV Institute, Universitat Polite`cnica de Vale`ncia, Valencia, Spain MD HASSANUR RAHMAN • Department of Biological Sciences, Tokyo Metropolitan University, Tokyo, Japan JOSE M. SEGUI´-SIMARRO • Cell Biology Group—COMAV Institute, Universitat Polite`cnica de Vale`ncia, Valencia, Spain A. MARK SETTLES • Horticultural Sciences Department and Plant Molecular and Cellular Biology Program, University of Florida, Gainesville, FL, USA CE SHI • State Key Laboratory of Hybrid Rice, College of Life Sciences, Wuhan University, Wuhan, China DANIEL SLANE • Department of Cell Biology, Max Planck Institute for Developmental Biology, Tu¨bingen, Germany RICHARD SMITH • Department of Comparative Development and Genetics, Max Planck Institute for Plant Breeding Research, Cologne, Germany YVONNE STAHL • Institute for Developmental Genetics, Heinrich-Heine University, Universit€ atsstr. 1, Du¨sseldorf, Germany SOEREN STRAUSS • Department of Comparative Development and Genetics, Max Planck Institute for Plant Breeding Research, Cologne, Germany MENG-XIANG SUN • State Key Laboratory of Hybrid Rice, College of Life Sciences, Wuhan University, Wuhan, China SUVI SUTELA • Natural Resources Institute Finland (Luke), Helsinki, Finland ERIKA TODA • Department of Biological Sciences, Tokyo Metropolitan University, Tokyo, Japan DANIEL UDDENBERG • Physiological Botany, Department of Organismal Biology and Linnean Center for Plant Biology, Uppsala University, Uppsala, Sweden MINAKO UEDA • Division of Biological Science, Graduate School of Science, Nagoya University, Nagoya, Japan; Institute of Transformative Bio-Molecules (ITbM), Nagoya University, Nagoya, Japan SARA VON ARNOLD • Department of Plant Biology and Linnean Center for Plant Biology, Uppsala BioCenter, Swedish University of Agricultural Sciences, Uppsala, Sweden JAANA VUOSKU • Ecology and Genetics Research Unit, University of Oulu, Oulu, Finland

Contributors

xi

LUCAS WASER • Department of Plant and Microbial Biology and Zurich–Basel Plant Science Center, University of Zurich, Zurich, Switzerland STEFANIE WEIDTKAMP-PETERS • Center for Advanced Imaging, Heinrich Heine University Du¨sseldorf, Universit€ a tsstraße 1, Du¨sseldorf, Germany DOLF WEIJERS • Laboratory of Biochemistry, Wageningen University, Wageningen, The Netherlands VIOLA WILLEMSEN • Department of Plant Sciences, Wageningen University and Research, Wageningen, The Netherlands JANIKA WITTHO¨FT • Department of Cell Biology, Max Planck Institute for Developmental Biology, Tu¨bingen, Germany PENG ZHAO • State Key Laboratory of Hybrid Rice, College of Life Sciences, Wuhan University, Wuhan, China XUEMEI ZHOU • State Key Laboratory of Hybrid Rice, College of Life Sciences, Wuhan University, Wuhan, China TIANQING ZHU • State Key Laboratory of Tree Genetics and Breeding, Chinese Academy of Forestry, Beijing, China

Part I Mutagenesis and Genetic Screens

Chapter 1 Genetic Screens to Target Embryo and Endosperm Pathways in Arabidopsis and Maize C. Stewart Gillmor, A. Mark Settles, and Wolfgang Lukowitz Abstract The major tissue types and stem-cell niches of plants are established during embryogenesis, and thus knowledge of embryo development is essential for a full understanding of plant development. Studies of seed development are also important for human health, because the nutrients stored in both the embryo and endosperm of plant seeds provide an essential part of our diet. Arabidopsis and maize have evolved different types of seeds, opening a range of experimental opportunities. Development of the Arabidopsis embryo follows an almost invariant pattern, while cell division patterns of maize embryos are variable. Embryo–endosperm interactions are also different between the two species: in Arabidopsis, the endosperm is consumed during seed development, while mature maize seeds contain an enormous endosperm. Genetic screens have provided important insights into seed development in both species. In the genomic era, genetic analysis will continue to provide important tools for understanding embryo and endosperm biology in plants, because single gene functional studies can now be integrated with genome-wide information. Here, we lay out important factors to consider when designing genetic screens to identify new genes or to probe known pathways in seed development. We then highlight the technical details of two previous genetic screens that may serve as useful examples for future experiments. Key words Embryo defective, Defective kernel, Seed mutant, Enhancer trap, Reporter, Parent-oforigin effect, Germline sector

1

Introduction Mutational analysis has been spectacularly successful in identifying the molecular determinants underpinning biochemical pathways, cellular structures, and developmental processes. But does the genetic approach still hold value in a post-genomics era? After all, mutant screens for a host of different traits have been performed for more than 40 years, often on a large scale—how many informative mutants can possibly remain undiscovered? We argue that genetic screens remain a valuable explorative tool, in particular when a genetic pathway of interest has been sensitized using chemicals or existing genetic variation, where novel techniques for visualizing a phenotype or a trait of interest are available, or if applied to

Martin Bayer (ed.), Plant Embryogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2122, https://doi.org/10.1007/978-1-0716-0342-0_1, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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nonmodel species. In such circumstances, screens can be a prelude to original and rewarding work. However, they are probably best approached as “high risk, high reward” projects—not all ideas will prove fruitful. It follows that screens should be designed to be as nimble as possible: the first goal should be to determine quickly and with as little effort as possible if informative mutations can be found, so it may be decided early in the process whether to expand or abandon an experiment. Once a pilot screen has yielded promising results, the procedure can be repeated on a larger scale. General guiding principles for designing genetic screens with the two most widely used plant models, maize and Arabidopsis, have been summarized in two excellent reviews [1, 2]. Here, we briefly consider some of the potential difficulties as well as opportunities of such approaches when applied to mutations affecting the embryo and endosperm. At the end of the chapter, we describe the technical details of two such screens: a combined enhancer and fluorescent marker screen for embryo patterning mutants in Arabidopsis, and a screen for mutations with a parent-of-origin effect on seed development in maize. Mutagenesis protocols for both species are outlined in Chapters 2 and 3 of this issue. We hope our discussion encourages researchers in seed biology to explore how this powerful tool may be reimagined to benefit their programs.

2

Designing Genetic Screens for Seed Phenotypes

2.1 Use of Morphological Criteria

Abnormalities in the development, cell biology, or physiology of embryos or the endosperm are often reflected in visibly aberrant mature seeds, making it easy to establish large collections of embryo- or endosperm-defective mutations. From such collections, it is clear that many hundreds of genes can mutate to seed phenotypes (discussed in [3] for Arabidopsis and [4–6] for maize). However, the likely primary defects of embryo- or endospermdefective mutations are rarely obvious, since many of them result in superficially similar phenotypes—for example, about half of the mutant embryos uncovered in a microscopy-based screen with Arabidopsis arrested as globular-shaped structures that lacked obvious distinctive features [7]. Thus, identifying robust and informative traits or phenotypic criteria that can be used to specifically probe a process of interest is arguably the most important step toward designing a successful screen. Maize embryos develop surrounded by a large endosperm, occupying most of the mature seed, and an opaque seed coat. Maize seed mutants are commonly subdivided into three classes: embryo-specific (emb) mutants; endosperm-specific mutants; and defective kernel (dek) mutants, which are typically embryo lethal but also affect endosperm morphology [4, 5, 8]. Due to the difficulty in visualizing maize embryos, only emb mutants with relatively

Genetic Screens in Arabidopsis and Maize

5

drastic changes in embryo morphology have been reported to date, and several of these loci function in the plastids [9–13]. Perhaps not surprisingly, the dek and endosperm-specific mutants characterized so far represent a variety of biological functions [14–17]. Compared to maize, Arabidopsis offers several advantages for morphology-based screens: flowers are self-fertile and on average produce about 50 seeds that develop in relative synchrony—a substantial and relatively homogeneous sample; the siliques of an inflorescence axis provide a natural developmental series; the seed coat is thin and transparent while the endosperm is delicate and short-lived, facilitating whole-mount imaging techniques. Perhaps most importantly, Arabidopsis embryos follow an invariant pattern of cell division and cell expansion during the early stages of embryogenesis [18, 19]. Thus, formative cell divisions or the primordia of specific tissues and organs can often be identified on the basis of cell size, shape, and arrangement, particularly before the heart stage or in the incipient root meristem of older embryos. Morphological criteria have also been used to identify defects in basic cell biological functions, such as cell division and cell expansion [7, 20, 21]. 2.2 Use of Specific Genetic Backgrounds

Potentially great improvements over purely morphological criteria can be gained by incorporating molecular markers for a trait of interest, such as reporters of cell and tissue identity or of cellular structures. In Arabidopsis, several sizeable collections of gene and enhancer trap lines employing fluorescent reporter genes as well as reporter genes visualized by histological staining have been described and made publicly available [22–28]. Establishing transgenic plants harboring molecular reporters for a gene of interest or a specific promoter element is relatively straightforward. That said, relatively few embryo screens that were aided by cell- or tissuespecific markers have been reported (for example refs. 29, 30). Using transgenes or existing genetic variation, specific genetic backgrounds may also be created with the aim of sensitizing a pathway or process of interest. Such backgrounds often enable the recovery of a different spectrum of mutants, for example, enhancer or suppressor mutations that in otherwise wild-type plants would have too small an impact to be noticeable or would be masked by genetic redundancy. Subheading 3 describes a screen for developmental embryo mutants in Arabidopsis that combines a sensitized genetic background as well as a fluorescent marker. It is tempting to mention genome-editing technologies in this context: the custom-designed genetic backgrounds they promise to make available, ranging from targeted manipulations in protein coding sequences to rearrangement of cis-regulatory elements, or insertion of foreign elements, seem like an invitation for novel approaches to designing genetic screens.

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Imaging Embryos

Assessing embryo and endosperm phenotypes will in most cases involve some manual dissection followed by microscopic analysis. Given this slow and labor-intensive process, the cost for recovering a mutation can be high, and effective protocols for preparing and visualizing samples are critically important. In the case of maize, simple methods for visualizing whole-mount seeds have not yet been reported; typically, candidate mutant embryos are dissected from the seed and examined with a dissecting microscope. In Arabidopsis, whole-mount seeds can be imaged in a number of ways. In case only the morphology of embryos is of interest, immature seeds can be embedded in Hoyer’s solution (containing chloral hydrate as a clearing agent [31]) and viewed after a few hours by differential interference contrast (DIC or Nomarski optics; for an example see [7]). While this procedure is fast and simple, chloral hydrate is incompatible with fluorescent proteins and other commonly used fluorophores, posing a significant limitation. Good progress toward developing clearing methods that are compatible with fluorescent microscopy has been made over the past few years, although perhaps no clear favorite has yet emerged. Hydrogels provide excellent optical properties, but sample processing is laborious and time-consuming [32, 33]. Perfluorodecalin readily infiltrates samples containing air spaces, such as leaves, increasing their transparency with minimal impact on physiological processes [34–36]; however, seeds usually don’t contain air spaces. For imaging seeds, clearing with ClearSee or thiodiethanol may be the best currently described options. ClearSee contains xylitol, deoxicholate, and urea as active ingredients [37], an improvement over previous agents containing urea and glycerol [38], and has been used for clearing a wide variety of plant tissues with excellent results [39, 40]. Tissue samples are briefly fixed with formaldehyde and then cleared before microscopy; clearing time depends on the sample and may be up to several days. Thiodiethanol (TDE) can be mixed with water at any ratio to create mounting media with a refractive index matching the sample [41]. Tissue samples are briefly fixed and directly embedded in diluted TDE for microscopy, making for a comparatively rapid procedure (also known as TOMEI; [42–44]). In many cases, clearing may not be necessary at all: young Arabidopsis seeds containing pre-globular embryos are naturally quite transparent, as are older embryos released from their seeds by application of gentle pressure to the cover slip. Musielak et al. [45] recently described a very simple one-step protocol for simultaneously fixing and counterstaining young seeds or older embryos released from the seeds; SR2200 (Renaissance Chemicals) was used to specifically label the cell wall, and the resulting slides can be directly imaged by fluorescence or confocal microscopy.

Genetic Screens in Arabidopsis and Maize

2.4 Screens Involving Crosses

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Plant seeds are mosaics consisting of a maternal seed coat encasing the two products of double fertilization, the triploid endosperm and the diploid embryo. The regulatory networks coordinating these compartments are intricate and only beginning to be understood—for example, gene expression in the endosperm is heavily influenced by maternal imprinting, while many genes expressed in the zygote and early embryo show delayed activation of the paternal allele [46–48]. In addition, seed development can be affected by genes acting in the male or female gametophyte before fertilization. These complex genetic interactions give rise to mutations with specific gametophytic and sporophytic parent-of-origin effects. Genetic screens for parent-of-origin effects often involve mass crosses of mutagenized plants with tester plants of a known genotype. Arabidopsis flowers are hermaphroditic and self-fertilizing. Although male- and female-sterile lines are available to facilitate crossing experiments on a large scale, we are not aware of a screen based on crossing that has been described in the literature. The relative ease of crossing maize may offer a unique experimental advantage in this respect; Subheading 4 describes screening procedure for mutations with a parent-of-origin effect capitalizing on this advantage. Similarly, crossing-based screens can also be used for identifying particular allelic interactions, such as noncomplementation.

3 Example of Screening Protocol: Use of a Fluorescent Marker for Identifying Patterning Defects in Embryos Produced by Mutagenized Arabidopsis Plants In a lucky (but probably not uncommon) coincidence, the enhancer trap insertion E2023 [30, 49] (ABRC stock number CS65893) creates a tissue-specific fluorescent marker combined with a loss-of-function mutation. E2023 harbors a chromosomal inversion that places a green fluorescent protein (GFP) under regulation of the KANADI2 (KAN2) gene, causing GFP expression in the peripheral-abaxial domain of the embryo beginning at the heart stage. The same inversion also creates a kan2 mutant, sensitizing the genetic background for a loss of peripheral-abaxial identity. Thus, mutagenesis of E2023 at once uncovers second-site enhancers of the weak peripheral-abaxial polarity defects conditioned by loss of KAN2 function, as well as factors directly or indirectly affecting the regulation of KAN2 transcription. Two genes identified in a mutant screen with this line have been characterized [30]. Following EMS treatment, the M2 embryos produced by selfpollination of M1 plants grown from mutagenized seed were examined directly for alterations in the E2023 expression pattern and/or morphological alterations. The M1 generation is mosaic, and in the

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main inflorescence 2–3 sectors on average contribute to gamete production [50] (also see Chapter 2 of this issue); several siliques were examined to include all or most of these sectors. As predicted, subsequent siliques on the same inflorescence typically represented different sectors: a mutation affecting seed morphology observed in one silique would be absent from the subsequent 1 or 2 siliques and then reappear again. Recovering a mutation of interest in the following generation was not problematic, suggesting that most sectors are well represented in the mature seed harvested from an M1 plant. A tremendous practical advantage was that E2023 created a surprisingly strong fluorescent signal, likely due to displacement of KAN2 repressor elements by the chromosomal rearrangement. GFP fluorescence was bright enough to be readily observable through the seed coat, which greatly simplified the initial assessment of expression patterns and level. For each M1 plant, three consecutive siliques containing embryos of the bent cotyledon stage were removed and fixed to a microscope slide with doublesided tape. Siliques were then sliced along the valve-septum junction using #5 forceps, and the valve pushed aside to expose the developing seed. Seeds were examined using a dissecting microscope fitted with epifluorescence illumination and a long-pass GFP filter. Under these conditions, red chlorophyll auto-fluorescence outlined the overall morphology of the embryo, while green E2023 fluorescence marked the peripheral domain of hypocotyl and cotyledons (Fig. 1). When deviations for normal morphology or marker expression were noted, embryos were dissected out of the ovules for closer inspection: an incision was made in the bottom of ovule between the hypocotyl and cotyledons, and the embryos pushed out. Heart- and torpedo-stage embryos were visualized in a similar manner to provide a more detailed assessment of the phenotype (Fig. 1). Only M1 plants producing candidate mutants of interest (about 1% of all plants examined) were documented and saved for seed production. The remaining M1 plants were discarded. This screening procedure required that plants be examined during an approximately 2-week window, when they contained seeds of the appropriate ages. To secure a constant supply of M1 plants, a mutagenesis was performed every 2 months. Mutagenized seed were distributed to 20 standard trays fitted with 24-well inserts, and the trays placed in a cold room for up to 4 weeks. Every week 5 trays were transferred to a growth chamber. Ten days after transfer, the flats were weeded to leave four healthy-looking M1 seedling per well. One tray of mutagenized plants could be examined per day (in about 6 h). Thus, a pilot screen of 10 flats containing about 1000 M1 plants (representing about 2000 sectors) was accomplished with 2 weeks’ work.

Genetic Screens in Arabidopsis and Maize

a

b

9

c

*

* * d

M1 plant with two germline sectors

e

wild type

serrate

Fig. 1 Outline of a seed screen with a fluorescent reporter. (a) Schematic drawing of an M1 plant after seed mutagenesis; two different recessive seed mutants arising in two independent germ line sectors after self-pollination are marked in blue and orange. (b–e) The enhancer trap line E2023 (yellow fluorescence) marks the peripheral-abaxial domain of developing embryos (outlined by red chlorophyll fluorescence). (b) Partly opened silique exposing late-stage wildtype and serrate seeds. (c) Wild-type and serrate seeds removed from the silique and viewed with a fluorescence stereo microscope: seeds containing embryos with reduced E2023 expression domains marked by stars. (d) Bent cotyledonstage wild-type and (e) serrate mutant embryos dissected from seeds of the same silique. Scale bars: (a, b) 500 μm, (c, d) 100 μm

4 Example of Screening Protocol: Crossing-Based Schemes for Identifying Parent-of-Origin Effects in Maize Seed Development The maize UniformMu transposon-tagging population was developed by introgressing active Robertson’s Mutator (Mu) transposons into an inbred line (color-converted W22). To avoid the accumulation of Mu elements or induced mutations over time, lines were maintained by continuous backcrossing [51]. In this experiment, Mu-active pollen was crossed to normal inbred plants (W22) to generate M1 kernels. A large number of the kernels were planted, the resulting plants self-pollinated, and the M2 ears screened for defective kernel (dek) mutations. More than 2000 dek mutants were identified in this manner. Mutations with a dominant or a gametophytic parent-of-origin effect are expected to result in ears with 50–75% dek kernels upon self-pollination, but such ears were rarely observed. Most dek mutants recovered from the screen were recessive lethals, segregating 25% dek kernels. The unexpected scarcity of mutants producing 50% or more dek kernels upon self-pollination may be explained by

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x Tester pollen

3

1 Selfpollination

x Tester ear 2

1

Paternal effect

2 Normal 3

Candidate mutant

dek (control)

or

dek Maternal effect

Dominant zygotic effect

Fig. 2 Reciprocal crossing scheme used for uncovering mutations with a dominant zygotic effect or gametophytic parent-of-origin effect (see text for details)

“grain-fill”: ovary and early seed abortion is common in maize inbred lines, and the positions of aborted seeds are filled by sibling neighboring kernels [52]. Alternatively, most dominant- and parent-of-origin-effect mutations in maize may show low penetrance. In order to differentiate between these possibilities, potential parent-of-origin effects of a subset of 146 dek mutants that develop a rough, etched, or pitted endosperm were investigated in detail [53]. This secondary screen was performed in conjunction with recombination mapping of these mutants and relied on reciprocal crosses (Fig. 2). The normal kernels of segregating families were planted for each of the dek mutant isolates. The first or apical ear of the resulting plants was then self-pollinated, while the second ear was crossed with normal inbred pollen (B73 or Mo17). In addition, pollen of each plant was crossed to normal inbred plants (B73 or Mo17). In this experiment, self-pollination served to identify the heterozygous dek plants: only these plants produced dek kernels upon self-pollination, while their wild-type siblings from the same family did not. The reciprocal crosses were then screened for segregation of dek kernels. This analysis unambiguously placed each dek mutant in one of three genetic classes (Fig. 2). Recessive zygotic-effect mutants segregated for dek phenotypes only in self-pollinated ears, but were complemented by wild-type alleles from either parent—the majority of the mutants were in this class. Dominant zygotic-effect mutants would segregate for dek kernels in the self-pollinated ears as well as ears from both reciprocal crosses—however, no such mutants were recovered. Maternal-effect mutants segregated for dek kernels in the self-pollinated ears and in the ears resulting from crosses with inbred pollen. Paternal-effect mutants segregated in the self-pollinated ears and the ears resulting from crosses onto inbred plants. This screen identified 15 candidate parent-of-origineffect mutants, 9 of which showed consistent maternal or paternal effects in subsequent crosses [53]. Perhaps not surprisingly, many

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of the parent-of-origin mutations primarily affected the endosperm. Notably, only two of them produced dek kernels close to the expected 1:1 ratio, indicating that the vast majority of parentof-origin effects likely show relatively low penetrance. Consistent with this conclusion, further analysis of three maternal-effect mutations showed normal transmission through the gametophytic generation. An independent screen for sporophytic maternal mutations and female gametophytic mutations was reported by Chettor et al. [8]. This screen capitalized on the observation that many transposons are most active in the generative cells, causing small mutant sectors that typically encompass only a single gametophyte. Ears of plants harboring active transposons with such properties (UniformMu and Ac/Ds) were crossed with normal pollen. These crosses produced a low number of dek kernels in most ears, due to either environmental factors, aneuploid gametes resulting from an aberrant meiosis, or novel mutations that originated in a single gametophyte. After replanting, about 20% of these dek kernels germinated and produced a seedling. Any resulting fertile plants were then crossed with normal pollen to determine whether the dek phenotype was maternally inherited. Seven gametophytic maternaleffect mutants were isolated from ~750 M1 ears and 566 dek kernels, corresponding to, ~1.2% of the dek kernels initially isolated. Most of these mutations also showed poor transmission through pollen, indicating that the affected genes are required in the gametophytes of both sexes. Gametophytic factors with poor transmission and viability are prone to be lost or underrepresented in genetic screens, suggesting that the decision to limit the analysis to the relatively low number of dek kernels that appeared immediately after mutagenesis and crossing with normal pollen was critical to the success of the experiment.

Acknowledgements This work was supported by grants from CONACYT Mexico (237480 to C.S.G.), NIFA (SCRI-2018-51181-28419 to A.M. S.), the NSF (IOB-04541201 to W.L., IOS-1444456 to A.M.S.), and the Vasil-Monsanto Endowment (to A.M.S.). References 1. Candela H, Hake S (2008) The art and design of genetic screens: maize. Nat Rev Genet 9:192–203 2. Page D, Grossniklaus U (2001) The art and design of genetic screens: Arabidopsis thaliana. Nat Rev Genet 3:124–136

3. Tzafrir I, Pena-Muralla R, Dickerman A, Berg M, Rogers R, Hutchens S, Sweeney TC, McElver J, Aux G, Patton D, Meinke D (2004) Identification of genes required for embryo development in Arabidopsis. Plant Physiol 135:1206–1220

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4. Neuffer MG, Sheridan WF (1980) Defective kernel mutants of maize. I. Genetic and lethality studies. Genetics 95(4):929–944 5. Scanlon MJ, Stinard PS, James MG, Myers AM, Robertson DS (1994) Genetic analysis of 63 mutations affecting maize kernel development isolated from Mutator stocks. Genetics 136(1):281–294 6. Brunelle DC, Clark JK, Sheridan WF (2017) Genetic screening for EMS-induced maize embryo-specific mutants altered in embryo morphogenesis. G3 (Bethesda) 7:3559–3570 7. Gillmor CS, Roeder AHK, Sieber P, Somerville C, Lukowitz W (2016) A genetic screen for mutations affecting cell division in the Arabidopsis thaliana embryo identifies seven loci required for cytokinesis. PLoS One 11:e0146492. https://doi.org/10.1371/jour nal.pone.0146492. 8. Chettoor AM, Phillips AR, Coker CT, Dilkes B, Evans MMS (2016) Maternal gametophyte effects on seed development in maize. Genetics 204:233–248 9. Magnard J-L, Heckel T, Massonneau A, Wisiewski J-P, Cordelier S, Lassagne H, Perez P, Dumas C, Rogowsky PM (2004) Morphogenesis of aize embryos requires ZmPRPL35-1 encoding a plastid ribosomal protein. Plant Physiol 134:649–663 10. Sosso D, Canut M, Genderot G, Dedieu A, Chambier P, Barkan A, Consonni G, Rogowsky PM (2012) PPR8522 encodes a chloroplasttargeted pentatricopeptide repeat protein necessary for aize embryogenesis and vegetative development. J Exp Bot 63:5843–5857 11. Shen Y, Li C, McCarty DR, Meeley R, Tan B-C (2013) Embryo defective 12 encodes the plastid initiation factor 3 and is essential for embryogenesis in maize. Plant J 74:792–804 12. Li C, Shen Y, Meeley R, McCarty D, Tan B-C (2015) Embryo defective 14 encodes a plastidtargeted cGTPase essential for embryogenesis in maize. Plant J 84:785–799 13. Yang J, Suzuki M, McCarty DR (2016) Essential role of conserved DUF177A protein in plastid 23S rRNA accumulation and plant embryogenesis. J Exp Bot 67:5447–5460 14. Gontarek BC, Neelakandan AK, Wu H, Becraft PW (2016) NKD transcription factors are central regulators of maize endosperm development. Plant Cell 28:2916–2936 15. Li C, Yue Y, Chen H, Qi W, Song R (2018) The ZmbZIP22 transcription factor regulates 27-kD γ-Zein gene transcription during maize endosperm development. Plant Cell 30:2402–2424

16. Zhan J, Li G, Ryu C-H, Ma C, Zhang S, Lloyd A et al (2018) Opaque-2 regulates a complex gene network associated with cell differentiation and storage functions of maize endosperm. Plant Cell 30:2425–2446 17. Bai F, Corll J, Shodja DN, Davenport R, Feng G, Mudunkothge J, Brigolin CJ, Martin F, Spielbauer G, Tseung C-W, Siebert A, Barbazuk WB, Lal S, Settles AM (2019) RNA binding motif protein 48 is required for U12 splicing and maize endosperm differentiation. Plant Cell 31:715–733 18. Mansfield SG, Briarty LG (1991) Early embryogenesis in Arabidopsis thaliana II: the developing embryo. Can J Bot 69:461–476 19. Mayer U, Ju¨rgens G (1994) Arabidopsis. In: Brad J (ed) EMBRYOS, color atlas of development. Mosby Year Book Ltd, London 20. Gillmor CS, Poindexter P, Lorieau J, Palcic MM, Somerville C (2002) Alpha-glucosidase I is required for cellulose biosynthesis and morphogenesis in Arabidopsis. J Cell Biol 156:1003–1013 21. Gillmor CS, Lukowitz W, Brininstool G, Sedbrook JC, Hamann T, Poindexter P, Somerville C (2005) Glycosylphosphatidylinositolanchored proteins are required for cell wall synthesis and morphogenesis in Arabidopsis. Plant Cell 17:1128–1140 22. Sundaresan V, Springer P, Volpe T, Haward S, Jones JDG, Dean C, Ma H, Martienssen R (1995) Patterns of gene action in plant development revealed by enhancer trap and gene trap transposable elements. Genes Dev 9:1797–1810 23. Campisi L, Yang Y, Yi Y, Heilig E, Herman B, Cassista J, Allen DW, Xiang H, Jack T (1999) Generation of enhancer trap lines in Arabidopsis and characterization of expression patterns in the inflorescence. Plant J 17:699–707 24. Geisler M, Jablonska B, Springer PS (2002) Enhancer trap expression patterns provide a novel teaching resource. Plant Physiol 130:1747–1753 25. Laplaze L, Parizot B, Baker A, Richaud L, Martiniere A, Auguy F, Franche C, Nussaume L, Bogusz D, Haseloff J (2005) GAL4-GFP enhancer trap lines for genetic maniupulation of lateral root development in Arabidopsis thaliana. J Exp Bot 56:2433–2442 26. Ckurshumova W, Koizumi K, Chatfield SP, Sanchez-Buelna SU, Gangaeva AE, McKenzie R, Berleth T (2009) Tissue-specific GAL4 expression patterns as a resource enabling targeted gene expression, cell typespecific transcript profiling and gene function

Genetic Screens in Arabidopsis and Maize characterization in the Arabidopsis vascular system. Plant Cell Physiol 50:141–150 27. Gardner MJ, Baker AJ, Assie J-M, Poethig RS, Haseloff JP, Webb AR (2009) GAL4 GFP enhancer trap lines for analysis of stomatal guard cell development and gene expression. J Exp Bot 60:213–226 28. Radoeva T, ten Hove CA, Saiga S, Weijers D (2016) Molecular characterization of Arabidopsis GAL4/UAS enhancer trap lines identifies novel cell-type-specific promoters. Plant Physiol 171:1169–1181 29. Jurkuta RJ, Kaplinsky NJ, Spindel JE, Barton MK (2009) Partitioning the apical domain of the Arabidopsis embryo requires the BOBBER1 NudC domain protein. Plant Cell 21:1957–1971 30. Gillmor CS, Park MY, Smith MR, Pepitone R, Kerstetter RA, Poethig RS (2010) The MED12-MED13 module of mediator regulates the timing of embryo patterning in Arabidopsis. Development 137:113–122 31. Anderson LE (1954) Hoyer’s solution as a rapid permanent mounting medium for bryophytes. Bryologist 57:242–244 32. Chung K, Wallace J, Kim S-Y, Kalanasundaram S, Andalman AS, Davidson TJ, Mirzabekov JJ, Zalocusky KA, Mattis J, Denisin AK (2013) Structural and molecular interrogation of intact biological systems. Nature 497:332–337 33. Palmer WM, Martin AP, Flynn JR, Reed SL, White RG, Furbank RT, Grof CPL (2015) PEA-CLARITY: 3D molecular imaging of whole plant organs. Sci Rep 5:13492. https:// doi.org/10.1038/srep13492. 34. Littlejohn GR, Gouveia JD, Edner C, Smirnoff N, Love J (2010) Perfluorodecalin enhances in vivo confocal microscopy resolution of Arabidopsis thaliana mesophyll. New Phytol 186:1018–1025 35. Littlejohn GR, Love J (2012) A simple method for imaging Arabidopsis leaves using perfluorodecalin as an infiltrative medium. J Vis Exp 16:3394. https://doi.org/10.3791/3394 36. Littlejohn GR, Mansfield JC, Christmas JT, Witterick E, Fricker MD, Grant MR, Smirnaoff N, Everson RM, Moger J, Love J (2014) An update: omprovements in imaging perfluorocarbon-mounted plant leaves with implications for studies of plant pathology, physiology, development and cell biology. Front Plant Sci 5:140. https://doi.org/10. 3389/fpls.2015.00140. 37. Kurihara D, Mizuta Y, Sato Y, Higashiyama T (2015) ClearSee: a rapid optical clearing

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reagent for whole-plant fluorescence imaging. Development 142:4168–4179 38. Warner CA, Biedrzycki ML, Jacobs SS, Wisser RJ, Caplan JL, Sherrier DJ (2014) An optical clearing technique for plant tissues allowing deep imaging and compatible with fluorescence microscopy. Plant Physiol 166:1684–1687 39. Ursache R, Andersen TG, Mahavy P, Geldner N (2018) A protocol for combining fluorescent proteins with histological stains for diverse cell wall components. Plant J 93:399–412 40. Mizuta Y, Tsuda K (2018) Three-dimensional multiphoton imaging of transcription factor by ClearSee. In: Yamaguchi N (ed) Plant transcription factors: methods and protocols, methods in molecular biology. Springer, New York, p 1830. https://doi.org/10. 1007/978-1-4939-8657-6_15 41. Staudt T, Lang MC, Medda R, Engelhardt J, Hell SW (2007) 2,20 -thiodiethanol: a new water soluble mounting medium for high resolution optical microscopy. Microsc Res Tech 70:1–9 42. Hasegawa J, Sakamoto Y, Nakagami S, Aida M, Sawa S, Matsunaga S (2016) Threedimensional imaging of plant organs using a simple rapid transparency technique. Plant Cell Physiol 57:462–472 43. Musielak TJ, Slane D, Liebig C, Bayer M (2016) A versatile optical clearing protocol for deep tissue imaging of fluorescent proteins in Arabidopsis thaliana. PLoS One 11:e0161107. https://doi.org/10.1371/journal.pone. 0161107. 44. Slane D, Bu¨rgel P, Bayer M (2017) Staining and clearing o Arabidopsis reproductive tissue for imaging of fluorescent proteins. In: Schmidt A (ed) Plant Germline development: methods and protocols, metholds in molecular biology. Springer, New York, p 1669. https:// doi.org/10.1007/978-1-4939-7286-9_8 45. Musielak TJ, Schenkel L, Kolb M, Henschen A, Bayer M (2015) A simple and versatile cell wall staining protocol to study plant reproduction. Plant Reprod 28:161–169 46. Bai F, Settles AM (2015) Imprinting in plants as a mechanism to generate seed phenotypic diversity. Front Plant Sci 5:780. https://doi. org/10.3389/fpls.2014.00780. 47. Garcia-Aguilar M, Gillmor CS (2015) Zygotic genome activation and imprinting: parent-oforigin gene regulation in plant embryogenesis. Curr Opin Plant Biol 27:29–35 48. Gehring M, Satyaki PR (2017) Endosperm and imprinting, inextricably linked. Plant Physiol 173:143–154

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49. Haseloff J (1999) GFP variants for multispectral imaging of living cells. Methods Cell Biol 58:139–151 50. Redei GP, Koncz C (1992) Classical mutagenesis. In: Koncz C, Chua NH, Schell J (eds) Methods in Arabidopsis research. World Scientific Publishing, Singapore, pp 16–82 51. McCarty DR, Settles AM, Suzuki M, Tan BC, Latshaw S, Porch T, Robin K, Baier J, Avigne W, Lai J, Messing J, Koch KE, Hannah LC (2005) Steady-state transposon mutagenesis in maize. Plant J 44:52–61

52. Gustin JL, Boehlein SK, Shaw JR, Junior W, Settles AM, Webster A, Tracy WF, Hannah LC (2018) Ovary abortion is prevalent in diverse maize inbred lines and is under genetic control. Sci Rep 8:13032. https://doi.org/10.1038/ s41598-018-31216-9. 53. Bai F, Daliberti M, Bagadion A, Xu M, Li Y, Baier J, Tseung CW, Evans MM, Settles AM (2016) Parent-of-origin-effect rough endosperm mutants in maize. Genetics 204:221–231

Chapter 2 EMS Mutagenesis of Arabidopsis Seeds C. Stewart Gillmor and Wolfgang Lukowitz Abstract The ethylating agent ethyl methanesulfonate (EMS) is widely used for inducing random point mutations. In Arabidopsis, treatment with EMS causes GC-to-AT transitions with great efficiency: it has been estimated that a population of 50,000 well-mutagenized plants harbors one or more transitions in almost every GC pair of the genome. These properties, combined with ease of use, make EMS a mutagen of choice for genetic screens. Here, we describe a protocol for mutagenizing Arabidopsis seed with EMS. In addition, we briefly consider the germ line sectors typically induced by this treatment, and approaches for estimating the rate of induced mutations. Key words Mutagenesis efficiency, Embryo defective, Seed mutant, Genetic mosaic, Germ line sector

1

Introduction Induced mutations are at the heart of experimental biology, both in directed and in open-ended settings. Targeted approaches, such as genome editing, empower the functional analysis of specific genes or genetic pathways. In contrast, unguided, broad approaches, such as mutant screens, can serve as tools for uncovering novel functions or identifying previously unknown interactions. Screening procedures for embryo or seed traits are typically labor-intensive, posing a practical limit to the number of individuals that can be examined (see Chapter 1 of this issue for examples and discussion). Thus, it is important to ensure that a large number of mutant alleles are represented in a small mutagenized population. The resulting requirement for a highly effective mutagen is met remarkably well by ethyl methanesulfonate (EMS). Treatment with EMS is inexpensive, easy to implement, and can be used to induce point mutations at a high rate with good consistency in many different genetic backgrounds or species. EMS is an ethylating agent that can modify several positions in the bases of DNA [1]. The mutagenic effect is almost exclusively due to O6-ethylation of guanidine. O6-ethylguanidine pairs more

Martin Bayer (ed.), Plant Embryogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2122, https://doi.org/10.1007/978-1-0716-0342-0_2, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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stably with thymine as opposed to cytosine, such that an O6-GC pair is frequently converted to an AT pair during replication. Such transitions were essentially the only class of induced changes uncovered by the Arabidopsis TILLING facility among more than 1000 sequenced mutations from an EMS-mutagenized population [2]. Importantly, no evidence for positional bias was uncovered, suggesting that GC pairs are targeted at random, irrespective of their genomic or sequence context [3]. In the vast majority of cases, mutant phenotypes result from loss of gene function due to premature stop codons, disruption of splice sites, or deleterious amino acid substitutions. Point mutations conferring novel properties are obtained in rare cases (for example resistance to micro-RNAs [4]; resistance to auxin-dependent degradation [5]; or temperaturesensitivity [6]). Mutations resulting in ectopic gene expression are not well represented in EMS mutagenized populations and should be screened by alternative methods, such as activation tagging or transposon insertion. Two studies using different methodologies came to very similar estimates on the efficiency of EMS treatment: in well mutagenized populations of Arabidopsis, each diploid genome harbors about 700 point mutations [2, 7]. This is an impressively high rate—it implies, for example, that nearly all GC pairs of the Arabidopsis genome are mutated once or more in a sample of 50,000 mutagenized plants. Thus, a relatively small sample of about 1000 mutagenized plants, while certainly not exhaustive, should be sufficiently large for validating or dismissing a novel screening idea (see Chapter 1 of this issue). This protocol of Arabidopsis seed mutagenesis describes in detail how to administer EMS-treatment, how to assess the effectiveness of the mutagenesis, and which basic parameters to consider when analyzing the resulting population of mutants.

2

Materials

2.1 Laboratory and Safety Equipment

1. Chemical hood. 2. Lab coat, safety goggles, gloves. 3. Magnetic stir plate and stir bars.

2.2 Glass and Plasticware

1. 1 mL disposable syringe with needle. 2. 10 mL plastic pipette (for stirring). 3. 5 L plastic beaker or container. 4. 2
100–250 mL glass bottles with tightly closing screw cap. 5. 500 mL glass beaker. 6. 1 L bottle of water.

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2.3 Arabidopsis Seeds

1. 100 mg, corresponding to about 5000 seeds, provide sufficient material for most purposes. Although treatment with EMS can be expected to reduce germination rates and to cause sterility in a portion of treated plants, we have never obtained less than 2000 fertile plants from 5000 seeds mutagenized as described below. It is a good idea to grow the seed from a single or a few quarantined plant(s) to avoid potential contamination by pollen of a different genotype (see Note 1).

2.4 Reagents and Solutions

1. Ethyl methanesulfonate (EMS; MW 124; CAS# 62-5-0): If possible, a fresh batch should be purchased for the experiment. Only a fraction of a gram will be needed (see Note 2). 2. Inactivation solution: In a 5 L beaker, dissolve 100 g sodium thiosulfate pentahydrate (MW 248; CAS# 10102-17-7) in 4 L of water to make a 0.1 M solution. Stir briefly with 10 mL plastic pipette until dissolved. Leave the plastic pipette in the beaker for later use. The solution can be prepared ahead of time (see Note 3). 3. Dilute agar solution: In a 1 L bottle, mix 0.4–1 g of Bacto agar with 500 mL water and autoclave. The solution should be liquid but dense enough to prevent immersed seeds form sinking. The most effective concentration will depend on the batch of agar, but solutions between 0.04% and 0.1% should all be adequate. More or less of the solution may be needed dependent on how many seeds are mutagenized and how densely they are sown. 4. Soil mixture, pots, trays, etc. (for growing plants).

3

Methods

3.1 EMS Mutagenesis of Arabidopsis Seeds

Pretreatment (see Note 4) 1. Weigh out the desired amount of seeds (see Subheading 2 and Note 1). 2. Option A (5 days): Spread seeds out on a filter paper, taking care to avoid clumps or crowding. Soak the paper in water and incubate in a moist chamber at 4  C for 2–4 days to prime the seeds for germination. Then dry the seeds back by placing the paper in a growth chamber or warm room for a day. Carefully scrape the dry seeds off their support, weigh out the desired amount and proceed to step 4. Option B (overnight): Place the seeds in a 100 mL glass bottle with tightly closing screw cap and imbibe in 0.01% Tween 20 for 15 min. Pour out the Tween 20 solution (along with any seeds that are still floating) and rinse the seeds several times with water, until no more bubbles form.

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Add 40 mL water and leave the bottle at 4  C overnight on a rocker or small shaker providing gentle agitation. Pour out the water and proceed to step 4. Day 1: Mutagenesis (start early, the seed will incubate with EMS for 6–8 h) 3. Prepare working space in chemical hood: Clear an adequate area, turn on ventilation, and post a sign to caution coworkers of the ongoing mutagenesis. Place the container with decontamination solution, a magnetic stirrer, and a rocker or small shaker on the working space. Cover the stirrer and the rocker/ shaker with bench-liner paper. Fill 50 mL of water in a 100 mL glass bottle with tightly closing lid, add a small stir bar, and place the bottle on the stirrer. Have ready: EMS bottle and 1 mL disposable syringe with needle. 4. Protective equipment: When handling EMS, wear a lab coat, safety goggles, and double gloves. 5. Prepare 0.3% EMS solution: EMS is typically sold in bottles that have a screw cap as well as a plastic plug—remove the cap but not the plug. Pierce through the plug with the needle and draw 0.15 mL of the EMS into the syringe. Inject the EMS into the glass bottle on the stirrer, making sure that the tip of the needle is submerged in the water. Drops of the EMS will sink to the bottom of the bottle. Fill the syringe with decontamination solution and drop it into the decontamination container. Close the glass bottle and turn on the magnetic stirrer—mix for about 5 min until no more droplets are visible. Remove the magnetic stirrer from the hood (see Note 5). 6. Discard EMS bottle (optional): If the remaining EMS is not to be stored for further use, it can now be inactivated. Carefully remove the plastic plug of the bottle. Submerge screw cap, plug, and bottle in the decontamination container. Remove outer gloves, place in the decontamination solution and submerge them using the plastic pipette. Put on new pair of outer gloves. 7. Start mutagenesis: If option A was followed for pretreatment (step 1), add the pretreated dry seeds to the EMS solution. Close the bottle, place it on rocker/shaker and gently agitate to disperse the seeds. If option B was followed for pretreatment (step 1), pour the EMS solution into the glass bottle containing the pretreated seeds. Carefully submerge the empty bottle used to prepare the EMS solution including the stir bar and lid in the decontamination container. Close the bottle containing the seeds, place it on the rocker/shaker and gently agitate.

Arabidopsis EMS Mutagenesis

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8. Incubation: Incubate the seeds for 8 h. Monitor from time to time to make sure that seeds are covered by the EMS solution (see Note 5). 9. Rinsing out the EMS: Have a 250 mL glass beaker and a bottle with 1 L of water ready. Remove bottle with seeds and EMS solution from the rocker/shaker and let the seeds sink to the bottom. Remove rocker/shaker from the hood. Carefully open the bottle and decant the EMS solution (including any seeds that may be floating on the surface of the solution) into the decontamination container. Rinse the seeds three times with water: fill the bottle about half way, briefly swirl, let the seed sink to the bottom, and carefully decant the rinse water into the decontamination container. Fill the bottle with water one more time, swirl to suspend the seeds, and pour the seed/water slurry into a 250 mL glass beaker; repeat if necessary to recover all seeds. Place the empty glass bottle and the lid in the decontamination container. Cover the decontamination container with tin foil and leave in the hood overnight. 10. Washing the seeds: The seeds can now be handled outside of the chemical hood. Continue washing the mutagenized seeds thoroughly with tap water—go through 10 or more cycles of rinsing/decanting over the period of 1 h, such that remaining traces of EMS can diffuse out of the seeds. Rinse water can be discarded in the sink (see Note 6). 11. Sowing the seeds: Suspend seeds in dilute agar (see Subheading 2) as needed. We usually start with 500 mL of slurry, make aliquots of about 200–500 seeds each in 100 mL bottles, and then distribute the seeds of a bottle to one tray of pots by pipetting or pouring. Depending on the screening strategy for the plants, lower or higher densities may be required. Day 2: Clean-up 12. Overnight incubation in decontamination solution is more than sufficient to destroy the EMS. The syringe/needle should be discarded in a sharps container. Glass or plastic ware, and the magnetic stirrer can be washed and reused. The decontamination solution can be disposed in the sink. 3.2 Arabidopsis M1 Generation: Sectors, Mutation Rate, and Seed Pooling

1. Mosaic nature of mutagenized plants: The plants growing from the mutagenized seeds are commonly referred to as M1 generation. Since the EMS treatment affects each cell in a unique way, M1 plants are genetic mosaics. Indeed, their leaves occasionally show mutant sectors of white or light green color. Only about two sectors on average contribute to the generative cells of the main inflorescence (in other words: all gametes produced by the main inflorescence can be traced back to about two cells in the shoot apical meristem of the mature seed; see Fig. 1).

20

C. Stewart Gillmor and Wolfgang Lukowitz M1 seeds: EMS-treatement induces different mutations in each cell +/-

M1 plants: ~ two mutant sectors contribute to the germline

+/+

-/-

+/-

M2 seeds: siliques are clonal, producing families with segregating mutations

Fig. 1 Seed mutagenesis and the creation of germ line sectors. Blue and orange colors represent independent mutations segregating in the siliques of mosaic M1 plants. The seeds produced within a silique after self-pollination form an M2 family

Lateral inflorescences as well as individual siliques of the main inflorescence are thought to be clonal. 2. Assessing the efficiency of mutagenesis: While accurately measuring the rate of induced mutation may not be simple, a reasonable estimate can readily be obtained from the frequency of “embryo-defective” mutations. This group is defined loosely here, including any mutation with a clearly noticeable effect on the appearance of mature or nearly mature seeds—from dark colored, collapsed seeds that seem empty to shrunken or irregular seeds containing misshaped or rudimentary embryos. In a well-mutagenized M1 population, an average cell contains about one induced embryo-defective mutation. These mutations (as long as they follow Mendelian segregation) can easily be detected in self-fertilized siliques: about 25% of the contained seeds will be homozygous and thus abnormal. The average number of embryo-defective mutations can most easily be assessed indirectly, by estimating the frequency of M1 sectors that produce all normal seeds. To do so, mature siliques from about 100 different M1 plants are examined: if no embryo-defective seeds are contained in the silique, the corresponding M1 sector is said to produce all normal seeds. According to the Poisson distribution, the average frequency of embryo-defective mutations is 1 or greater as long as fewer than about 40% of the sectors produce all normal seeds [8, 9]. A similarly stringent cut-off was used by the Arabidopsis TILLING facility, and the selected mutagenized population was found to harbor about 700 induced point mutations per genome [2].

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3. Harvesting the M1 generation in pools: Screens for mutant embryo or seed phenotypes can be performed directly on M1 plants, by examining siliques produced by self-pollination. A benefit of this approach is that each plant harbors on average two different mutant sectors, which can help to increase screening efficiency. But since EMS treated seeds cannot be easily stored, mutagenesis and screening must be tightly coordinated (see Chapter 1 of this issue for an example). Alternatively, the mutagenized plants can be harvested and the screen performed with plants of the M2 generation. This approach allows for more flexibility, since M2 seeds can be stored for long periods of time. Another advantage is that any physiological stress caused by the mutagenesis treatment, a potentially confounding factor in the M1 generation, should be overcome in the M2 generation. For convenience, M2 seeds are typically bulk-harvested from pools of 100 to 1000 M1 plants. Allelic mutations recovered in different M2 plants from the same pool must be assumed to be siblings (unless revealed as different by molecular analysis). Therefore, M2 pools are typically undersampled to maximize the recovery of independent mutations (discussed in [8]; see [10] for an example).

4

Notes 1. The average weight of Arabidopsis seeds can differ substantially between genotypes (see [11] for an assessment of commonly used wild accessions) and strongly depends on the physiological condition of the fruiting plant. The value of ~0.2 mg/dry seed is a rough but reasonable estimate. 2. EMS is classified as Category 1B germ cell mutagen and Category 2 carcinogen: it is known to cause germ cell mutations and suspected to cause cancer. At room temperature, EMS is liquid, with a slightly yellow or brownish color and a density of 1.2 g/ L. EMS may decay over time, suggesting that the “potency” of old stocks in a mutagenesis experiment can vary. If a freshly purchased batch is not available, we recommend trying several different EMS concentrations (as described in Note 5). 3. Sodium thiosulfate is also known as “hypo,” from sodium hyposulfate. It is widely used as a fixative in black and white photographic processing, and readily available at minimal cost. Handling poses little or no risk. De Meo and colleagues tested several protocols for decontaminating alkylating agents, including EMS [12]. Among the solutions tried, 1 M sodium thiosulfate had the strongest effect, cutting the half-life of EMS to less than a minute. No mutagenic breakdown products were

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C. Stewart Gillmor and Wolfgang Lukowitz

created in the process. Treatment with 1 N sodium hydroxide, a procedure often described in mutagenesis protocols, was also found effective. 4. The pretreatment aims at breaking dormancy and establishing similar physiological conditions within the seeds of the batch. Anecdotal evidence suggests that seeds with different physiological states react differently to the EMS treatment, introducing a potentially large plant-to-plant variability. 5. Treatment with 0.3% EMS for 8 h, as suggested here, typically induces an adequate rate of mutations. However, the results of mutagenesis may vary—different genotypes or seed batches can show different sensitivity to EMS; the “potency” of EMS stocks will decrease in storage. Should the EMS treatment be too harsh, the mutagenized plants will be largely sterile. Should the treatment be too weak, relatively few mutations will be found (see Subheading 3.2 for how to estimate the rate of induced mutations). In these cases, we recommend setting up three (or more) parallel treatments with varying concentrations of EMS that cover an appropriate range between 0.1% and 0.5%. Since EMS decays during the mutagenesis, the effects of varying the incubation time seem less predictable. 6. Repeated rinsing over a prolonged period of time is important: residual EMS may cause sterility.

Acknowledgments This work was supported by grants from CONACYT Mexico (237480 to C.S.G.) and the NSF (IOB-04541201 to W.L.). We gratefully acknowledge I. Furner (Cambridge, UK), M. Hu¨lskamp (Cologne, Germany), J. Langdale (Oxford, UK), O. Leyser (Cambridge, UK), M. Muszynski (Honolulu, HI), and E. Vollbrecht (Ames, IA) for posting or communicating mutagenesis protocols we have consulted over the years. References 1. Sega GA (1984) A review of the genetic effects of ethyl methanesulfonate. Mutat Res 134:133–142 2. Till BJ, Reynolds SH, Greene E, Codomo CA, Enna LC, Johnson JE, Burtner C, Odden AR, Young K, Taylor NE, Henikoff JG, Comai L, Henikoff S (2003) Large-scale discovery of induced point mutations with highthroughput TILLING. Genome Res 13:543–530 3. Greene EA, Codomo CA, Taylor NE, Henikoff JG, Till BJ, Reynolds SH, Enns LC, Burtner C,

Johnson JE, Odden AR, Comai L, Henikoff S (2003) Spectrum of chemically induced mutations from a large-scale reverse-genetic screen in Arabidopsis. Genetics 164:731–740 4. Mallory AC, Reinhart BJ, Jones-Rhoades MW, Tang G, Zamore PD, Barton MK, Bartel DP (2004) MicroRNA control of PHABULOSA in leaf development: importance of pairing to the microRNA 50 region. EMBO J 23:3356–3364 5. Hamann T, Benkova E, B€aurle I, Kientz M, Ju¨rgens G (2002) The Arabidopsis

Arabidopsis EMS Mutagenesis BODENLOS gene encodes an auxin response protein inhibiting MONOPTEROS-mediated embryo patterning. Genes Dev 16:1610–1615 6. Arioli T, Peng L, Betzner AS, Burn J, Wittke W, Herth W, Camillieri C, Ho¨fte H, Plazinski J, Birch R, Cork A, Glover J, Redmont J, Williamson RE (1998) Molecular analysis of cellulose biosynthesis in Arabidopsis. Science 279:717–720 7. Jander G, Baerson SR, Hudak JA, Gonzales KA, Gruys KJ, Last RL (2003) Ethylmethanesulfonate saturation mutagenesis in Arabidopsis to determine frequency of herbicide resistance. Plant Physiol 131:139–146 8. Redei GP, Koncz C (1992) Classical mutagenesis. In: Koncz C, Chua NH, Schell J (eds) Methods in Arabidopsis research. World Scientific Publishing, Singapore, pp 16–82 9. Weigl D, Glatzebrook J (2002) EMS mutagenesis of Arabidopsis seeds. Cold Spring Harb

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Protoc. https://doi.org/10.1101/pdb. prot4621 10. Gillmor CS, Roeder AHK, Sieber P, Somerville C, Lukowitz W (2016) A genetic screen for mutations affecting cell division in the Arabidopsis thaliana embryo identifies seven loci required for cytokinesis. PLoS One 11:e0146492. https://doi.org/10.1371/jour nal.pone.0146492. 11. El-Lithy M, Clerkx EJM, Ruys GJ, Koorneef M, Vreugdenhil D (2004) Quantitative trait locus analysis of growth-related traits in a new Arabidopsis recombinant inbred population. Plant Physiol 135:444–458 12. De Meo M, Laget M, Castegnaro M, Dumenil G (2010) Evaluation of methods for destruction of some alkylating agents. Am Ind Hyg Assoc J 51:505–509

Chapter 3 EMS Mutagenesis of Maize Pollen A. Mark Settles Abstract Effective mutagenesis is critical for connecting traits of interest to specific plant genes. The development of site-directed mutagenesis and sequenced-indexed genetics resources in maize allows for targeted analysis of individual genes. These reverse genetics approaches have the potential for confirmation bias by only studying candidate genes for association with traits of interest. Genetic screens of induced, random mutations are important for identifying novel loci as well as interacting factors for known mutant loci. Chemical mutagenesis provides very high mutation rates and can be used for a variety of screen designs. This chapter provides an updated protocol for ethyl methanesulfonate (EMS) mutagenesis of maize pollen using paraffin or mineral oil. Mutagenesis occurs in mature pollen causing nonconcordant endosperm and embryo genotypes as well as sectored M1 plants. Considerations for these factors in genetic screens are discussed. Key words Mutagenesis efficiency, Sperm cell, Germinal mutation, Nonconcordant kernel, Defective kernel (dek), Embryo defective (emb), Endosperm

1

Introduction Maize is versatile model organism that has major importance for our food supply. Worldwide production of maize is over 1 billion metric tons, making it the primary cereal crop in agriculture (www. fao.org). Maize has been a focus of genetic studies since Gregor Mendel and has been used to study a large diversity of genetic phenomena [1]. Transposon-induced mutagenesis was first observed in maize [2]. The diversity of endogenous transposons enabled many of the maize genetics resources to be based on transposon-induced mutagenesis [3, 4]. Efficient chemical mutagenesis was developed that enabled much higher rates of mutation than that observed with transposons [5]. This protocol suspends ethyl methanesulfonate (EMS) in paraffin or mineral oil to allow EMS to diffuse into pollen and sperm cells without germinating the pollen grain (Fig. 1a). Treated pollen is then applied to ear silks to fertilize nonmutagenized ovules. Mature pollen has two sperm cells and EMS treatment will

Martin Bayer (ed.), Plant Embryogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2122, https://doi.org/10.1007/978-1-0716-0342-0_3, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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segregating ear emb

germinal

sagittal normal

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mutant

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emb + end EMS-treated pollen: sperm cells affected independently

M1 kernels: embryo and endosperm M1 plants: form distinct heterozygous sectors

emb + end end

Fig. 1 Outline of a pollen mutagenesis and examples of kernel phenotypes. (a) Two sperm cells of pollen are drawn in different colors (right); after double fertilization, mutations induced in one sperm are contributed to the endosperm, while mutations induced in the other sperm are contributed to the embryo of a kernel (middle); each kernel is an independent event and will typically give rise to a heterozygous M1 plant (left). (b) Mutant kernels segregating on a heterozygous ear: defects can be specific to the embryo (row labeled emb), to the endosperm (row labeled end), or can appear in both parts of the kernel (rows labeled emb + end); germinal surface views and in sagittal hand sections of mutant and normal kernels are shown; arrowheads point to defective kernels of the ear (white), the root–shoot axis of the embryo (red), the embryonic root (orange), and the embryonic shoot (green). Scale bar: 5 mm

mutagenize these cells independently. Consequently, M1 plants and M2 progeny are screened for germinal mutations that were transmitted into the M1 embryo. Initial EMS mutagenesis focused on defective seed phenotypes [6]. These studies showed distinct classes of loci that affect the development of the endosperm, embryo, or both products of double fertilization (Fig. 1b). EMS mutagenesis is still a common approach for a multitude of genetic screen designs (discussed in [3] and Chap. 1 of this issue). In nondirected mutagenesis, pollen is treated and crossed onto normal plants. The resulting kernels will have different endosperm and embryo genotypes due to EMS impacting the sperm cells independently. The M1 plants are then self-pollinated to generate M2 families. These families can be screened for specific phenotypes of interest, such as embryo defective [7] or inflorescence development phenotypes [8]. Recessive mutants will be expected to segregate within an M2 family. EMS-treatment of maize pollen can also be used to generate additional alleles by crossing mutagenized pollen to a reference mutant [9]. In these screens, normal pollen is mutagenized and crossed to mutant plants. It is imperative that the mutagenized,

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normal parent have distinct flanking marker polymorphisms relative to the reference mutant. This allows for new alleles to be genetically separated after an allele is identified. Finally, EMS is used for enhancer and suppressor screens by mutagenizing pollen from the reference mutant and crossing to the reference mutant [10]. M1 plants are then self-pollinated to identify more severe or suppressed phenotypes that segregate as unlinked dominant or recessive loci. The main advantage of EMS for these applications is that a much smaller number of mutagenized progeny need to be screened to identify mutants of interest. This chapter describes maize pollen mutagenesis that can be conducted in field grown plants. The protocol details best strategies for managing a mutagenesis experiment to minimize handling of this potent carcinogen as well as factors to consider due to the potentially sectored M1 plants that will be generated by treating mature pollen.

2

Materials EMS is a potent carcinogen. This protocol produces EMS-oil liquid and solid waste. Consult your chemical safety office for local waste containment and disposal procedures prior to initiating mutagenesis.

2.1 Laboratory and Safety Equipment

1. Chemical hood. 2. In the laboratory: Lab coat, safety goggles, and gloves. 3. In the field: Tyvek-type disposable clothing, respirator with combination filter cartridge, safety goggles (if respirator is half-face), and gloves. 4. Magnetic stir plate and stir bars.

2.2 Supplies and Plasticware

1. Tea strainer. 2. Disposable plastic centrifuge tubes with volume graduations. 3. 100–250 mL glass bottles with tightly sealing caps. 4. 100–250 mL plastic applicator squeeze bottles.

2.3

Chemicals

1. Ethyl methanesulfonate. 2. Paraffin or mineral oil (referred to as “oil”).

2.4

Plant Material

1. Maize plants: Pollen treatments will reduce kernel set to about 10% of normal hand pollinations. Depending upon the genetic stock, 100–200 plants should be crossed with mutagenized pollen. This will yield 3000–10,000 M1 kernels. Pollen parents with robust shed are preferred, but poor shedding genotypes can be used as long as sufficient plants are available.

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Field Supplies

1. Maize pollination (tassel) bags, ear shoot bags. 2. Permanent marker, paper clips, plier stapler, staples.

3

Methods

3.1 EMS Mutagenesis of Maize Pollen

Field preparations 1. If mutagenesis will involve crosses of mutagenized pollen parent onto a female parent with a different genetic background, it is important to characterize the flowering time of each line in the local field or greenhouse environment. Plant the two parents for pollen shed to be coincident with ear silking. Planting the genotypes on multiple dates ensures crosses can be made with robust pollen shed and fresh silks. 2. Follow typical controlled pollination practices for maize. When tassels begin to be visible, scout and cap all first ear shoots prior to silking. In order to avoid outcross contamination, ear shoots with exposed silks should not be used. 3. It is ideal to have sufficient plants that all tassels can be removed from plants that will be used as female parents in the mutagenesis. If this strategy is to be used, plant male pollen parents in a separate block from female parents. Day 0: prepare for mutagenesis crosses 4. Monitor both male and female planting blocks to judge when 35–50% of plants are at an optimum for crosses. There should be at least 35 ears with sufficient silks for cut-back and pollination on Day 1. Cut back each of these ears and mark the shoot bag to indicate the plant is ready to pollinate on Day 1. If you have separate blocks, all tassels should be removed from the female block of plants. 5. At least 30 tassels should be ready to give optimum pollen shed for Day 1. Typically, select tassels with exposed anthers throughout the central spike with a few anthers exposed on lateral tassel branches. Bag these tassels and secure them with a paper clip. Day 0: prepare ems stock solution 6. Clear an adequate area in your laboratory chemical fume hood, turn on ventilation, and post a sign to caution coworkers of the ongoing mutagenesis. Place a solid hazardous chemical waste container labeled for contamination with EMS in the work area. Place a magnetic stirrer on the work space. Cover the stirrer with bench-liner paper. Fill 50 mL of oil in a 100 mL glass bottle with a tightly closing lid. Add a small stir bar, and

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place the bottle on the stirrer. Have ready: EMS bottle and 1 mL disposable syringe. 7. Protective equipment: When handling EMS, wear a lab coat, safety goggles, and double gloves. 8. EMS is typically sold in bottles that have a screw cap as well as a plastic plug—remove both cap and plug. Draw 0.5 mL of the EMS into the syringe. Add the EMS into the glass bottle on the stirrer, making sure that the syringe is submerged in the oil. Dispose of the syringe in the solid hazardous waste container. Close the glass bottle and turn on the magnetic stirrer and mix for at least 1 h (see Note 1). Day 1: mutagenesis (start early to ensure pollen collection as soon as shed begins) 9. If a chemical hood is available near your field site, prepare the chemical working space and use protective clothing as described for the EMS stock solution. Stir the EMS stock solution for at least 1 h. Alternatively, prepare the treatment solution at your laboratory and transport to the field site. 10. Prepare 0.067% EMS treatment solution: The treatment solution is a 1:15 dilution of the EMS stock solution in oil. Prepare an appropriate amount of treatment solution for the number of ears cut back on Day 0. Budget 1 mL of treatment solution per pollination. Assuming 50 pollinations, add 56 mL of oil in a 100 mL glass bottle with a tightly closing lid. Add a small stir bar, and place the bottle on the stirrer. Add 4 mL of the EMS stock solution. Stir vigorously for 1 h (see Note 2). 11. Monitor pollen shed for your genotype of interest. It is important to get dry, viable pollen. If there was rain or heavy dew overnight, it is a good idea to change the tassel bags early in the morning prior to shed. Once pollen is shedding for your pollen parent genotype, collect and pool the pollen. Sift the pollen through the tea strainer into a plastic tube with volume gradations. A paper or plastic funnel can be used to help guide pollen into the tube. A 1/10 volume of pollen relative to the mutagenesis treatment solution is needed. The pollen should be free-flowing without anthers or insect contaminants (see Note 3). 12. In the chemical fume hood, add 6 mL of pollen to the 60 mL EMS treatment solution. Use the magnetic stirrer to stir the pollen gently in the solution for 45–60 min. Transfer the treated pollen into a plastic applicator squeeze bottle and return to the field. 13. Protective equipment: All personnel handling EMS in the field need to wear Tyvek-type disposable clothing, a respirator with a combination filter cartridge, and gloves. A full-face respirator

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is recommended but safety goggles can be used in combination with a half-face respirator. Be sure to double-contain all EMS solutions whenever transferring between locations. 14. Mix the pollen and oil solution by inversion prior to each pollination. Remove the cap for the squeeze applicator bottle and apply 0.5–1 mL of pollen/oil mix to the silks of each cut-back ear. Cover the silks immediately after application of pollen by covering the ear with a tassel bag and stapling the bag around the plant. Pollinations should be completed within 1 h after the mutagenesis treatment. 15. Contain the applicator squeeze bottle with any remaining EMS-oil for transport back to your laboratory and hazardous chemical waste disposal. Change gloves after you have finished handling EMS solutions. 16. Prior to removing safety equipment, survey the nonpollinated plants in your mutagenesis block. Typically, you would skip 1 day between mutagenesis treatments to minimize the number of days required for complete coverage of the block. If the remaining 50% of plants are ready for pollination, cut back ears and bag tassels as in Day 0. Days 2–3: complete mutagenesis 17. Continue to monitor the planting for mutagenesis until the remaining plants can be pollinated or you have reached your target number of pollinations. Each additional cycle of pollinations follows the same procedure. 18. Safety precautions: Tassel bags covering EMS-mutagenized pollinations should be clearly labeled. The mutagenized block should be cordoned off to discourage entry into the field and a pesticide application sign should be used to indicate that part of the field was treated with EMS. Expected half-life of EMS in the environment is approximately 4 days (http://toxnet.nlm. nih.gov), and the EMS mutagenized block should not be entered without safety equipment for at least 12 days after final pollinations. Days 30–45: harvest 19. Mature ears can be harvested with standard procedures and require no special safety precautions. Mutagenized kernels should be dried to 10–15% moisture prior to planting. 3.2 Maize M1 Generation: Nonconcordant Kernels, Production of M2

1. Mosaic nature of mutagenized kernels and plants: Pollen mutagenesis independently affects the two sperm cells in the pollen grain. One sperm cell fuses with the central cell to create the endosperm, while the other sperm cell fuses with the egg to form the embryo. Kernel mutations observed in the M1 will

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have nonconcordant endosperm and embryo genotypes. Thus, dominant kernel endosperm phenotypes are unlikely to be inherited, since the embryo will not carry the causative mutation. M1 plants will typically be heterozygous for induced mutations. However, it is also possible that only one strand of the sperm cell DNA is mutant, giving rise to half-plant sectors in the M1 generation. Self-pollinating plants sectored for a recessive mutant will give less than 3:1 mutants in the M2 generation and may only show segregation in part of the ear. 2. Assessing the mutagenesis: Mutagenesis efficiency can be estimated based on the frequency of recessive defective kernel (dek) and embryo-specific (emb) mutations observed in the ears of self-fertilized M1 plants or M2 ears. These mutations segregate with a frequency of approximately 12–25%, depending upon whether the M1 plant is sectored. The defective kernel class includes any mutation with a clearly noticeable effect on the appearance of mature seeds. Embryo-specific mutants cause the embryo to abort and can also be identified in mature kernels. But since most of the kernel consists of endosperm, embryo-specific mutations are associated with relatively subtle visual defects [7, 11]. In genotypes expressing endosperm anthocyanins, embryo-specific mutants cause more intense color expression, allowing them to be distinguished as darker kernels [12]. In backgrounds without endosperm anthocyanins, embryo-specific mutants can be more clearly observed in a side or germinal view. In a well-mutagenized M1 population, approximately 25–30% of the M2 ears will segregate for defective kernels and about 10% for embryospecific phenotypes [6, 7]. 3. Screening and testing for heritability: Screens for embryospecific and defective kernel phenotypes can be performed on mature M2 ears. M2 ears stored at 10  C and 50% relative humidity will retain viability for 10–15 years, allowing the mutagenized population to be stored and screened as time permits. Mature mutant kernels can be screened for phenotypes of interest to select isolates for heritability tests. Defective kernel and all embryo-specific mutants are typically lethal, such that inheritance is tested in 15–30 normal siblings from the same M2 ear by self-pollination reciprocal crosses with a normal genetic stock. Recessive mutations will show a 3:1 normal– mutant segregation pattern in the self-pollinations and no mutant kernels in crosses. Dominant mutations will become apparent by segregating for mutant kernels in all crosses. Dominant mutations will often have incomplete penetrance and segregate at ratios below the expected 3:1 mutant–normal ratio. Parent-of-origin effect mutants will segregate for mutant kernels in self-pollinations and only one direction of reciprocal

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crosses [13]. In Arabidopsis, parent-of-origin mutations resulting in aborted seeds can be recognized by empty spaces accounting for approximately 50% of the positions normally filled with seeds in the siliques [14]. In maize, the position of aborted ovaries tends to be overgrown by neighboring seeds, making similar estimates more difficult [15].

4

Notes 1. The 1% EMS stock solution in oil is stable for years and can be stored at room temperature [16]. If long-term storage is planned, double containment of the stock solution is highly recommended. It is also possible to make 0.067% EMS-oil solution directly by pipetting 67 μL of EMS into 100 mL of oil with vigorous stirring for 1 h prior to adding pollen. 2. The mutagenesis treatment is temperature sensitive and best completed in a temperature-controlled laboratory. However, mutagenesis in the field is possible. If a field treatment is necessary, hot temperature conditions should be avoided. The experimenter will hand mix the pollen and EMS treatment solution every 3 min during the 45–60 min mutagenesis. 3. A separate block of pollen parents is highly recommended even if the mutagenesis cross is between siblings of inbred lines. EMS-treated pollen has low viability. Leaving tassels that are shedding pollen close to the female plants will cause outcross contamination with nonmutagenized pollen, effectively reducing the mutation load in the M1 generation.

Acknowledgments This work was supported by grants from NIFA (SCRI-201851181-28419), the NSF (IOS-1444456), and the Vasil-Monsanto Endowment. References 1. Coe EH Jr (2001) The origins of maize genetics. Nat Rev Genet 2:898–905 2. McClintock B (1948) Mutable loci in maize. In: Carnegie Inst. wash. Year book, vol 47. Cold Spring Harbor, New York, pp 155–169 3. Candela H, Hake S (2008) The art and design of genetic screens: maize. Nat Rev Genet 9:192–203 4. Gault CM, Settles AM (2014) Functional genomics. In: Wusirika R, Bohn M, Lai J

(eds) Genetics, genomics, and breeding of maize. CRC Press, Boca Raton, FL, USA, pp 131–154 5. Neuffer MG, Coe EH Jr (1977) Paraffin oil technique for treating mature corn pollen with chemical mutagens. Maydica 23:21–28 6. Neuffer MG, Sheridan WF (1980) Defective kernel mutants of maize. I. Genetic and lethality studies. Genetics 95:929–944

Maize Mutagenesis 7. Brunelle DC, Clark JK, Sheridan WF (2017) Genetic screening for EMS-induced maize embryo-specific mutants altered in embryo morphogenesis. G3 (Bethesda) 7:3559–3570 8. Bai F, Reinheimer R, Durantini D, Kellogg EA, Schmidt RJ (2012) TCP transcription factor, BRANCH ANGLE DEFECTIVE 1 (BAD1), is required for normal tassel branch angle formation in maize. Proc Natl Acad Sci U S A 109:12225–12230 9. Fouquet R, Martin F, Fajardo DS, Gault CM, Go´mez E, Tseung CW, Policht T, Hueros G, Settles AM (2011) Maize rough endosperm3 encodes an RNA splicing factor required for endosperm cell differentiation and has a nonautonomous effect on embryo development. Plant Cell 23:4280–4297 10. Gallavotti A, Long JA, Stanfield S, Yang X, Jackson D, Vollbrecht E, Schmidt RJ (2010) The control of axillary meristem fate in the maize ramosa pathway. Development 137:2849–2856 11. Clark JK, Sheridan WF (1991) Isolation and characterization of 51 embryo-specific mutations of maize. Plant Cell 3:935–951

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12. McCarty DR, Settles AM, Suzuki M, Tan BC, Latshaw S, Porch T, Robin K, Baier J, Avigne W, Lai J, Messing J, Koch KE, Hannah LC (2005) Steady-state transposon mutagenesis in inbred maize. Plant J 44:52–61 13. Bai F, Daliberti M, Bagadion A, Xu M, Li Y, Baier J, Tseung C, Evans M, Settles A (2016) Parent-of-origin-effect rough endosperm mutants in maize. Genetics 204:221–231 14. Grossniklaus U, Vielle-Calzada JP, Hoeppner MA, Gagliano WB (1998) Maternal control of embryogenesis by MEDEA, a polycomb group gene in Arabidopsis. Science 280:446–450 15. Gustin JL, Boehlein SK, Shaw JR, Junior W, Settles AM, Webster A, Tracy WF, Hannah LC (2018) Ovary abortion is prevalent in diverse maize inbred lines and is under genetic control. Sci Rep 8:13032 16. Neuffer MG (1994) Mutagenesis. In: Freeling M, Walbot V (eds) The maize handbook. Springer, New York, p 212

Part II Microscopic Imaging and Phenotypic Analysis of Arabidopsis Embryos

Chapter 4 Live-Cell Imaging of Zygotic Intracellular Structures and Early Embryo Pattern Formation in Arabidopsis thaliana Minako Ueda, Yusuke Kimata, and Daisuke Kurihara Abstract Plant embryogenesis begins with fertilization and ends with the generation of the basic body plan of the future plant. Despite its importance, the dynamics of flowering plant ontogeny have long been a mystery, because the embryo develops deep in the maternal tissue. Recently, an embryonic live-cell imaging system was established in Arabidopsis thaliana by developing an in vitro ovule cultivation method and utilizing two-photon excitation microscopy (2PEM), which is suitable for deep imaging. This system enabled us to visualize intracellular dynamics during zygote polarization and monitor the cell division pattern during embryogenesis from the zygote until organ formation. In this chapter, we describe a method that allows for high-resolution imaging of cytoskeletal rearrangements in the zygote and long-term tracing of embryo patterning. Key words Zygote, Embryo, Live-cell imaging, Two-photon excitation microscopy, Arabidopsis thaliana

1

Introduction In most plants, the zygote divides asymmetrically along the apical– basal axis to generate apical and basal daughter cells, which develop into the shoot and the root, respectively [1–4]. Therefore, it is important to understand how the zygote polarizes before this asymmetric cell division and how the daughter cells produce a proper plant body during embryogenesis. Detailed observations of fixed embryos have revealed that the Arabidopsis thaliana zygote is highly polarized, with a nucleus at the apical tip and large vacuoles in the basal region [5], and the embryo exhibits a stereotypical cell division pattern [6–8]. Therefore, Arabidopsis has been used as a model system to analyze the molecular mechanisms underlying embryo formation. Nevertheless, effective tools for real-time observation of zygote polarization and embryo pattern formation have been lacking, because embryogenesis occurs in the ovule, which is deeply embedded in the flower. Therefore, an

Martin Bayer (ed.), Plant Embryogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2122, https://doi.org/10.1007/978-1-0716-0342-0_4, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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in vitro ovule cultivation system was recently developed to grow the embryo from the zygote stage to a fertile plant under a microscope [9]. This system was combined with 2PEM, which is compatible with deep plant tissue imaging [10] to perform high-resolution imaging of various intracellular dynamics, such as nuclear migration and cytoskeleton organization, in the polarizing zygote [11, 12]. Furthermore, the development of a PDMS (poly [dimethylsiloxane]) microcage array and micropillar array device to fix the growing ovule in the liquid medium, and an automatic water-supply system to use a water immersion objective lens with a long working distance enabled long-term live imaging for more than 4 days [9, 13]. This system allows for the monitoring of cell division timing and cell fate specification from the zygote-stage to torpedo-stage embryo [9]. This live-cell imaging system was further combined with inhibitor assays to examine the roles of various specific factors, such as microtubules (MTs), on zygote elongation [12] and the effects of novel chemical compounds on embryonic cell proliferation [14]. In addition, optical manipulations, such as laser disruption, were employed to reveal the function of intercellular communication in embryo pattern formation [9].

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Materials 1. In vitro ovule cultivation medium (N5T medium): 1
Nitsch basal salt mixture, 5% (w/v) trehalose dihydrate, 0.05% (w/v) MES-KOH (pH 5.8), and 1
Gamborg’s vitamin solution. Sterilize the medium by filtering it through a 0.22-μm filter or autoclaving it (121  C, 20 min). The medium can be stored at 4  C for 2–3 months. 2. Fresh reproductive tissues of Arabidopsis plants harboring fluorescent markers to label specific structures of the zygote and/or embryo (e.g., actin filaments (F-actin) and nucleus) (Fig. 1, see Note 1). 3. PDMS micropillar array device (Fig. 2a, see Note 2). 4. Dissection microscope. 5. Fine-tip tweezers, square-tip tweezers, and dissecting needles (gauge 0.40 mm). 6. 70% ethanol. 7. Vacuum chamber. 8. Clean bench with ultraviolet (UV) light. 9. Glass slide (76
26 mm) and cover glass (18
18 mm). 10. 35 mm culture dish and 35 mm glass-bottom dish (singlewell). 11. Double-sided adhesive tape.

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Fig. 1 Live-imaging of zygote polarization and embryo pattern formation. (a) Nucleus and actin filament (F-actin) of the zygote were labeled with EC1p::H2B-tdTomato (magenta) and EC1p::Lifeact-Venus (green), respectively. (b) Nucleus of the embryo was labeled with WOX2p::H2B-GFP (green). The maximum-intensity projections (MIP) were generated from z-stack images. Numbers indicate the time (h:min) from the first frame. Arrowheads and brackets show the nucleus and the dividing nucleus, respectively. The corresponding stages of the zygote are shown. Bars ¼ 10 μm (a) and 30 μm (b)

Fig. 2 PDMS micropillar array. (a, b) Micrographs of the PDMS micropillar array. (b) Ovules were successfully attached. Numbers indicate the pillar pitch (μm). Bars ¼ 1 mm

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Fig. 3 Automatic water-supply system. (a) Schematic diagram of 2PEM for long-term live imaging of the embryo. (b) Syringe pump to a needle with a holder. (c) The automatic water-supply system retained a droplet on the objective lens

12. Paraffin film. 13. Two-photon excitation microscope with an inverted microscope stand (Fig. 3a, see Note 3). 14. Water-immersion objective lenses (see Note 4). 15. Water-supply system for water-immersion objective lenses (Fig. 3b, see Note 5).

3

Methods

3.1 Preparation of PDMS Micropillar Array Device

1. Cut open the PDMS device with scissors to set on the cover glass (14 mm diameter) of a glass-bottom dish (approximately 9
9 mm). 2. Sterilize the PDMS device with UV light irradiation for 15 min on a clean bench. 3. Invert the PDMS device using tweezers and sterilize the other side for an additional 15 min.

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Fig. 4 Setup of the PDMS micropillar array in a glass-bottom dish. (a) The cut and sterilized PDMS device was soaked in a culture dish. (b) After degassing, the extra medium on the side and bottom of the PDMS device was removed. (c) The medium was placed on the micropillar array. (d) After extracting the ovules from the dissected silique (see Fig. 5), the ovules were completely soaked by pushing a cover glass. (e) The extra medium was removed by pulling a cover glass horizontally. (f) The PDMS device placed sideways and (g) inverted into a glass-bottom dish. (h) After slightly pushing the PDMS device to attach a cover glass, the N5T medium was gently poured by decanting the medium to completely soak until it exceeds the upper surface of the PDMS device. (i) The glass-bottom dish was sealed with paraffin film

4. Place the sterilized PDMS device in a culture dish and push it to attach tightly. 5. Pour approximately 5–7 mL of N5T medium to soak until it exceeds the upper surface of the PDMS device (Fig. 4a). 6. Transfer the dish into a vacuum chamber and degas the PDMS device for 3 h up to overnight (see Note 6). 7. Place the PDMS device on a paper towel to clean off the extra medium around the device (Fig. 4b). Be careful to retain the medium on the micropillar array (Fig. 4c, see Note 7).

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8. Transfer the device onto a glass slide, with the micropillar part facing the upper side (Fig. 4c). Cover the PDMS device with the lid of a culture dish to maintain humidity during the subsequent silique dissection. 3.2 Silique Dissection to Extract Ovules

1. Sterilize a needle and fine-tip tweezers by wiping with 70% ethanol. 2. Collect siliques at appropriate developmental stages for the experiments by using fine-tip tweezers (see Note 8). 3. Place the silique onto double-sided adhesive tape on a glass slide (Fig. 5a). 4. Under a dissection microscope, incise both sides of the replum using a needle (Fig. 5, see Note 9).

Fig. 5 Silique dissection to extract ovules. (a) The silique was placed onto a double-sided adhesive tape on a glass slide. (b) The silique was incised with a needle along the valve margin, between the replum and valve. (c) The valve was opened. (d) The other side of valve margin was incised and that valve was also opened

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5. Dip the opened silique into the N5T medium at the micropillar part of the device (Fig. 4c) and gently scrape the ovules from the silique into the medium using tweezers or a needle. 6. Place a cover glass on the PDMS device and push it to soak the ovules between the micropillars (Figs. 3d, 2b). 7. Take off the cover glass by pulling it horizontally to remove extra medium (Fig. 3e, see Note 10). 8. Invert and place the PDMS device into a glass-bottom dish (Fig. 3f, g). 9. Push the device using square-tip tweezers to prevent the removal of the device from the cover glass at the next step. 10. Pour the N5T medium gently into the glass-bottom dish to soak until it exceeds the upper surface of the PDMS device (Fig. 3h). Seal the dish with a paraffin film (Fig. 3i, see Note 11). 3.3 Live-Cell Imaging

1. Place the glass-bottom dish on the stage of an inverted microscope and observe the ovules under bright field illumination by adjusting the position and focus (see Note 12). 2. Observe the fluorescent signals of the zygotes/embryos and select suitable ovules to focus on. 3. Set the parameters for image acquisition (e.g., zoom and laser power) according to the manufacturer’s instructions of the microscope system (see Note 13). 4. Set the center plane of the z-position, the number and interval of z-slices for z-section image acquisition, and the duration and interval for time-lapse images (see Note 14). 5. Once the target events, such as zygotic division and embryo patterning, are completed, stop time-lapse imaging and review the acquired image sequence (see Note 15). 6. Analyze the acquired images and convert them to a general movie file, with extensions .avi or .mov (see Note 16).

4

Notes 1. Because the zygote and embryo are observed through the ovule, it is important to use specific fluorescent markers that are not expressed in other ovule tissues, such as endosperm and integument. Otherwise, the background fluorescent signals in the ovule will mask the zygotic/embryonic signals. 2. The PDMS device is necessary to fix the growing ovule in the liquid medium during long-term imaging. If a PDMS device is not used, the ovule will frequently move away from the observation field (Fig. 6a). The complete procedure of PDMS device

Fig. 6 Examples of failed imaging. (a) Time-lapse observation of an embryo: The growing ovule detached from the glass bottom and thus escaped from the observation field (13:50). (b–d) EC1p::H2B-tdTomato, magenta: zygotes expressing markers for the nucleus. (b and d) EC1p::Clover-TUA6, green: zygotes expressing markers for microtubules. (c) EC1p::Lifeact-Venus, green: zygotes expressing markers for F-actin. (b) The damaged ovule shrunk, and thus the zygote was crushed. The width of the embryo sac (yellow brackets) suddenly and rapidly reduced after 13:00. (c) The drifting ovule rapidly changed from the z-position at 0:00 and finally caused the image to be out of focus. (d) All fluorescence was gradually bleached due to intense laser irradiation. The MIP (b and d) and center plane (c) images are shown. Numbers indicate the time (h:min) from the first frame. Bars ¼ 50 μm (a) and 10 μm (b–d)

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construction has been described previously [9, 13]. Short-term imaging is possible without the PDMS device. 3. A confocal microscope is sufficient to observe cellular organization in the embryo, but visualization of intracellular structures in the zygote requires a two-photon excitation microscope with a femtosecond pulse laser and a highsensitivity detector (e.g., Nikon A1R MP system equipped with a Ti:sapphire femtosecond pulse laser and external non-descanned GaAsP PMT detectors). 4. We recommend using 40
and 25
objective lenses for zygote and globular embryo observation, respectively. Waterimmersion lenses with high numerical aperture (NA) and long working distance (WD) (e.g., Nikon CFI Apo LWD WI (NA ¼ 1.15, WD ¼ 0.59–0.61 mm) as the 40
lens and Nikon CFI Apo LWD 25
WI (NA ¼ 1.10, WD ¼ 2.00 mm) as the 25
lens) are suitable for deep imaging. 5. Because the water on objective lenses is easily dried, immersion oils suitable for water-immersion lenses (e.g., Zeiss Immersol W 2010) can be used for the 40
objective. However, because of their viscosity, such oils are not compatible with long working distance objective lenses. Therefore, an automatic watersupply system is necessary, in which a syringe pump applies water on the objective lens through a needle. The needle is attached to the lens by a handmade holder. The water flow rate to apply depends on room conditions, such as humidity and temperature. Our flow rates are 0.50–1.00 μL/min using a Fusion 100 syringe pump (Chemyx, Stafford, TX, USA). 6. The device can be detached from the bottom of the culture dish by air bubbles under a strong vacuum, but this does not affect the device. 7. The medium on the micropillar array is used for ovule cultivation. Therefore, if the medium is not sufficient, add more using a pipette. 8. The developmental stages of the ovules in a silique are roughly synchronized and, under our growth conditions, the zygotes and young globular embryos are mainly contained in approximately 5 mm and 8–10 mm siliques, respectively. 9. Cut only the silique surface (ovary wall) without damaging the ovules on the inside. It is important to dissect the silique gently but quickly, because injured or dried ovules easily die during live imaging (Fig. 6b). 10. If extra medium remains in the micropillar part, the PDMS device is easily detached from the glass bottom when the medium is poured (Subheading 3.2, step 10).

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11. The device may float if it is not completely degassed (Subheading 3.1, step 5) or if the medium is poured too quickly. Once the device floats, the ovules scatter into the medium and thus, the whole preparation needs be repeated from the beginning. 12. Usually, after switching on the mechanical system of the microscope, several hours are required for the temperature to stabilize due to thermal changes in the z-position of the stage and thus, the glass-bottom dish. If live imaging is immediately started after placing the glass-bottom dish on the stage, the z-position will shift during imaging (Fig. 6c). To avoid this “drift,” leave the dish on the stage for several hours before imaging or use a weight to accelerate z-position stabilization. 13. It is important to use the minimum laser power and irradiation to reduce damage to the ovules. If laser irradiation is too strong, the ovules are damaged (Fig. 6b) or the fluorescence is severely bleached (Fig. 6d). The microscopic parameters are diverse and depend on the equipment used. Therefore, appropriate settings should be chosen to fit the purpose of the experiment. 14. Our settings for zygote imaging are as follows: 31 z-slices with 1-μm intervals for 24 h with 20 min intervals; for embryo imaging: 9–11 z-slices are obtained with 5-μm intervals for 72–120 h at 10–15 min intervals. 15. The time-lapse imaging should be examined from time to time. Adjust the xy- and z-positions if the ovule position has changed. 16. Image analysis and conversion can be performed using ImageJ or Fiji, which are the open source programs inspired by NIH Image [15, 16].

Acknowledgments M.U., Y.K., and D.K. are supported by the Japan Society for the Promotion of Science (Grant-in-Aid for Scientific Research on Innovative Areas [JP17H05838 for M.U.], Grant-in-Aid for Challenging Exploratory Research [JP16K14753 for M.U.], Grant-inAid for JSPS Research Fellow [JP18J10512 for Y.K.], Grant-in-Aid for Scientific Research [B; JP17H03697 for D.K.], and Challenging Research [Exploratory; JP 18 K19331 for D.K.]). This work was supported by the Institute of Transformative Bio-Molecules of Nagoya University and the Japan Advanced Plant Science Network.

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References 1. Sato A, Toyooka K, Okamoto T (2010) Asymmetric cell division of rice zygotes located in embryo sac and produced by in vitro fertilization. Sex Plant Reprod 23(3):211–217. https://doi.org/10.1007/s00497-009-01299 2. Sakakibara K, Reisewitz P, Aoyama T, Friedrich T, Ando S, Sato Y, Tamada Y, Nishiyama T, Hiwatashi Y, Kurata T, Ishikawa M, Deguchi H, Rensing SA, Werr W, Murata T, Hasebe M, Laux T (2014) WOX13-like genes are required for reprogramming of leaf and protoplast cells into stem cells in the moss Physcomitrella patens. Development 141(8):1660–1670. https://doi.org/ 10.1242/dev.097444 3. He YC, He YQ, Qu LH, Sun MX, Yang HY (2007) Tobacco zygotic embryogenesis in vitro: the original cell wall of the zygote is essential for maintenance of cell polarity, the apical-basal axis and typical suspensor formation. Plant J 49(3):515–527. https://doi.org/ 10.1111/j.1365-313X.2006.02970.x 4. Natesh S, Rau MA (1984) The embryo. In: Johri BM (ed) Embryology of angiosperms. Springer Berlin Heidelberg, Berlin, Heidelberg, pp 377–443. https://doi.org/10.1007/ 978-3-642-69302-1_8 5. Mansfield SG, Briarty LG, Erni S (1991) Early embryogenesis in Arabidopsis thaliana. I. The mature embryo sac. Can J Bot 69:447–460 6. Juergens G, Mayer U (1994) Arabidopsis. In: Bard J (ed) Embryos: colour atlas of development. Wolfe, London 7. Mansfield SG, Briarty LG (1991) Early embryogenesis in Arabidopsis thaliana. II. The developing embryo. Can J Bot 69:461–476 8. Yoshida S, Barbier de Reuille P, Lane B, Bassel GW, Prusinkiewicz P, Smith RS, Weijers D (2014) Genetic control of plant development by overriding a geometric division rule. Dev Cell 29(1):75–87. https://doi.org/10.1016/ j.devcel.2014.02.002 9. Gooh K, Ueda M, Aruga K, Park J, Arata H, Higashiyama T, Kurihara D (2015) Live-cell

imaging and optical manipulation of Arabidopsis early embryogenesis. Dev Cell 34 (2):242–251. https://doi.org/10.1016/j. devcel.2015.06.008 10. Mizuta Y, Kurihara D, Higashiyama T (2015) Two-photon imaging with longer wavelength excitation in intact Arabidopsis tissues. Protoplasma 252(5):1231–1240. https://doi.org/ 10.1007/s00709-014-0754-5 11. Kurihara D, Kimata Y, Higashiyama T, Ueda M (2017) In vitro ovule cultivation for live-cell imaging of zygote polarization and embryo patterning in Arabidopsis thaliana. J Vis Exp 127. https://doi.org/10.3791/55975 12. Kimata Y, Higaki T, Kawashima T, Kurihara D, Sato Y, Yamada T, Hasezawa S, Berger F, Higashiyama T, Ueda M (2016) Cytoskeleton dynamics control the first asymmetric cell division in Arabidopsis zygote. Proc Natl Acad Sci U S A 113(49):14157–14162. https://doi. org/10.1073/pnas.1613979113 13. Park J, Kurihara D, Higashiyama T, Arata H (2014) Fabrication of microcage arrays to fix plant ovules for long-term live imaging and observation. Sens Actuators B Chem 191:178–185. https://doi.org/10.1016/j. snb.2013.09.060 14. Nambo M, Kurihara D, Yamada T, NishiwakiOhkawa T, Kadofusa N, Kimata Y, Kuwata K, Umeda M, Ueda M (2016) Combination of synthetic chemistry and live-cell imaging identified a rapid cell division inhibitor in tobacco and Arabidopsis thaliana. Plant Cell Physiol 57 (11):2255–2268. https://doi.org/10.1093/ pcp/pcw140 15. Schneider CA, Rasband WS, Eliceiri KW (2012) NIH image to ImageJ: 25 years of image analysis. Nat Methods 9(7):671–675 16. Schindelin J, Arganda-Carreras I, Frise E, Kaynig V, Longair M, Pietzsch T, Preibisch S, Rueden C, Saalfeld S, Schmid B, Tinevez JY, White DJ, Hartenstein V, Eliceiri K, Tomancak P, Cardona A (2012) Fiji: an opensource platform for biological-image analysis. Nat Methods 9(7):676–682. https://doi.org/ 10.1038/nmeth.2019

Chapter 5 Analyzing Subcellular Reorganization During Early Arabidopsis Embryogenesis Using Fluorescent Markers Che-Yang Liao and Dolf Weijers Abstract Virtually all growth, developmental, physiological, and defense responses in plants are accompanied by reorganization of subcellular structures to enable altered cellular growth, differentiation or function. Visualizing cellular reorganization is therefore critical to understand plant biology at the cellular scale. Fluorescently labeled markers for organelles, or for cellular components are widely used in combination with confocal microscopy to visualize cellular reorganization. Early during plant embryogenesis, the precursors for all major tissues of the seedling are established, and in Arabidopsis, this entails a set of nearly invariant switches in cell division orientation and directional cell expansion. Given that these cellular reorganization events are genetically regulated and coupled to formative events in plant development, they offer a good model to understand the genetic control of cellular reorganization in plant development. Until recently, it has been challenging to visualize subcellular structures in the early Arabidopsis embryo for two reasons: embryos are deeply embedded in seed coat and fruit, and in addition, no dedicated fluorescent markers, expressed in the embryo, were available. We recently established both an imaging approach and a set of markers for the early Arabidopsis embryo. Here, we describe a detailed protocol to use these new tools in imaging cellular reorganization. Key words Arabidopsis thaliana, Embryogenesis, Subcellular structure, Fluorescent protein, Confocal microscopy

1

Introduction The process of embryogenesis of the flowering plant Arabidopsis thaliana provides an ideal model for addressing the fundamental question of how a single cell can give rise to a multicellular organism consisting of various cell types arranged in a precise pattern [1]. The connection between cell identity and gene activity has been the main focus of studies focusing on mechanisms underlying pattern formation during Arabidopsis early embryogenesis. As a consequence, essential techniques for analyzing cellular pattern and gene activity have been established [2–4]. However, the subcellular reorganization during oriented cell division and cell type specification, and their connection to the genetically controlled

Martin Bayer (ed.), Plant Embryogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2122, https://doi.org/10.1007/978-1-0716-0342-0_5, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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pattern formation process during early embryogenesis, remains to be discovered. While genetic tools and imaging methods to dissect cell biology underlying various developmental processes have been established, three challenges obstruct the direct examination of subcellular events during embryogenesis. Firstly, most established fluorescent-protein based reporters labeling specific cellular structures are driven by constitutive promoters that have either no (e.g., CaMV 35S promoter [5, 6];) or weak (e.g. pUBQ10 [7, 8];) activity during early embryogenesis. Secondly, the embryo is encapsulated in the seed and fruit. While seed culture and live imaging of gene activity and cytoskeleton reorganization in zygote have been established [9, 10], imaging subcellular structures in more developed embryos drastically suffers from the refraction and scattering resulting from the development of the seed coat and endosperm (unpublished observation). Therefore, embryo extraction is necessary for imaging subcellular structures in embryos. A third challenge thus follows: while techniques for embryo extraction and cellular profile counterstaining for analysis of gene activity were well established [2, 3], they suffer from low embryo extraction efficiency and are not meant to preserve delicate subcellular structures, such as microtubule and actin cytoskeletons. To overcome these challenges, we recently developed and reported [11] a set of fluorescent protein reporters labeling cellular structures, and driven by an embryo-specific promoter. In addition, we optimized imaging procedures that aimed to preserve cellular structures [11]. In this chapter, we describe the detailed procedure including seed extraction, embryo isolation, microscopy specimen preparation, and key points on confocal microscope setting aim to preserve and document subcellular reorganization during early plant embryogenesis.

2 2.1

Materials Tools

1. Forceps for collecting silique. Working surface of the tip should be less than 1 mm wide and flat to collect silique. 2. 92 mm diameter petri dish with double-sided tape. Divide the surface of the double-sided tape into a 15
10 mm grid (Fig. 1a). 3. Double edged blade for safety razor. We use the ones for shaving, not industrial use (see Note 1). Preferably without grease. With the blade still in its paper sheath, bend the blade longitudinally in the middle several times to snap it in two. 4. Stereo microscope, 10
–40
magnification, with bottom illumination.

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Fig. 1 Modified tools for seed extraction. (a) Petri dish with adhesive doublesided tape. The cutting pattern for the cover of the double-sided tape is marked in red. (b) Needle is bent toward the reader; note that the opening of the needle (arrow) is also toward the reader

5. 0.4
15 mm needle mounted (by screw) on a 1 mL syringe. Bend the needle as in Fig. 1b, note the orientation of the needle (see Note 2). 6. For collecting seeds containing 1-cell to globular stage embryos: Jeweler’s forceps, straight tip with smooth working surface, tip dimension: 0.1
0.06 mm or smaller (see Note 3). 7. For collecting seeds containing globular to torpedo stage embryos: 0.6
15 mm needle screw mounted on 1 mL syringe. Dip the tip of the needle in superglue to seal the channel of the needle. Remove excessive glue with a utility knife or sand paper after the glue is dried. 8. 92 mm diameter petri dish lid. 9. 200 μL pipette tip. 10. P100 pipette. 11. Timer. 12. Permanent marker, fine. 13. Coverslips, 24
50 mm. 14. Microscopy slide. 15. Tissues or paper towel. 16. Surgical glove. 17. Confocal microscope. Upright confocal microscope with UV (405 nm) for SR2200 and other light source for the fluorescent protein of choice.

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Mounting Media

Prepare all solutions with demineralized water, if not specified otherwise. 1. 20% glucose solution: dissolve 20 g glucose in 100 mL demineralized water. Autoclave. Make 500 μL aliquots in microcentrifuge tubes stored in 20  C. 2. 400 mM PIPES (piperazine-N,N0 -bis(2-ethanesulfonic acid)) stock solution: add 24.2 g PIPES with 60 mL demineralized water while adjusting pH to 6.8 with 10 N NaOH. Add demineralized water to final volume of 100 mL. Autoclave and store at room temperature. 3. 100 mM EGTA (ethylene glycol-bis(β-aminoethyl ether)-N,N, N0 ,N0 -tetraacetic acid): dissolve 3.8 g EGTA with 60 mL demineralized water while adjusting pH to 7 with 10 N NaOH. Add demineralized water to final volume of 100 mL. Store at room temperature. 4. 40 mM MgCl2: dissolve 0.812 g MgCl2·6H2O with 90 mL demineralized water and add demineralized water to final volume of 100 mL. Autoclave and store at room temperature. 5. 2
microtubule stabilizing buffer (2
MTSB): 200 mM PIPES, 10 mM EGTA, 4 mM MgCl2. Diluted with demineralized water. Make 500 μL aliquot in microcentrifuge tubes and store at 20  C. 6. 10% SR2200 (SCRI Renaissance 2200): Mix 50 μL of SR2200 with 500 μL demineralized water. Make 20 μL aliquot in microcentrifuge tubes and store at 20  C. 7. 10 mM Taxol (Paclitaxel): Dissolve in 100% DMSO (dimethyl sulfoxide) (see Note 4). Make 20 μL aliquot in microcentrifuge tubes and store at 20  C. 8. Embryo general mounting solution (EGM): 10% glucose solution. 9. Embryo general counterstaining solution (EGC): 10% glucose +0.1% SR2200. 10. Embryo microtubule mounting solution (EMTM): 10% glucose, 10 μM Taxol in 1
MTSB. 11. Embryo microtubule counterstaining solution (EMTC): 10% glucose, 10 μM Taxol, 0.1% SR2200 in 1
MTSB.

3

Methods Every operation described below is from the perspective of a righthanded operator.

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3.1 Transferring Seeds from the Silique to the Microscope Slide

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1. Water the plant the day before imaging (see Note 5). 2. One will need stable hands, so avoid any activity that could increase heart rate or strain the arms prior to sample preparation. 3. With the forceps (not jeweler’s), place the silique on the double-sided adhesive tape with the replum facing upward and the stigma pointing toward the operator (Fig. 2a). Place the stigma on the double-sided tape and secure it with a finger gently and “pull” the forceps gently to gently “stretch” the silique before placing the pedicel end on the double-sided tape (see Note 6). 4. Tap the silique gently along the axis of the replum with the index finger. This gives the silique firm contact with the double-sided tape to secure its position. 5. Place the petri dish under a stereomicroscope with the stigma pointing toward the operator. Following steps will be described from the perspective via stereomicroscope with 10x magnification. The prime directive is to maintain the intact structure of the seed. Embryos can hardly be extracted from damaged seed. 6. Switch on the bottom illumination of the stereomicroscope (see Note 7). 7. Secure the safety razor blade between the thumb and middle finger with index finger on the blade for fine control (Fig. 2b), cut through both ends of the silique (Fig. 2c, cut #1 and #2) (see Note 8). 8. From the pedicel to the stigma, cut through only one side of the right valve as close to the replum as possible with the razor blade (Fig. 2c, cut #3) via bending the thumb and index finger (see Note 9). 9. Hold the 0.4
15 mm needle like a pen with the point of the needle pointing downward perpendicular to the valves, opening the right valve by poking the point of the needle into opening made from cut #3 and push the valve to the right with the thumb. Secure the valve on the double-sided tape through continuing moving the needle to the right side. The geometry of the valve will make itself against the double-sided tape providing additional stability for the following operations (Fig. 2d). 10. Rotate the petri dish so the stigma is pointing away from the operator, repeat Subheading 3.1, steps 8 and 9 to open another valve (Fig. 2e). 11. Rotate the petri dish so the stigma is pointing toward from the operator.

Fig. 2 Seed extraction. (a) The silique secured on the adhesive double-sided tape with its replum facing toward the reader. (b) Holding the shaving razor blade. The arrow indicates the point of contact of the razor blade with the valve. (c) Cuts to open the valve. The arrow, open arrow, and arrowhead indicate cut #1, cut #2, and cut #3, respectively. (d) Silique with its right valve opened. (e) Silique with both valves opened. (f) Holding the jeweler’s forceps. The arrow indicates the area of working surface contacts the seed. (g) Positions of the working surface of the jeweler’s forceps during seed extraction. The black triangles indicate the working surface when extracting seed in the right or left valve. Note when extracting the seeds in the left valve, the point of the working surface is under the seed. The open arrows indicate the directions of the movement of the jeweler’s forceps. (h) Extracted seed on the working surface of the jeweler’s forceps. (i) Extracted seeds in the mounting medium. Note all seeds only aggregate at the same plane as the result of transferring them one-byone. (j) Petri dish covering the whole specimen to minimize mounting medium evaporation

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12. Take a microscope slide and place a drop of the mounting medium in the following volume corresponding to the seeds’ developmental stage: 2–16 cell, 10 μL; 16-early heart stage, 15 μL; Heart to torpedo 20 μL (see Note 10). Keep the medium in a shape of a droplet. 13. Place the microscope slide with the mounting medium droplet where it can be easily accessible and well lit when working with the stereomicroscope but wouldn’t be disturbed during the operation. The well-lit surface of the droplet makes it easier to transfer the extracted seeds to the droplet. 14. Back to the stereomicroscope. Hold the jeweler’s forceps like a pen with one working surface facing away from the operator (Fig. 2f). 15. Transfer each seed in the right valve one-by-one to the mounting medium. Start from the right valve first, from the stigma toward the pedicel (see Note 11). Place forceps’ tip between the septum and the seed, gently move the tip downward to cut the funiculus, then with the working surface of the forceps against the seed, move the working surface with the seed outward (toward 2 o’clock direction) to disconnect the seed from the septum (Fig. 2g). Do not move the seed outside of the valve because once the seed contacts with the double-sided tape, any attempt to lift it from the double-sided tape is likely to damage the seed. 16. Transfer the seeds on the forceps tip (Fig. 2h) to the mounting medium on the slide by touching the seeds to the surface of the droplet (see Note 12). Once the seed is transferred to the droplet (Fig. 2i), carefully wipe tip of the forceps’ tip with the tissue/paper towel to ensure a dry working surface. 17. Transfer each seed in the left valve one-by-one, from the stigma toward the pedicel, to the mounting medium. Place forceps’ tip under the funiculus between the septum and the seed; gently move the tip to the right. The septum will be folded to the right with the forceps now on the funiculus (Fig. 2h). Gently move the tip downward to cut the funiculus, then with the working surface of the forceps against the seed, move the working surface with the seed outward (toward 2 o’clock direction) to disconnect the seed from the septum. 18. Transfer the seed to the mounting medium as described in Subheading 3.1, step 16. 19. Repeat Subheading 3.1, steps 17 and 18 until all seeds are transferred. 20. Add mounting medium over the first mounting medium droplet counting seeds in the following volume corresponding to

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the seeds’ developmental stage: 2–16 cells, 5 μL; 16-early heart stage, 7.5 μL; heart to torpedo 10 μL (see Note 13). 21. Cover the slide with the petri dish lid (Fig. 2j) and incubate the seed in the mounting medium for 15 min (see Note 14). 3.2 Embryo Extraction

1. At the left side of the mounting medium containing the extracted seeds, 5 mm apart (Fig. 3a), add staining medium in the following volume corresponding to the seeds’ developmental stage: 2–16 cell, 15 μL; 16-early heart stage, 22.5 μL; Heart to torpedo 30 μL (see Note 15). 2. Put on a surgical glove on the left hand if Taxol is used (see Note 16). 3. With the forceps (not jeweler’s) in the right hand, hold the coverslip with the forceps and position the left edge of coverslip, secured by the left hand, on the left side of the droplet of the staining medium with the angle between the coverslip and the slide at 45 (Fig. 3a). 4. Open the forceps but keep contact with the coverslip to support the coverslip with the tip of the forceps. 5. Lower the forceps slowly to close the coverslip without introducing air bubbles as in (Fig. 3a) (see Note 17). 6. Place the slide under the stereomicroscope. Switch on the bottom illumination. 7. Secure the slide with left thumb and middle finger and secure the coverslip with the left index finger a shown in (Fig. 3c) (see Note 18). 8. Start the timer, the specimen can be used for imaging for an hour from now on. 9. Secure a 200 μL pipette tip in the right hand with the thumb, index and middle finger (Fig. 3b). The index finger guides and provides the downward force while the thumb and middle finger support the 200 μL pipette tip. 10. The following steps will be described from the perspective via stereo microscope with 40
magnification. The prime directive is to extract embryo from each seed one-by-one through visual confirmation. 11. Disregard the orientation of the seed, position the tip at the 4–5 o’clock position at the proximity of the seed (see Note 19). 12. Apply downward force on the tip until seeing the seed deforms (getting bigger, pressed) (see Note 20). 13. Maintain the same downward force and slide the tip toward the 10 o’clock position to the edge of the seed. An expulsion of materials from the seed, including an embryo, would be expected indicating successful embryo extraction (Fig. 3d). If no expulsion occurs, repeat twice without applying more

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Fig. 3 Specimen preparation and embryo extraction. (a) Position of the staining medium (left side of the specimen) and placement of the coverslip. (b) Holding the pipette tip. (c) Hands position during embryo extraction. Note the position of the thumb, index, and middle finger of the left hand. (d) Seed with extracted embryo. The green area is the suitable area to apply pressure by the pipette tip during embryo extraction. (e) The imaging area marked by the permanent marker in the backside of the specimen. (f) Mounting the specimen on the microscope stage. Note the stage clip is on the coverslip and the relative position between the immersion medium and the marked imaging area (light blue quadrilateral). The operation of the objective lens must be remaining inside the marked imaging area

downward force than in the first attempt, but change the position and orientation of the slide, for example, change from 4-to-10-o’clock to 5-to-11-o’clock. If an expulsion occurs but with no embryo in the expelled materials or no expulsion after three attempts, move on to the next seed (see Note 21).

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14. Repeat Subheading 3.2, steps 12 and 13 till all seeds are processed. 15. Turn the specimen to vertical position so the coverslip is facing away from the operator. 16. Identify the position of the seeds and mark this area with permanent marker on the slide (the coverslip remains facing away from the operator) (Fig. 3e) (see Note 22). 3.3 Imaging the Subcellular Structure Using an Upright Confocal Microscope

1. Mount the specimen on the specimen stage of the microscope. Lift the working surface of the stage clip, position it above the left or right edge of the coverslip, and gently lower it to secure the coverslip (Fig. 3f) (see Note 23). 2. Add immersion medium on one of the seeds close to the mark (Fig. 3f) and raise the stage till the immersion medium contacts the objective lens. 3. Find the embryo and acquire images as ordinary imaging procedure with following recommendation (see Note 24). 4. Use photon counting mode: The combination of live sample and repetitive scanning to generate optical stacks necessitates low laser input to prevent phototoxicity and photobleaching while the minute nature of subcellular structure and weaker promoter activity (while pWOX2 is considered strong embryo promoter, it is weaker than the commonly used p35S and pUBQ10 in more developed organs, such as roots) generates small quantities of photons per pixel. Therefore, photon counting is the preferred detector operation mode. Setting a discriminator equal to the background (an area without biological material), immensely increase image contrast and reduces background noise.

4

Notes 1. The difference between the blade for shaving and for industrial use is their sharpness and thickness. The sharper and thinner nature of razor for shaving make it ideal for cutting the silique valves open without damaging the seeds. 2. This orientation allows the operator to see the exact position of the point of the needle as the actual point of the needle will be at the left-hand side when the tip is pointing downward perpendicularly. 3. Essential for collecting seeds containing 1–16 cell embryo. Smaller tip dimension such as 0.05
0.01 mm or 0.025
0.005 mm will be preferred for seeds with developmental defect or containing 1–4 cell embryos.

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4. We ordered 5 mg Taxol and add 585 μL of 100% DMSO directly to the stock vial with a 1000 μL pipette tip whose first 5 mm point was cut off to accommodate the viscosity of 100% DMSO. Gently pipet up and down ten times for mixing. Alternatively, 50% DMSO dissolved in demineralized water can be used. 5. This ensures firm valves and facilitates dissecting the siliques as firmer valves offer stronger mechanical feedback when cut by razor blade. Watering the plant a day in advance also ensures that seeds are turgid and increases the success rate of embryo extraction. 6. For wild types or mutants without a seed abortion phenotype, one silique is generally sufficient for each specimen. Otherwise, no more than 50 seeds per specimen should be used. This ensures that seeds are well separated from each other and provides sufficient room not occupied by the seed to be operated by the pipette tip during embryo extraction. 7. The light from beneath makes the silique, especially the younger siliques containing seeds with embryos younger than heart stage, transparent, making each seed in the silique more distinguishable so that the operator can evade the seeds when necessary. 8. Keep the fingers in one half of the razor blade to maintain clear view in stereomicroscope. Make the cuts 0.5 mm from both ends of the valves. This may eliminate 2–4 seeds, but will make it easier to open the valves as these cuts remove the more tapered parts of the silique. 9. With the wrist and forearm rest on the table provides a stable platform for the finger movement. Try different path of the razor blade (with the Petri Dish and silique in the corresponding orientation), for example 9–3 o’clock, 11–5 o’clock, or 12–6 o’clock, to find the most stable movement for each operator. The safety razor blade is so sharp that as long as one feels the contact between the razor blade with the valve, the blade is making the cut already. If not, replace the razor blade. 10. These volumes are guidelines as the combined total volume of mounting and staining medium depends on the size of the seed, which is correlated to embryo’s developmental stage in wild type plants. This step requires 1/3 of the combined total volume of mounting and staining medium. Two criteria of the optimal combined total volume of mounting and staining medium are: firstly, sufficient liquid between the coverslip and the slide so no bubble occurs during embryo extraction; secondly, no liquid flowing outside the coverslip after embryo extraction.

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11. Start from the stigma end of the right valve as the funiculi of the seeds in the right valve is most accessible with this orientation of working. For seeds containing heart to torpedo stage embryos, 0.6
15 mm needle is preferable with larger working surface to accommodate larger seeds. 12. Do not dip the whole tip of the needle/forceps into the droplet as this may encapsulate the seed within a small droplet on the tip instead of transfer the seeds to the mounting medium. Keep the needle/forceps dry as seeds are prone to stick to wet surface and this will only make it difficult to transfer the seeds to the mounting medium. 13. Similar to Subheading 3.1, step 13, these numbers are a guideline, but shall be 1/6 of the optimal combined total volume of mounting and staining medium. 14. The lid prevents the minute volume of the mounting medium being evaporated causing change in the osmotic pressure of the mounting medium. The incubation makes the seed easier to be extracted; reducing the force required to push the embryo out of the seed and allows Taxol to enter the seed to stabilize microtubule. 15. Similar to Subheading 3.1, step 13, these numbers are a guideline, but shall be 1/2 of the optimal combined total volume of mounting and staining medium. 16. Surgical gloves also increase friction between the finger and the glass and facilitate to secure the specimen. 17. It is crucial to have no air bubble as the air bubble will become an air cushion between the coverslip and the slide that hinders embryo extraction. In addition, the movement of the bubble could cause movement of extracted embryos during imaging. There will be some room left at the right edge of the coverslip that is not contacting the slide. This is intentional and will be sealed in the following steps. 18. The thumb and the middle finger provide support for the whole specimen. The index finger provides the support to secure the coverslip to prevent the movement of the coverslip during the popping. This is crucial as any horizontal coverslip movement will damage the seed and make it impossible to extract the embryo from the seed. Maintain left index finger position supporting the coverslip until the end of embryo extraction. 19. Never position the tip directly above the seed as the following pressure will likely squash the seed but not “pop” the seed. 20. Pay attention to the seed, especially those containing an embryo from globular or later stages. The seed could burst at this point if an expulsion of materials from the seed, including an embryo, is observed.

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21. Attempting to extract embryos from damaged seeds is futile because it is impossible to build up sufficient pressure in the seed to expel the embryo. The expected efficiency of embryo extraction is between 25% and 50%. 22. This marks the area for objective lens operation. Each side of this marked area must be at least 8 mm, or the diameter of immersion medium when the objective lens is at its operating position against the specimen, away from the edge of the coverslip. This mark serves as a landmark when searching for the embryos and prevents the objective lens being too close to the edge of the coverslip causing the contact/mixture of the immersion medium with the mounting medium that could drastically alter the property, such as osmotic pressure, of the environment surrounding the extracted embryo. 23. This prevents any horizontal movement of the coverslip that could damage the embryos. 24. Always take optical stacks: The isodiametric geometry of the embryo makes 3D reconstruction necessary for the analysis of subcellular organization. References 1. Wendrich JR, Weijers D (2013) The Arabidopsis embryo as a miniature morphogenesis model. New Phytol 199:14–25 2. Llavata-Peris C, Lokerse A, Mo¨ller B, De Rybel B, Weijers D (2013) Methods Mol Biol 959:137–148 3. Musielak TJ, Schenkel L, Kolb M, Henschen A, Bayer M (2015) A simple and versatile cell wall staining protocol to study plant reproduction. Plant Reprod 28:161–169. https://doi.org/ 10.1007/s00497-015-0267-1 4. Yoshida S et al (2014) Genetic control of plant development by overriding a geometric division rule. Dev Cell 29:75–87. https://doi. org/10.1016/j.devcel.2014.02.002 5. Benfey PN, Ren L, Chua NH (1990) Tissuespecific expression from CaMV 35S enhancer subdomains in early stages of plant development. EMBO J 9:1677–1684 6. Vo¨lker A, Stierhof YD, Ju¨rgens G (2001) Cell cycle-independent expression of the Arabidopsis cytokinesis-specific syntaxin KNOLLE results in mistargeting to the membrane and is not sufficient for cytokinesis. J Cell Sci 114:3001–3012

7. Sun CW, Callis J (1997) Independent modulation of Arabidopsis thaliana polyubiquitin mRNAs in different organs and in response to environmental changes. Plant J 11:1017–1027. https://doi.org/10.1046/j. 1365-313X.1997.11051017.x 8. Norris SR, Meyer SE, Callis J (1993) The intron of Arabidopsis thaliana polyubiquitin genes is conserved in location and is a quantitative determinant of chimeric gene expression. Plant Mol Biol 21:895–906. https://doi.org/ 10.1007/BF00027120 9. Gooh K et al (2015) Live-cell imaging and optical manipulation of Arabidopsis early embryogenesis. Dev Cell 34:242–251. https://doi.org/10.1016/j.devcel.2015.06. 008 10. Sauer M, Friml J (2008) In vitro culture of Arabidopsis embryos. Methods Mol Biol 427:71–76 11. Liao CY, Weijers D (2018) A toolkit for studying cellular reorganization during early embryogenesis in Arabidopsis thaliana. Plant J 93:963–976. https://doi.org/10.1111/tpj. 13841

Chapter 6 From Stained Plant Tissues to Quantitative Cell Segmentation Analysis with MorphoGraphX Merijn Kerstens, Soeren Strauss, Richard Smith, and Viola Willemsen Abstract Development and growth of plant organs is determined by a myriad of molecular processes that occur in each individual cell. As a direct consequence of these processes, cells alter in size and shape. They therefore serve as excellent parameters to thoroughly understand gene function. However, conventional single-plane analyses fail to accurately capture cell metrics. Here, we present a comprehensive illustrated guide that demonstrates how SCRI Renaissance 2200 staining of Arabidopsis thaliana embryos and roots can be combined with the open-source application MorphoGraphX to quantify cell parameters in 3D. We compare this staining method with other common staining techniques and provide examples of embryo and root tissue segmentation. With our novel approach, subtle single-cell phenotypes can be identified in their native context, providing new possibilities to dissect gene networks. Key words Embryos, Roots, Lateral roots, 3D imaging, SCRI Renaissance, MorphoGraphX, 3D segmentation, Cell volume

1 1.1

Introduction Information in 3D

Faithful development of living organisms necessitates precise coordination of all cellular processes in both space and time. Many studies have been performed to elucidate the genetic principles, mechanisms, and dynamics of fundamental developmental events. Concurrently, advances in genome and transcriptome sequencing technologies have allowed to dissect the genetic networks driving these processes. The size and shape of plant organs are wellestablished readouts from these networks, but it remains very challenging to accurately extract such metrics on a cellular level by single-plane imaging. For instance, a conventional 2D analysis on erratic cell shapes or cell division planes is prone to misinterpretation due to the layered nature of tissues. Hence, 3D reconstitutions of organs are indispensable to analyze their size and shape in a precise and quantitative manner. The volume of cells that form a tissue, as well as cell wall orientation, area, and position, is an

Martin Bayer (ed.), Plant Embryogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2122, https://doi.org/10.1007/978-1-0716-0342-0_6, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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attractive candidate parameter to examine with this novel approach in order to characterize mutation phenotypes or gene-editing effects. Complementing our existing knowledge, such crucial analyses will lead to new insights on how genetic networks contribute to the multiscale progressions from cellular dimensions to organ shape and, ultimately, whole plant stature. The first time we were able to visualize, analyze, and manipulate root tissues using optical sections by making use of confocal scanning laser microscopy (CLSM), is almost 25 years ago [1]. But, in the past decades, existing techniques have been continuously improved. This has facilitated the extraction of more information from the images, leading to rapid uncovering of various molecular networks underlying plant growth and development. Integrating these data into the spatial and temporal context of dynamic organ growth remains a technical challenge. The introduction of MorphoGraphX (MGX), which is an open-source application for the visualization and analysis of 3D biological datasets obtained by confocal imaging, was a major breakthrough [2]. Although the program makes it possible to analyze organs at a single-cell resolution, the learning curve can be discouraging. Furthermore, obtaining high-quality images that can be used in MGX can also be challenging. Here, we will describe a comparison between different established staining techniques, and we will provide a quick guide on using MGX with standard settings. In this protocol we demonstrate how the program can be used for segmenting cells in Arabidopsis thaliana embryos, seedling roots, and various stages of lateral root formation (Fig. 1). 1.2 Comparison Between Different Staining Techniques

Using a method that can stain the cell wall or plasma membrane sufficiently to obtain good quality high resolution images of your tissue is of major importance to obtain reliable 3D information. To find the optimal staining protocol we have qualitatively compared different staining techniques, and our findings are described in the following paragraph. Conventional propidium iodide (PI) staining provides a nonfixating approach to accurately and rapidly stain undamaged cell walls. This allows living tissues to be visualized in near-native conditions [1]. Consequently, PI staining can be used to contextualize fluorescence to observe gene expression and protein localization. While tissue penetration is swift (seconds) in thin specimen such as root tips and lateral roots, cell files situated more centrally in thicker regions require prolonged staining. Even then, resolution in these areas is low when median cross sections are made. An additional problem that can occur with PI is that dead cells are extensively stained due to specimen handling during the imaging process. Even a small lesion allows PI to permeate the cell, which presents a serious issue to a segmentation analysis with MGX. In order to improve resolution and overcome the dead cell problem, an

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Fig. 1 Segmentation of embryos (a), individual root cell files (b), and lateral roots (c) of A. thaliana Col-0 plants using MGX and Renaissance staining. (a) One-cell stage, two-cell stage, 16-cell stage (from left to right), (b) segmented epidermis (left), segmented cortex (right), (c) stage I lateral root (left), stage II lateral root (right)

adapted PI staining called modified pseudo-Schiff propidium iodide (mPS-PI) [3] has been developed. This method greatly improves tissue penetration and can also visualize amyloplasts, a convenient tool for diagnosing differentiating meristems (Fig. 2). However, staining takes up to 3 days, and the resulting tissue is extremely fragile. This can lead to cell wall breakage, which again complicates segmentation (Fig. 3). With the introduction of the versatile cell wall staining SCRI Renaissance 2200 [4, 5], the abovementioned problems could be solved. Additionally, the SCRI Renaissance staining can be combined with a wide variety of fluorescent proteins, which can be useful for studying various aspects of plant development. We have compared the different staining and have listed the cons and pros in the following table (Table 1).

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Fig. 2 Cross sections of primary A. thaliana Col-0 roots (a–c) and longitudinal sections (d–f). (a, d) PI staining; (b, e) Renaissance staining; (c, f) Schiff staining. Imaged derived from MGX

Fig. 3 Cell wall disruption by handling fragile mPS-PI-stained root material. (a) Main stack view in MGX, (b) resulting segmentation errors (Asterisks mark the lesion sites.)

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Table 1 Comparison between different staining techniques

Staining name

Shape of root cross Procedure Allows section Resolution Fixation Preservation duration fluorescence Disadvantage

PI

Round

Low

No

No

Instantly

Yes

Stains dead cells, low penetration

mPS-PI

Flat

High

Yes

Yes

3–5 days

No

Cell wall breakage

High

Yes

Yes

1–3 h

Yes

Sometimes stains nuclei

Renaissance Oval

To analyze tissues, organs, or complete organisms, it is important that the staining and imaging procedures affect the shapes of the cells as little as possible. The abovementioned staining methods have different effects on the shapes, which are listed in Table 1 represented in Fig. 2.

2

Materials

2.1 Chemicals and Solutions

1. Propidium iodide staining solution [1]: 10 μg/mL propidium iodide (PI) in water. 2. SCRI 2200 Renaissance staining solution (adapted from [5]): 0.1% SCRI Renaissance Staining 2200 Chemicals, 1% DMSO, 0.05% Triton X-100, 5% glycerol, 0.13 M paraformaldehyde in PBS buffer, pH 8.0 (see Note 1). 3. Fixative solution (adjusted from [3]): 10% Acetic acid, 50% methanol. 4. 1% periodic acid solution. 5. mPS-PI Schiff reagent solution (adjusted from [3]): 100 mM sodium metabisulfite, 0.15 N HCl. This solution can be stored at room temperature for a long time. PI must be freshly added to a final concentration of 0.1 mg/mL. 6. Chloral hydrate solution: 4 g chloral hydrate, 1 mL glycerol, and 2 mL water. Keep at 4  C. 7. Hoyer’s solution: 30 g gum arabic, 200 g chloral hydrate, 20 g glycerol, and 50 mL water.

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Equipment

1. Confocal laser microscope with 405 nm excitation laser. 2. Computer requirements for MorphoGraphX: MorphoGraphX (MGX) requires a computer running Windows or Linux with enough main memory (at least 8 GB of RAM) and a graphics card supporting OpenGL 3. A multicore CPU and an nVIDIA graphics card with CUDA support and 4 GB or more video memory are beneficial for the processing of large images.

3

Methods

3.1 Preparation for Microscopy

1. Collect material into fixative solution (10% acetic acid, 50% methanol solution) and keep it at least for 12 h at 4  C.

3.1.1 Schiff Staining

2. Rinse the tissue with water. 3. Remove the water and add 1% periodic acid for 40 min to 1 h at RT. Work in fume hood from this step until step 9. 4. Remove the periodic acid and wash the sample with water. 5. Incubate samples in Schiff reagent solution with propidium iodide for 1–2 h at RT or until the material stains pink. 6. Place the material (root tip) on the microscope slides (after the Chloral Hydrate treatment, material is very sensitive, so be gentle). 7. Add a drop of chloral hydrate solution (enough to cover samples but not too much) and incubate O/N at RT in a closed environment to prevent drying out. In case of root tips, this step can be limited to few hours of incubation or it can also be omitted. 8. Remove excess chloral hydrate with a piece of paper. Samples can be correctly oriented at this step. 9. Add several drops of the mounting medium Hoyer’s solution and place coverslips. 10. Leave slides in the dark for at least 3 days to allow the mounting solution to set properly.

3.1.2 Renaissance Staining

1. Collect your plant material and incubate the tissue according to Table 2 in SCRI Renaissance 2200 staining solution. (a) For roots and lateral roots: The staining solution tends to run off microscope slides. Therefore, clean the seedlings in water prior to positioning them in 100 μL of SCRI Renaissance staining on an objective slide. Gently (use tweezers) lower an 18  18 mm coverslip onto the seedling.

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Table 2 Plant tissue characteristics for SCRI Renaissance staining Plant tissue

Age

Incubation time

Embryo

Flowering plant

None

Root

3–5 dpg

1.5 h

Lateral root

7–10 dpg

O/N

(b) For embryos: ovules have to be isolated from siliques directly in SCRI Renaissance 2200 solution on an objective slide. Pop out the embryos from the ovules by applying gentle pressure to the coverslip. 3.2 Confocal Microscopy

As an example, we have thoroughly explained the imaging for the Zeiss LSM 710 in Fig. 4. 1. Use DAPI settings on the confocal—an excitation wavelength of 405 nm and emission filter of 410–514 nm. Change the bit depth to “16-bit”. Use a pinhole of 1 Airy Unit (AU) and adjust the laser power and digital/master gain to a level at which the median of the tissue clearly displays all cell walls, but the upper epidermis is not subjected to severe overexposure. Especially in roots, the lower epidermis will always be quite dark. 2. Check “Z-stack” in the upper left corner, expand the Z-stack window under “Multidimensional acquisition” and change the interval to 0.50 μm. Focus on a plane a few microns above the sample and press “Set First,” then focus on a plane a few microns below the sample and press “Set Last” (These settings are for a 40 objective). 3. Use a pixel dwell time of 6.30 μs by dragging the slider next to “Speed.” You can then initiate scanning by selecting “Start experiment” (see Note 2).

3.3 Image Processing and Analysis Using MorphoGraphX

After acquiring the images, we can now move on to process the imaged data using MorphoGraphX.

3.3.1 Getting Started with MorphoGraphX

1. The software is open-source and available on the website www. morphographx.org, where installation packages and the source code can be found [2]. A detailed installation and user guide can be found on the website in the “Help” section as well as in MGX under Help/User Manual.

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Check Z stack Will display “start experiment” prior to scanning

Speed slider

Bit depth

405 nm laser Laser power 1 Airy Unit button Master Gain Digital Gain

Set last plane

Interval; Z-resolution Range; depth of stack

Fig. 4 The Zeiss LSM 710 microscope settings are displayed in a screenshot

Set first plane

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Table 3 MGX tools and their symbols Tool name

Tool symbol

Anchor Delete Label in Volume Fill Label Main Stack Color Map Pick Label Select Connected Area Select Points in Mesh Voxel Edit

2. Upon start-up, MGX opens a graphical user interface (the main window) together with a console window in the background. The console window can show additional information and needs to remain open as long as MGX is running. 3. MGX can process two types of data: stacks and meshes. Stacks are 3D voxel images, whereas meshes consist of triangles of vertices to define a surface or a volume. Stacks typically represent the raw data, and meshes (that can be generated from stacks) are used for extended data analysis steps. 4. The main window consists of toolbars on the top (for stack processing) and the left side (for mesh processing) and a tab menu on the right side containing the tabs “Main,” “View” (for visualization options), and “Process” (see below). 5. MGX uses modular plug-ins (“processes”) categorized into folders to process data. They can be found under the “Process” tab in the main window. Processes often come with parameters that can be found in the bottom right corner after clicking on a process. To execute a process, double-click on it or click on the “Run” button in the top-right of the main window. 6. MGX does not have an “undo” function. We advise to back up your original data and frequently save your stacks and meshes. 7. The MGX symbols for the different tools used in this protocol are listed in Table 3. 3.3.2 Analysis of Embryo Data

1. Convert the confocal .lsm file to .TIFF with Fiji [6] by dragging your file into the Fiji GUI and save it in TIFF format using (“File > Save as > TIFF”). Now you should also check the voxel size of your image (“Info >Show Info > Voxel Size”).

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Fig. 5 Screenshots of MGX displaying the first steps of the segmentation procedure. (a) The main stack of a four-cell stage embryo. The green circles indicate the “Process” tab, the “Stack” sub-tab and the “Change Voxel Size” process within the “Canvas” folder, the blue circle highlights its associated parameters. (b) The main stack has been subjected to a Gaussian blur (purple circle), and afterward the Watershed process has been run (pink circle) with a level of 1500 (orange circle). The resulting segmentation can be seen in the main window; it is visible as a rectangular block. In order to make the data visible, the segmentation of the surrounding air (blue color) has to be deleted using the “Delete Label in Volume” tool and Alt+Left clicking on it (gray circles)

2. Start MGX and open your TIFF file by dragging it into the main window. This will open the file as “main” store. You can rotate the image by dragging the mouse holding the left mouse key, move the image by holding the right mouse key and zoom using the mouse wheel (see Notes 3 and 4). 3. Change the voxel size using the process “Stack > Canvas > Change Voxel Size” (in Fig. 5a, the green circle and blue circles) to the dimensions extracted from Fiji in the previous step. Wrong voxel sizes cause scale changes (see Note 5). 4. Apply a Gaussian blur of 0.3 μm in all dimensions, by running the process “Stack > Filter > Gaussian Blur Stack” (Fig. 5b, purple circle). The processed stack will now be transported to the “work stack” and turn blue. 5. Use the Auto-seeded Watershed process with a threshold of 1500–2000 (Process “Stack > ITK > Segmentation > ITK Watershed Auto Seeded”; Fig. 5b, pink circle). Your segmentation will result in a rectangular block around the tissue representing the empty outside space of the image. To make the segmented cells visible, delete this block by selecting the “Delete Label in Volume” tool in the stack tool bar and using Alt+Left click on the block (Fig. 5b, blue circles). Check your sample for errors in the segmentation (Figs. 6 and 7, see Notes 6 and 7).

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Fig. 6 Oversegmentation; the effect of different threshold values in the watershed process (a) threshold 1000: the sample is highly oversegmented. Almost all cells consist of several segments, (b) Threshold 2000: Slightly oversegmented, (c) Threshold 3000: slightly undersegmented: some cells are merged with their neighbors. This should be avoided as merged cells cannot be split up later on

6. Do not forget to frequently save your intermediate results as there is no undo function. To save a current state (use the upper toolbar!): (a) Save the main stack: “Stack > Stack 1 > Main > Save”. (b) Save the work stack: “Stack > Stack 1 > Work > Save”. (c) It is also possible to save a project file (∗.mgxv) via File > Save. This will save the locations of the currently opened stacks and meshes and all viewing and processing parameters. 7. Correct errors in the segmentation (see Fig. 7). 8. Once you are satisfied with the segmentation, create a cell mesh with the process “Mesh > Creation > Marching Cubes 3D” (cube size: use 1.0 μm). This will generate a triangular mesh of volumetric cells (Fig. 7f, see Note 8). 9. Ensure that the mesh is correct and that you cannot find any additional segmentation errors! You can use the clipping planes to scroll through the entire tissue cell by cell to see whether the segmentation matches the actual cells. If you do find errors, you have to go back to the Watershed (or an earlier saved stack) and to merge/delete cells (see Note 9). Save the mesh using the function “Mesh > Mesh 1 > Save” in the upper toolbar. The remaining analysis requires only the mesh. 10. To conceal the stacks, go to the “Main” tab and uncheck “Main” and “Work.” To make the mesh visible, check the “Surface” checkbox. Vertices, edges and selection of the mesh can be made visible using the “Mesh” checkbox and selection of the desired option in the drop-down menu (see Note 10).

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Fig. 7 Correction for oversegmentation errors. (a) The main stack view. (b) The segmented stack after using the watershed process. (c) The segmented stack after removing unnecessary cells with Alt+Left clicking on them using the “Voxel Edit” tool or the “Delete label in Volume” tool . (d) Check the outline of the cells with the main stack. (e) Correct the oversegmented cells with Alt+Left click using the “Pick Label” tool on one cell , and using Alt+Left click on the other cell using the “Fill Label” tool . Use the clipping planes and main stack reference to verify errors. (f) Mesh created from the corrected embryo. (g) Correction procedure on a single cell level: from the main stack it can be seen that there is one cell; (h) however, the watershed segmented it into two cells. (i) Segmentation after correction

11. We can now quantify the volume and the wall area of the cells using the process “Mesh > Heat Map > Heat Map Classic” selecting either “Volume” or “Area” as Heat Map Type and pressing “OK.” This will color the cells according to their value, and a scale bar will appear on the top left with the range of the heat map’s values (Fig. 8b). 12. After creating the heat map, the values for the individual cells can be saved to a csv-file using the process “Mesh > Heat Map > Heat Map Save” (see Note 11).

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Fig. 8 Quantifying the volume and the area of the embryonic cells. (a) Heat map coloring the cells according to their volume (as indicated by the scale bar), (b) Heat map of the cell wall area 3.4 Additional Analysis Tools Using MorphoGraphX 3.4.1 Changing the Color (and the Cell Label) of a Cell in a Mesh

MGX has numerous tools to change the visualization of stack and meshes. Here we introduce a few of them that might come in handy. 1. Select the cell you wish to relabel with Alt+Left click on the “Select Connected Area” tool symbol. Ensure that “Mesh” is checked in the “Main” tab! 2. Run the process “Mesh > Segmentation > Label Selected Vertices” with the desired label parameter. 3. The label number corresponding to a certain color can be found by going to the “Edit Labels” menu in the upper toolbar. You can change the given colors by double-clicking a certain color and then selecting a new one (see Note 12).

3.4.2 Changing the Color of the Main Stack

1. In the “Main” tab, open the “Main Stack Color Map” by clicking on next to “Main.” 2. Select your desired color in the drop-down menu titled “Predefined color maps.” Choose only the scaled color maps!

3.5 Image Analysis of Root Tissue Using MorphoGraphX 3.5.1 Segmentation of Root Images

The 3D Cell Atlas Add-On for MGX allows detailed analysis of individual cells of radially symmetric organs such as roots [7]. This guide briefly introduces the additional steps that are required to run a basic analysis using 3D Cell Atlas and to quantify the cell geometry beyond volume and wall area. A detailed guide for 3D Cell Atlas that is beyond the scope of this protocol can be found in Stamm et al. [8]. 1. In this example, we only want to analyze the epidermal cells of the root. Thus, the first step will be to create a segmented mesh of epidermal cells.

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Fig. 9 Segmentation and cell mesh analysis of the root epidermis. (a) A blurred root with broadly deleted columella during the segmentation process. (b) Rectangular selection of the surface mesh. (c) A section through the root is made by using a clipping plane. The mesh fragments in the inside that do not belong to the actual surface have to be removed. (d) The segmented mesh of epidermal cells inside the surface mesh. (e) A default Bezier grid with the main stack of a root. The grid has to be collapsed to a line and aligned to the root’s main axis. (f) Top view of the aligned Bezier line through the center of the vascular tissue of the cell mesh (visible as a dot in the center, see red arrow). (g) Same view as (f), but showing the original stack. (h) The bottommost cell has been selected in the epidermal cell mesh (highlighted in red, see yellow arrow)

2. Run the “Gaussian Blur” process (see step 3 in Subheading 3.3.2). 3. Broadly remove the columella region (Fig. 9a) by selecting the “Voxel Edit” tool in the upper toolbar and using Alt+Left mouse (click or drag). You can change the pixel edit size by going to “View” and under “Stack editing” dragging the “Pixel Edit Radius” bar. Be careful not to remove more than necessary (see Note 13)! 4. Segment the root using the “ITK Watershed” process (see step 4 in Subheading 3.3.2).

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5. For this protocol we are only interested in the epidermal cells and therefore will delete all other cells from the segmented root. Depending on the quality of the segmentation and the cell type of interest, this step can be omitted or adapted. You can delete whole cells by using the “Delete Label in Volume” tool . It is easiest to start in the median plane. You can view the median by using the clipping planes: “View > Clip1/2/3”. Do not forget to enable the Clips when you want to use them and disable them when you are done! You can view your actual scan (the main stack) by going to “Main > Stack 1” and checking “Main.” By comparing your stack to the segmentation, you can delete undesired cells. Be sure to save your work during the process, since you cannot undo an action! 6. Create the Mesh using the “Marching Cube” process (see step 6 in Subheading 3.3.2). 3.5.2 Root Cell Analysis Using 3D Cell Atlas

Having created a segmented mesh of root cells, we proceed to analyze the cells using the 3D Cell Atlas Add-On for MGX. Cell Atlas creates a root-specific coordinate system based on the center of the root and its surface. In order to be able to analyze and identify the positions of the cells, we first have to generate a surface mesh and to define the central axis of the root. 1. Open the original .TIFF stack in MGX. Change the X, Y and Z voxel size if necessary (check the dimensions in Fiji). This action should be performed regardless of any previous steps. 2. Blur the mesh a bit using the “Stack > Filter > Blur”. 3. In order to create the surface mesh of your stack, run the Process “Mesh.> Creation.> Marching Cubes Surface”. Leave the cube size at 5.0 as the surface mesh does not have to be very fine. A threshold of 2500 or lower is usually required. 4. In the “Main” tab, uncheck “Main” to conceal the main stack. This allows you to work only with the mesh. 5. Reset the view with “Anchor”

.

6. Next we need to delete the part of the mesh that does not represent the true organ surface. For this in the “Main” tab, check “Mesh.” Select the “Select Points in the Mesh” tool and use Alt+Left mouse and drag the mouse to create a rectangle at the top of the root (Fig. 9b). This will select the vertices of the rectangular selection. Press “Delete” to remove the selected vertices inside, which opens up the surface mesh. Be sure that you do not delete too much surface! If you do, your cell mesh might rise out of your surface mesh. 7. The inside of the root might contain some nonsurface mesh. In order to remove it, use the “Select Connected Area” tool and Alt+Left click on the surface mesh (that you want to keep).

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It will be selected in red. When it is selected, invert the selection using “Label and Selection/Invert Selection” which will select all other mesh fragments inside of the organ surface mesh. Now press “Delete” to remove them. 8. In order to reduce noise on the surface mesh it can be useful to smooth it. To do this use the Process “Mesh > Structure > Smooth Mesh” until you are satisfied with the result. 9. Now save the surface mesh via “Mesh > Mesh 1 > Save”. 10. Restart MGX and open your segmented cell mesh in “Mesh 1” and the surface mesh in “Mesh 2” (Fig. 9d). 11. In the “Main” tab, go to “Stack 2” and uncheck “Surface.” This hides the surface mesh. After this go back to “Stack 1,” which needs to be the active stack. 12. Next we have to accurately determine the center of the root. This will be done by placing the Bezier line through the center of the root. In order to this, it might be helpful to load your original .TIFF file (“Stack > Stack 1 > Main > Open”). Go to the “View” tab, check “Draw,” “Grid,” and “Bezier.” Press “Reset.” You will now see a Bezier grid in your root (Fig. 9e). 13. Collapse the Bezier grid to a single line with “Process > Mesh > Cell Atlas 3D > Tools > Collapse Bezier Points”. 14. Position the Bezier line precisely in the center of the root, that is, through the center of the vascular tissue and the root tip (Fig. 9f, g). Make sure that the line also goes through the center of the cell mesh “Main > Stack 1 > uncheck “Main” to see this. The line can be edited by selecting and moving its yellow control points. Select the points by using the “Select Points in Mesh” tool with the “Grid” checkbox checked in the “View” tab. Move selected points with Alt+Right mouse (see Note 14). 15. In “Main,” select “Stack 1” and make sure that “Mesh” is checked. Select the bottom most cell by using the “Select Connected Area” tool and Alt+Left click on the cell. It will be highlighted in red (Fig. 9h, yellow arrow). 16. After having created the central line and the surface mesh, the cells can be analyzed by Cell Atlas. Run the process “Mesh > Cell Atlas 3D > A – Analyze Cells 3D” with the default parameters. This process will calculate the cell center position and the organ centric coordinates as well as the cell sizes for each cell (see Note 15). Further processing can be done using the 3D Cell Atlas pipeline to annotate the different cell layers (for details see [7, 8]).

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17. For the purpose of this protocol, it is sufficient to quantify the cell sizes and their positions within the organ, which has been done using the “Analyze Cells 3D” process. In order to create heat maps of this data, use the process “Mesh > Cell Atlas 3D > Tools > Heat Map Cell Data” with the desired options. 18. To save and view the generated cell data, run process “Mesh > Cell Atlas 3D > Tools > Save Cell Data”. Input a file name (add .csv) before running the process. The file will be saved in the same folder as your meshes. The .csv file can be opened in excel. A single heat map can be saved using the process “Mesh > Heat Map > Heat Map Save”. An example of such an output file is visible in Table 4.

4

Notes 1. Paraformaldehyde needs gentle heating to dissolve. First dissolve it in PBS and let the solution cool down before adding any other reagents. Paraformaldehyde is very toxic: avoid breathing hot Paraformaldehyde vapors. 2. For roots: as a general indicator, a stack should be >80 μm in depth. If it is smaller, it is likely that the root is flattened by the coverslip to such extent that it is not usable for segmentation analysis. You can see the depth of an image in the Z-stack window under “Range.” 3. MGX can open two different images at a time (Stack 1 and Stack 2). Every stack can hold two versions of the same image: the “main” and the “work” store. Running any process that alters the image will create a result in the “work” store. 4. To visualize the inside of an organ or to slice through your sample, clipping planes can be used. They are accessible in the “View” tab. Clipping planes can be rotated and moved by selecting them in the “Control-Key Interaction” menu and using Ctrl+Left/Right mouse and dragging the mouse. Their thickness can be changed with the slider in the “Clip” tab. 5. If the image disappears after changing the voxel size by several orders of magnitude, use the “Anchor” button in the top toolbar to reset the view. 6. If the sample is highly oversegmented, increase the watershed threshold. If the sample is highly undersegmented, decrease the threshold. Your final segmentation is allowed to be oversegmented but not undersegmented because MorphoGraphX has tools to merge cell fragments but not to split segments of several cells.

Cell type

0

0

0

0

0

0

0

0

0

0

0

0

0

0

Cell label

2

37

54

61

66

105

136

165

174

176

205

221

264

340

0

0

0

0

0

0

0

0

0

0

0

0

0

0

Associated Cortical Cell

Table 4 Cell atlas output data Longitudinal length

0.705954 7.97003

0.207859 0.729247 6.68455

0.64487

0.390074 0.777354 10.8993

0.720723 0.719913 7.32756

0.309947 0.726058 7.49614

0.964118 0.411747 5.74734

0.684963 0.394749 10.9774

0.986338 0.366941 9.16689

0.273124 0.677378 14.2396

0.851043 0.564449 11.3136

0.152148 0.705512 7.69316

0.842824 0.636435 9.49705

0.919731 0.449439 8.68129

0.683572 0.733028 6.46967

Arclength Radius

15.5355

13.0751

12.954

11.4428

15.5267

6.18354

5.54654

6.893

14.8828

11.8615

12.7234

8.1678

6.22973

12.0398

Radial length

14.4011

10.2578

11.0557

9.72929

14.0827

7.81527

13.25

6.2539

13.4343

10.8708

17.4603

7.49975

9.48754

10.3059

Circumferential length

1239.01

900.801

1234.8

616.023

1391.01

205.969

593.787

269.393

1691.68

690.748

1292 67

428.841

360.303

637.64

Cell Volume

678.338

515.341

630.705

407.221

723.968

191.668

431.01

223.223

789.004

420.397

697.605

311.17

285.996

423.538

Cell Wall Area

80 Merijn Kerstens et al.

0

0

0

0

0

0

404

407

432

446

455

465

0

0

0

0

0

0

0 0.614192 8.50556

0.475548 0.717057 5.79442

0.165173 0.749551 14.964

0.609957 0.710944 7.7129

0.385271 0.713541 5.40315

0.748639 0.692288 7.66036

0.50794

0.142507 0.724209 5.94096

14.0018

14.4208

14.8887

17.7392

11.2885

14.0179

14.5015

11.6214

9.55463

10.5073

14.1211

10.8431

18.8247

16.8689

692.199

1581.09

952.344

1107.59

663.613

1445

1339.98

476.518

772.605

538.58

696.205

411.769

716.534

764.564

For each cell its label (column A, longitudinal (“Arclength,” D) and radial (“Radius,” E) position (in relative units) as well as cell lengths along its principal directions (in μm, columns F–H) and its volume and cell wall area (both in μm3). (I, J) are computed and exported to a .csv file. In this protocol we did not annotate the cell types; therefore columns B and C stay empty

0

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7. The segmentation assigns each cell a different label number. You can find out the cell label by using the “Pick Label” tool and Alt+Left click on a cell. The label then appears in the bottom left corner of the MGX interface. This can be very helpful to distinguish between cells with the same segmentation color! Also by checking the checkbox “Main >Stack 1” “16 Bit” (next to “Work”) the colors will be changed, which can help to distinguish different cells. 8. The cube size parameter specifies how fine the resulting mesh will be. Depending on the size of the cells and the quality of the segmentation, we suggest cube sizes between 1.0 and 5.0 μm Watershed (or an earlier saved stack) and to merge/delete cells. 9. The process “Stack > Segmentation > Join regions” allows you to merge cells even after the mesh is generated. Merge cells by clicking on them using the “Select Connected Area” tool and run the process with the desired cube size. This will merge the cells in the segmented stack and recompute the mesh for the selected cells only. 10. When working with the mesh, use the buttons on the left toolbar. The upper toolbar only shows the stack tools. 11. If you want to cross-check exported data with your actual cell mesh, it might be convenient to look up individual cells with, for example, aberrant volumes. For this use the process, “Mesh> Selection> Select label” and specifying the cell label number as parameter. The process will select the cell that will be highlighted in the mesh if the “Mesh” option in the “Main” tab is selected and the drop-down menu is on “Selected.” Additionally, all cell label numbers can be made visible when checking the “Map” option in the “Main “tab after creating a heat map. 12. Be careful changing the label of a mesh. If the same label is given to two different cells, the quantification of geometrical properties such as the volume might produce wrong results! 13. The pixel edit tool only works on the “Work” stack. The “Main” stack cannot be altered as it holds the original image. 14. You can also rotate and translate the Bezier line as a whole by checking the “Cut Surf” option in the “View” tab and dragging the mouse while pressing Ctrl+Left mouse (rotation) and Ctrl+Right mouse (moving). 15. The Cell Atlas coordinate system consists of three components: longitudinal (or arclength; along the Bezier line or the main axis of the organ, with values from 0 at the beginning of the Bezier line and 1 at the end); radial (or radius; from the Bezier line to surface, from 0 to 1) and circumferential (around the Bezier line, from 0 at the selected cell to 2π). Finally, for each cell its principal directions within this coordinate system are calculated and the cell size is measured along those directions.

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References 1. van den Berg C, Willemsen V, Hage W, Weisbeek P, Scheres B (1995) Cell fate in the Arabidopsis root meristem determined by directional signalling. Nature 378:62–65 2. de Reuille PB, Routier-Kierzkowska AL, Kierzkowski D, Bassel GW, Schu¨pbach T, Tauriello G, Bajpai N, Strauss S, Weber A, Kiss A, Burian A et al (2015) MorphoGraphX: a platform for quantifying morphogenesis in 4D. Elife 4:5864 3. Truernit E, Bauby H, Dubreucq B, Grandjean O, Runions J, Barthe´le´my J, Palauqui JC (2008) High-resolution whole-mount imaging of three-dimensional tissue organization and gene expression enables the study of phloem development and structure in Arabidopsis. Plant Cell 20:1494–1503 4. Smith ZR, Long JA (2010) Control of Arabidopsis apical-basal embryo polarity by antagonistic transcription factors. Nature 464:423–426 5. Musielak TJ, Schenkel L, Kolb M, Henschen A, Bayer M (2015) A simple and versatile cell wall

staining protocol to study plant reproduction. Plant Reprod 28:161–169 6. Schindelin J, Arganda-Carreras I, Frise E, Kaynig V, Longair M, Pietzsch T, Preibisch S, Rueden C, Saalfeld S, Schmid B, Tinevez JY, White DJ, Hartenstein V, Eliceiri K, Tomancak P, Cardona A (2012) Fiji: an opensource platform for biological-image analysis. Nat Methods 9:676 7. Montenegro-Johnson TD, Stamm P, Strauss S, Topham AT, Tsagris M, Wood AT, Smith RS, Bassel GW (2015) Digital single-cell analysis of plant organ development using 3D CellAtlas. Plant Cell 27:1018–1033 8. Stamm P, Strauss S, Montenegro-Johnson TD, Smith R, Bassel GW (2017) In silico methods for cell annotation, quantification of gene expression, and cell geometry at single-cell resolution using 3D Cell atlas. Methods Mol Biol 1497:99–123

Part III Expression Analysis and Transcriptomics

Chapter 7 Small RNA In Situ Hybridizations on Sections of Arabidopsis Embryos Katalin Pa´ldi, Magdalena Mosiolek, and Michael D. Nodine Abstract Small RNAs mediate posttranscriptional gene silencing in plants and animals. This often occurs in specific cell or tissue types and can be necessary for their differentiation. Determining small RNA (sRNA) localization patterns at cellular resolution can therefore provide information on the corresponding gene regulatory processes they are involved in. Recent improvements with in situ hybridization methods have allowed them to be applied to sRNAs. Here we describe an in situ hybridization protocol to detect sRNAs from sections of early staged Arabidopsis thaliana (Arabidopsis) embryos. Key words In situ hybridization, Small RNA, microRNA, Plant development, Embryo, Arabidopsis

1

Introduction RNA in situ hybridization is a technique that utilizes antisense oligonucleotide probes to detect complementary RNAs in a tissue of interest. This enables the characterization of RNA localization patterns and thus yields insights into their functions. The incorporation of locked nucleic acids (LNAs) in oligonucleotide probes increases hybridization probe affinity and thermal stability of the probe–target RNA duplex. Consequentially, the length of the probe required for stable target RNA duplex formation can be reduced facilitating the detection of small RNAs (sRNAs) from various species and tissue types [1–8]. An additional fixation step uses 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide (EDC) to immobilize the 50 monophosphates of sRNAs to the specimen’s protein matrix. This enhanced the sensitivity and robustness of sRNA in situ hybridization methods [9, 10].

The original version of this chapter was revised. The correction to this chapter is available at https://doi.org/ 10.1007/978-1-0716-0342-0_21 Martin Bayer (ed.), Plant Embryogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2122, https://doi.org/10.1007/978-1-0716-0342-0_7, © The Author(s) 2020, corrected publication 2021

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Fig. 1 Example images of small RNA in situ hybridizations on sectioned Arabidopsis embryos. Small RNA in situ hybridizations were performed with LNA-containing and dual DIG-labeled probes antisense to either the plantspecific miR156 (a) or animal-specific miR124 (b; negative control). Scale bars represent 20 μm. Oligonucleotide probe sequences and modifications were designed as described in [10]

Here, we present a detailed protocol for sRNA in situ hybridization on sections of Arabidopsis embryos. This protocol is based on previous mRNA and sRNA in situ protocols [3, 10–13]. Paraffin-embedded siliques are sectioned allowing access to young embryos, and hybridization is carried out with LNA-containing probes that are dual end-labeled with digoxigenin (DIG). Once probes are designed and prepared, the experiment takes 7 days to complete: tissue fixation and dehydration (days 1–3), clearing, embedding and sectioning (days 3–5), proteinase K digestion, EDC fixation and probe hybridization (days 5–6), washing and antibody reaction (day 6), and colorimetric reaction and mounting (days 6–7). Although this protocol was optimized for sectioned Arabidopsis embryos, it can be adapted to other tissue types with modifications as noted. Using this method, we were able to visualize the expression domain of miR156 from early Arabidopsis embryos (Fig. 1).

2

Materials

2.1 Reagents and Solutions

Solutions 1–12 must be made from RNase-free components. 1. DEPC-treated water: in the fume hood, add DEPC to a final concentration of 0.1% to deionized water, mix on a magnetic

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stirrer overnight, and then deactivate DEPC by autoclaving the next morning (see Note 1). 2. Formaldehyde-acetic acid-alcohol fixative (FAA; 50 ml): in the fume hood, add 17.5 ml DEPC-treated water, 5 ml 37% formaldehyde, 2.5 ml acetic acid and 25 ml 100% ethanol to 50 ml Falcon tube, mix and place on ice. 3. Eosin Y (0.1%): Dissolve 50 mg Eosin Y in 50 ml of 95% ethanol. 4. TE (10): 100 mM Tris, 10 mM EDTA, pH 7.5 (see Note 2). 5. Proteinase K buffer (5): 500 mM Tris, 250 mM EDTA in Milli-Q water, pH 8. 6. Proteinase K (10 mg/ml): Dissolve 100 mg in 100 ml 1 Proteinase K buffer, make single-use aliquots and store at 20  C. 7. Methylimidazole-NaCl: 0.13 M methylimidazole, 300 mM NaCl, pH 8. 8. Phosphate-buffered saline (PBS; 10): 1.37 M NaCl, 27 mM KCl, 100 mM Na2HPO4, 18 mM KH2PO4, pH 7.4. 9. EDC solution: 0.16 M N-(3-dimethylaminopropyl)-N0 -ethylcarbodiimide hydrochloride in Methylimidazole-NaCl. 10. Glycine (10): 20 mg/ml glycine, 0.15 M NaCl, 3 mM NaH2PO4, 7 mM Na2HPO4, pH 7. 11. Hybridization Salts (10): 3 M NaCl, 0.1 M Tris–HCl, 0.1 M Na2HPO4, 0.05 M EDTA, disodium salt dihydrate, pH 6.8. Make single-use aliquots and store at 20  C. 12. Dextran sulfate and Denhardt’s solution. Make single-use aliquots and store at 4  C. 13. tRNA: 50 mg/ml tRNA in DEPC-treated water, make singleuse aliquots and store at 20  C (see Note 3). 14. SSC (20): 3 M NaCl, 0.3 M trisodium citrate. 15. Maleic acid, sodium chloride buffer (MN buffer; 10): 1 M maleic acid, 1.5 M NaCl, pH 7.5 (see Note 4). 16. Blocking reagent solution (10): Dissolve 10 g of blocking reagent per liter 1 MN buffer and store at 4  C. 17. Tris-buffered saline (TBS; 10): 1 M Tris, 1.5 M NaCl, 5 mM MgCl2, pH 9.0. 18. BSA solution: Dissolve 6 g BSA with 1.6 ml Triton X-100, 60 ml 10 TBS and 540 ml Milli-Q water. 19. TNM5 buffer (1): 100 mM Tris, 100 mM NaCl, 5 mM MgCl2, pH 9.0. 20. TNP buffer (1): 100 mM Tris, 100 mM NaCl, 5 mM MgCl2, 10% PVA, pH 9.0.

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21. RNaseZap (ThermoFisher) or similar RNase decontamination solution. 22. HistoClear II or xylenes. 23. Paraplast Plus (required if manually embedding specimen). 24. EDC (N-(3-dimethylaminopropyl)-N0 -ethylcarbodiimide hydrochloride. 25. Ultrapure (GC 99.5%) and standard formamide. Make single-use aliquots of ultrapure formamide and store at 20  C. 26. DIG Nucleic Acid Detection Kit including Blocking reagent, NBT/BCIP stock solution and Anti-DIG-AP conjugate antibody (Roche). 27. Blocking reagent. 28. Anti-DIG-AP conjugate antibody. 29. NBT/BCIP Stock Solution (NBT/BCIP Stock Solution). Make single-use aliquots and store at 4  C. 30. Aqua Poly/Mount (Polysciences). 2.2

Equipment

1. 50 ml conical tubes (e.g., Falcon). 2. 20 ml glass scintillation vials. 3. Vacuum pump or access. 4. Automatic tissue embedding system (e.g., LOGOS Microwave Hybrid Tissue Processor from Milestone Medical) (optional). 5. Slide warming table. 6. Aluminum weighing dishes. 7. Insect pins. 8. Microtome. 9. Paintbrushes or cotton-tipped applicators. 10. Razor blades. 11. Hot plate. 12. Water bath. 13. Superfrost™ Ultra Plus Adhesion Slides. 14. Temperature-controlled incubators. 15. Staining dishes (~20–25) and holders (~5) (e.g., 10-slide dishes from Wheaton). 16. HybriSlip™HybridizationCovers(24mm60mm0.25mm; ThermoFisher). 17. Tupperware or similar plastic boxes with airtight seals (to use as a humidified chamber during hybridization). 18. Benchtop shaker. 19. Shallow plastic trays (e.g., 20 cm  30 cm).

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20. Slide mailers.

3 3.1

Methods Probe Design

3.2 Tissue Fixation and Dehydration (Days 1–3)

When designing probes, first place LNA modifications in central positions of the probe and then progressively add LNAs in evenly spaced positions along nonterminal positions of the probe until the desired RNA melting temperature between 80 and 90  C is reached (see Note 5). 1. Remove siliques from plant and gently slice both sides along the replum with a scalpel or needle (see Note 6). Place 15–20 siliques in 10 ml ice-cold FAA within a glass scintillation vial. Then vacuum infiltrate by pulling and releasing the vacuum as slowly as possible. Repeat it a few times until all siliques sink to the bottom of the vial. Gently tap desiccator occasionally to release air bubbles. Place vial at 4  C for 10–12 h (see Note 7). 2. Remove FAA from siliques and wash with 50% ethanol. 3. Add fresh 50% ethanol and incubate for 30 min at room temperature. Repeat with 65% ethanol, 80% ethanol and 95% ethanol (see Note 8). 4. Remove 95% ethanol and replace with 0.1% Eosin Y. Incubate overnight at 4  C (see Note 9). 5. Remove 0.1% Eosin Y and replace with 100% ethanol. Incubate for 1 h at room temperature. 6. Exchange 100% ethanol with fresh 100% ethanol and incubate for 30 min. Repeat 1 (see Note 10).

3.3 Clearing, Sectioning and Embedding (Days 3–5)

1. Samples can be cleared and embedded using an automatic tissue processor (e.g., we use LOGOS Microwave Hybrid Tissue Processor from Milestone Medical using the standard overnight cycle) (see Note 11). 2. Remove cassettes containing siliques from the tissue processor and place on a hot plate. Alternatively if embedding was done by hand, prepare paraffin blocks by pouring warm embedded material into an aluminum weighing dish on the hot end of slide warming table. Use insect pins to orient siliques so that they are in the same orientation. Carefully move aluminum dish to cooler part of the hot plate and rearrange as necessary. Let blocks harden for at least 1 h and store at 4  C until needed. 3. Using a standard microtome, cut paraffin-embedded siliques into 8–10 μm sections (see Note 12). Cut ribbons into approximately 2.5 cm strips with razor blades and carefully float strips in water bath filled with 42  C fresh deionized water for approximately 1 min. Then mount each ribbon on a glass slide by placing the slide underneath the section and carefully

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lifting up to capture the section on the slide (see Note 13). Place vertically for a couple of minutes to remove excess water. 4. Immediately place mounted slides on a hot plate (or in an incubator) set to 45–50  C overnight to bake the sections onto the slides. 3.4 Proteinase K Digestion, EDC Fixation and Probe Hybridization (Days 5–6)

3.4.1 Dewaxing

Before starting the experiment bake all glassware (i.e., staining dishes, slides holders) at 180  C. Be careful not to heat up or cool down too quickly as this will crack the dishes. Clean all plastic tools with RNaseZAP and rinse with fresh water (forceps, brushes, plastic boxes, etc.). Clean bench with RNaseZAP and always wear gloves while handling samples. Before the probe hybridization step only use DEPC-treated water and DEPC-treated PBS or solutions made with them (see Note 14). 1. Equilibrate slides to room temperature (see Note 15). 2. Put slides in a staining dish slide holder and fill with clearing reagent (i.e., HistoClear or xylenes (see Note 16)). Incubate for 10 min at room temperature. Dip slides up and down a few times during the incubation. Repeat 1 with fresh clearing reagent.

3.4.2 Hydration

1. Transfer slides from clearing reagent to a staining dish containing 100% ethanol. Dip slides up and down 15. Incubate for 5 min to remove clearing reagent. Repeat 1. 2. Process the slides through 95%, 85%, 70%, 50%, 30% and 15% ethanol. For each ethanol series, dip slides up and down 15 and incubate for 2 min (see Notes 17 and 18). 3. Incubate in 1 TE for 2 min. Repeat 1 with fresh 1 TE.

3.4.3 Proteinase K Digestion

1. Incubate slides for 10 min in 1 proteinase K buffer (without proteinase K) at room temperature. Dip slides up and down a few times to equilibrate the sections (see Note 19). 2. Add proteinase K to the prewarmed proteinase K buffer to a final concentration of 1 μg/ml. Transfer slides to staining dish containing proteinase K buffer (with proteinase K), and incubate at 37  C for 30 min (see Note 20). 3. Transfer slides to a staining dish containing 1 glycine. Dip up and down a few times to rinse off the proteinase K and incubate for 2 min at room temperature.

3.4.4 EDC Fixation

1. Transfer slides to a staining dish containing 1 PBS, dip up and down a few times to rinse off glycine and incubate for 2 min. Repeat 1.

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2. Transfer slides to a staining dish containing freshly prepared methylimidazole-NaCl and incubate for 10 min at room temperature. Repeat 1. 3. Transfer slides to EDC solution and incubate for 2 h at 60  C. 4. Transfer slides to 1 PBS, dip up and down a few times and incubate for 5 min. Repeat 2. 3.4.5 Dehydration

1. Transfer slides to a staining dish containing 15% ethanol, dip up and down a few times, and incubate at room temperature for 2 min. Repeat for 30%, 50%, 70%, 85%, 95%, and 100% ethanol. Dip slides up and down 15 and incubate for 2 min in each ethanol solution (see Note 21). 2. Let slides air-dry on bench with sections facing up for 2 h.

3.4.6 Probe Hybridization

Preheat an incubator to the preferred hybridization temperature. Prepare a humidified box for slides. We use airtight sealed plastic boxes filled with paper towels soaked in 50% formamide (standard grade) at the bottom, layered with glass pipettes on the top (Fig. 2). The pipettes prevent slides from coming into contact with the paper towels (see Note 22). 1. Prepare Hybridization Mix. Pipet the reagents in the following order to prepare 2 ml: 150 μl nuclease-free water, 250 ml 10 hybridization salts, 1 ml ultrapure and ice-cold formamide, 0.5 ml 50% dextran sulfate, 50 μl Denhardt’s solution, and 50 μl 50 mg/ml tRNA. Mix by pipetting up and down slowly to avoid air bubbles (see Notes 23 and 24).

Fig. 2 Assembly of probe hybridization chamber. In an airtight sealed plastic box (e.g., Tupperware), layer paper towels soaked in 50% formamide on the bottom to create a humidified chamber and place glass pipettes (or Parafilm) on top of paper towels to create a barrier between the formamide and slides. Cover chamber with an airtight lid and prewarm in an incubator to the desired hybridization temperature. Remove from incubator, add slides with specimen/probe, cover chamber, and place back in incubator for 10–12 h

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2. Prepare LNA probes: Add 160 μl of hybridization solution to separate Eppendorf tubes (one for each slide to be hybridized). Add appropriate amount of each probe to individual 200 μl PCR strip tubes and bring up to 40 μl with 50% formamide (see Note 25). Incubate at 80  C for 2 min and then 4  C in thermal cycler. Add 40 μl of probe/formamide mix to Eppendorf tubes containing hybridization solution and slowly mix by pipetting up and down 15. 3. Apply 160 μl of probe/hybridization mixture evenly to the slide by carefully pipetting along the section (see Note 26). 4. Carefully cover slides with HybriSlip covers without making bubbles. Use one pair of forceps to hold the HybriSlip in place while slowly lowering it onto the slide with another pair of forceps. 5. After applying the hybridization solution, immediately place the slides in a prewarmed box humidified with 50% formamide (standard grade). Incubate overnight at the optimal hybridization temperature (see Note 27). 3.5 Washing and Antibody Reaction (Day 6)

Preheat two 500 ml beakers containing 200 ml 0.2 SSC solution to 55  C in an incubator.

3.5.1 Coverslip Removal and High Stringency Washes

1. Fill two 500 ml beakers with 300 ml 2 SSC. Individually remove each slide from the hybridization box with forceps, and dip up and down in the first beaker of 2 SSC until the coverslip falls off (see Note 28). 2. Dip each slide up and down a few times in the second beaker to rinse off excess hybridization buffer, and place in a staining dish containing 2 SSC. Repeat this and the above step for each slide. 3. After each slide has been processed, transfer them to a second staining dish with fresh 2 SSC and incubate at room temperature for 5 min. 4. Transfer slides into the first 500 ml beaker containing prewarmed 0.2 SSC and incubate for 1 h at 55  C. Dip slide rack up and down a few times during the incubation. Alternatively, use a water bath with a shaking platform. 5. Transfer the slides to the second beaker with 0.2 SSC and incubate for another 1 h at 55  C dipping up and down a few times.

3.5.2 Blocking and Antibody Reaction

1. Remove slide holder from beaker and place it into a staining dish containing 1 MN buffer. Dip slides up and down 15 and then incubate at room temperature for 10 min.

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2. Transfer each individual slide into a shallow plastic tray filled with 1% blocking reagent solution. Incubate at room temperature for 30 min with gentle rotation on a rotating platform. Repeat 1 (see Note 29). 3. Transfer each individual slide into a shallow plastic tray containing 1% BSA solution. Incubate at room temperature for 30 min with gentle rotation. Repeat 1. 4. Dilute required amount of antibody in 1% BSA solution (see Note 30). 5. Fill slide mailers with 16 ml of antibody/BSA solution. Transfer each individual slide from 1% BSA solution to a slide mailer (5 slides/mailer). 6. Incubate at room temperature for 2 h with slow rotation. 7. Transfer slides to a plastic tray containing 1% BSA solution and incubate at room temperature for 20 min with slow rotation. Repeat 3 (see Note 31). 3.6 Colorimetric Reaction and Mounting (Days 6–7) 3.6.1 Colorimetric Reaction

1. Dip slides up and down 15 in TNM5 buffer and incubate for 2 min. Repeat 2. 2. In 50 ml conical tube, dilute 500 μl NBT/BCIP Stock Solution in 50 ml TNP buffer. Add 16 ml of this solution to each slide mailer (5 slides/mailer) and carefully place slides into mailers making sure that the sections are completely submerged (see Note 32). 3. Incubate at room temperature in complete darkness with no movement (see Note 33).

3.6.2 Mounting

1. When the color reaction is complete, wash the slides in 1 TE buffer for 5 min. Repeat 2. 2. Place slides on a clean sheet of aluminum foil with sections facing up. Apply 2–3 drops of Aqua-Poly/Mount and cover with a glass coverslip. Gently squeeze out the air bubbles and excess mounting medium by pressing down gently on the coverslip with a folded Kimwipes. 3. Let slides dry overnight at room temperature to harden the mounting medium. 4. Clean slides and examine under the microscope.

4

Notes 1. DEPC should not be used to treat solutions with high concentrations of Tris such as TE and proteinase K buffer.

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2. When adjusting pH for RNase-free solutions clean the glass electrode with RNaseZap and rinse with DEPC-treated water. 3. All solutions up to this point must be made from RNase-free components. 4. Use NaOH pellets during pH adjustment. 5. See https://www.qiagen.com for details and useful tools for calculating RNA melting temperatures and selfcomplementarities, but we recommend the following general principles during LNA probe design. LNA bases have significant differences in binding strengths (C > T > G  A). Try to design probes with low self-complementarity, 30–60% GC content, avoid 4 sequential LNAs, avoid 3 sequential LNA-modified G’s or C’s and do not place LNA-modified bases at oligo termini. 6. Be careful not to open siliques up too much in order to not lose ovules. We generate a perforated cut to keep valves attached but allow them to be subsequently removed easily. 7. Collection and fixation of siliques must be carried out on ice. 8. If siliques are floating in ethanol wash series (Subheading 3.2, steps 2 and 3), then vacuum infiltrate (described in Subheading 3.2, step 1) until siliques sink to the bottom of scintillation vials. 9. Eosin Y allows better visualization of tissues during embedding and sectioning. 10. At this point siliques can be stored for up to a few weeks at 4  C in 70% ethanol. First, wash siliques with 85% and 70% ethanol for 5 min each. 11. Paraffin embedding can also be done by hand in case an automatic embedding machine is not available. First, tissues need to be cleared with either xylenes or HistoClear as the clearing reagent. Remove 100% ethanol from previous step and replace with 25% clearing reagent/75% ethanol in glass scintillation vials. Incubate for 30 min and repeat with 50% clearing reagent/50% ethanol, 75% clearing reagent/25% ethanol and 100% clearing reagent. Repeat step with 100% clearing reagent. Place a beaker containing paraplast chips in an incubator and let melt for >6 h. Add 20 chips (approximately 2 g) to glass scintillation vials, place in incubator set to 42  C and swirl occasionally until paraplast is melted. Repeat this until vials are full (4–5) and increase temperature as needed to fully melt paraplast. Remove clearing reagent from vials containing fixed/cleared tissues and add melted paraplast. Mix by swirling vials and incubate at 60  C for 4 h. Exchange melted paraplast each morning and night for 2 days (4 total). 12. Place paraffin blocks on ice for 30 min before sectioning.

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13. Use paintbrushes or cotton-tipped applicators to handle ribbons and orient sections on slides. 14. Store solutions and buffers for in situ hybridizations on a designated RNase-free shelf/bench area separate from other lab chemicals. 15. Paraffin blocks can be stored for weeks at 4  C but after sectioning try to process the slides as soon as possible. 16. Originally xylenes were used to clear tissues and remove paraffin from tissue sections. However xylenes are toxic and HistoClear is a suitable replacement. 17. If the sections are still stained with eosin Y after the 85% ethanol wash, then let sections incubate an extra 5 min in 85% ethanol until the dye is not visible. 18. Keep the ethanol series (except 100%) in separate staining dishes to use again during the dehydration step. 19. Prewarm staining dish containing proteinase K buffer to 37  C before this step. 20. Proteinase K is used to partially digest the tissue to allow better probe penetration. However, over digestion reduces specimen integrity. Therefore, the ideal duration of proteinase K incubation needs to be carefully calibrated for each tissue type. These conditions were optimized for digesting sectioned embryos embedded within siliques. 21. Use the alcohol solutions from the hydration steps, except for the 100% ethanol because it is contaminated with clearing reagent. 22. Alternatively, a layer of Parafilm can be used to prevent direct contact between slides and paper towels soaked with formamide. 23. Dextran sulfate is viscous and very difficult to pipet. It helps to prewarm aliquots to room temperature before pipetting. 24. Hybridization solution should be made fresh for each set of hybridizations. 25. The optimal probe concentration should be determined for each probe individually. To reduce background, use the lowest possible probe concentration. For a new probe try different concentrations ranging from 5 nM to 100 nM. In our experience, 20 nM final concentration typically gives optimal results for early embryos. Prepare 1 mM working solutions from LNA probe stock (100 mM) and store at 20  C. 26. The amount of hybridization solution depends on the number of sections on the slide, and on the density and thickness of the sections. Based on our experience, 160 μl/slide works well.

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27. The optimal hybridization temperature needs to be experimentally determined. In our experience, 65  C works well for probes designed according to Note 5. 28. Do not remove coverslips manually with forceps and be careful not to damage the sections during this step. 29. Be careful not to rotate too fast. This can cause the slides to slide over each other and damage the specimen. 30. The optimal antibody dilution may change with different probes; however we typically use a 1:1500 dilution. 31. Add slides to mailers slowly and make sure all slides are covered in the color reagent evenly. Do not use slide mailers that were used for the antibody binding step. 32. Because TNP buffer is viscous, pour into conical tube and then pipet in the NBT/BCIP solution. Close screw cap and shake vigorously to mix. Cover with aluminum foil and protect from light. 33. For a typical probe, monitor the colorimetric reaction after 15 h. If you expect a very strong signal, begin monitoring after only a few hours. Choose a test slide and rinse with TNM5. Wipe off the back of the slide and observe using an old microscope. Put the test slide back into the color solution if the signal is weak.

Acknowledgments This work was supported by funding from the European Research Council under the European Union’s Horizon 2020 research and innovation program (Grant 637888 to M.D.N.). References 1. Carlsbecker A, Lee J-Y, Roberts CJ et al (2010) Cell signalling by microRNA165/6 directs gene dose-dependent root cell fate. Nature 465:316–321. https://doi.org/10.1038/ nature08977 2. Chitwood DH, Nogueira FTS, Howell MD et al (2009) Pattern formation via small RNA mobility. Genes Dev 23:549–554. https://doi. org/10.1101/gad.1770009 3. Nodine MD, Bartel DP (2010) MicroRNAs prevent precocious gene expression and enable pattern formation during plant embryogenesis. Genes Dev 24:2678–2692. https://doi.org/ 10.1101/gad.1986710 4. Valoczi A, Varallyay E, Kauppinen S et al (2006) Spatio-temporal accumulation of

microRNAs is highly coordinated in developing plant tissues. Plant J 47:140–151 5. Kloosterman WP, Wienholds E, de Bruijn E et al (2006) In situ detection of miRNAs in animal embryos using LNA-modified oligonucleotide probes. Nat Methods 3:27–29. https://doi.org/10.1038/nmeth843 6. Obernosterer G, Martinez J, Alenius M (2007) Locked nucleic acid-based in situ detection of microRNAs in mouse tissue sections. Nat Protoc 2:1508–1514. https://doi.org/10.1038/ nprot.2007.153 7. Darnell DK, Stanislaw S, Kaur S, Antin PB (2010) Whole mount in situ hybridization detection of mRNAs using short LNA containing DNA oligonucleotide probes. RNA

Small RNA In Situs in Plant Embryos 16:632–637. https://doi.org/10.1261/rna. 1775610 8. Javelle M, Timmermans MCP (2012) In situ localization of small RNAs in plants by using LNA probes. Nat Protoc 7:533–541. https:// doi.org/10.1038/nprot.2012.006 9. Pena JTG, Sohn-Lee C, Rouhanifard SH et al (2009) miRNA in situ hybridization in formaldehyde and EDC-fixed tissues. Nat Methods 6:139–141. https://doi.org/10.1038/nmeth. 1294 10. Ghosh Dastidar M, Mosiolek M, Bleckmann A et al (2016) Sensitive whole mount in situ localization of small RNAs in plants. Plant J

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88:694–702. https://doi.org/10.1111/tpj. 13270 11. Drews GN, Bowman JL, Meyerowitz EM (1991) Negative regulation of the Arabidopsis homeotic gene AGAMOUS by the APETALA2 product. Cell 65:991–1002 12. Vielle-Calzada JP, Thomas J, Spillane C et al (1999) Maintenance of genomic imprinting at the Arabidopsis medea locus requires zygotic DDM1 activity. Genes Dev 13:2971–2982 13. Long JA, Moan EI, Medford JI, Barton MK (1996) A member of the KNOTTED class of homeodomain proteins encoded by the STM gene of Arabidopsis. Nature 379:66–69. https://doi.org/10.1038/379066a0

Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made. The images or other third party material in this chapter are included in the chapter’s Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter’s Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

Chapter 8 Manual Isolation of Living Early Embryos from Tobacco Seeds Peng Zhao, Xuemei Zhou, Ce Shi, and Meng-xiang Sun Abstract Zygotic embryogenesis is one of key processes for fertile seed development and therefore has gained great attention for decades in the field of plant developmental biology. However, this process is deeply embedded in the maternal tissues. The inaccessibility of tiny early embryos has greatly hindered the study of early embryogenesis, especially limits direct observation and accurate omics investigations. In order to investigate the molecular mechanism regulating embryo development with modern technologies, it is necessary to develop a reliable method to isolate living embryos at different stages. For this purpose, plant scientists have been trying to develop different methods for isolating zygotes and early embryos in different plants such as maize, wheat, rice, and tobacco during past decades. Nicotiana tabacum has long been considered as an ideal model eudicot for the study of embryogenesis, which displays a traceable and predictable cell division pattern, spanning from the first zygotic division to the mature embryo formation. Here, we provide a detailed protocol for isolating living embryos from zygote to cotyledon embryo. Isolated living zygotes and early embryos could be used for several important studies such as cell type-specific transcriptome construction and clear GFP observation. Key words Tobacco, Embryo sac, Zygote, Early embryos, Microdissection

1

Introduction Zygotic embryogenesis is the beginning of a new sporophytic generation, which involves a series of important development events, such as parental genome fusion and interaction, zygotic genome activation (ZGA), the establishment of zygotic polarity and initiation of asymmetric zygote division and so on [1]. Therefore, the embryos are useful materials for researchers in the field of plant developmental biology. However, the inaccessibility of zygotes and early embryos, which are deeply embedded in endosperm and seed coats, has seriously hindered the researches on the molecular mechanism underlying embryogenesis, for example, the timing of ZGA or the characteristics of stage-specific transcriptome.

Martin Bayer (ed.), Plant Embryogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2122, https://doi.org/10.1007/978-1-0716-0342-0_8, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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Although it is technically difficult to isolate tiny embryos, the methods for manual isolation of early zygote or 1-cell embryo have been established in a few plants such as maize [2–4], rice [5–7], and wheat [8, 9]. Relevant techniques are still improving to be more efficient and more reliable concerning possible RNA degradation and contamination of other tissues. During past two decades, we have respectively developed a series of methods to isolate and collect living sperm cells, egg cells, zygotes, and embryos of tobacco for in vitro fertilization, zygote culture and cell typespecific cDNA library construction [10–13]. Here, we described a detailed method for isolation of living embryos from tiny zygote to mature cotyledon embryo by microdissection. These isolated embryos could be used for direct GFP observation and transcriptome analysis. Combined with the single-cell sequencing technology for transcriptome and epigenetic analysis, it will be also applicable in the studies on the timing of the ZGA, transcriptome dynamic and epigenetic modification during the whole process of embryogenesis.

2

Materials

2.1

Plant Materials

2.2

Reagents

Nicotiana tabacum var. SR1 and pNtCP14::H2B-GFP transgenic plants [14] were grown in the greenhouse under long-day conditions (16 h light:8 h dark) at 25
2  C. 1. Mannitol. 2. Cellulase R-10 (Yakult Pharmaceutical Industry Co. Ltd., Japan). 3. Macerozyme R-10 (Yakult Pharmaceutical Industry Co. Ltd., Japan). 4. Mineral oil. 5. 2-(N-Morpholino)ethanesulfonic acid hydrate (MES). 6. Fluorescein diacetate (FDA). 7. Dynabeads® mRNA Technologies).

DIRECT™

Micro

Kit

(Life

8. SMART-Seq® v4 Ultra® Low Input RNA Kit for Sequencing (Clontech). 9. Agencourt AMPure purification kit (Beckman Coulter). 2.3

Equipment

1. Inverted microscope. 2. Stereoscopic microscope. 3. Confocal microscope. 4. Fine tweezer (Fig. 1a).

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Fig. 1 Tools used for isolating early embryos of tobacco: (a) a fine tweezer; (b) handmade capillary pipette sealed with latex tubing at one end; (c) handmade glass pestle

5. Handmade capillary pipette with latex tubing (Fig. 1b). 6. Small glass pestle (Fig. 1c). 7. Petri dishes for microscope: diameter 3.5 cm. 8. Glass microscope slide. 9. 0.22-μm filter. 2.4 Solutions for Cell Isolation

1. Washing solution: 13% mannitol, 0.058%MES (pH ¼ 5.8). Dissolve 13 g of mannitol and 0.058 g of MES in about 80 ml deionized water in a 100 ml bottle. Mix well and adjust the pH to 5.8 with 1 M NaOH. Make up to 100 ml with deionized water. Then, autoclave at 121  C for 20 min. 2. Enzyme solution I: 1% Cellulase R-10 and 0.8% Macerozyme R-10 dissolved in Washing solution. 3. Dissolve 0.5 g Cellulase R-10 and 0.4 g Macerozyme R-10 in the 50 ml Washing solution. Filter-sterilize the enzyme solution I with a 0.22-μm filter and divide the enzyme solution into 1 ml per 1.5 ml Eppendorf tube. Store the enzyme solution I at 20  C. 4. Enzyme solution II: 0.25% Cellulase R-10 and 0.2% Macerozyme R-10 dissolved in Washing solution.

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5. Dissolve 25 mg Cellulase R-10 and 20 mg Macerozyme R-10 in the 10 ml Washing solution. Filter-sterilize the enzyme solution II with a 0.22-μm filter and divide them into 200 μl per tube. Store the enzyme solution II at 20  C. 6. FDA storage solution: Dissolve 2 mg FDA in 1 ml acetone and store at 20  C. 7. FDA staining solution: Add 1 μl FDA storage solution to a 1.5 ml Eppendorf containing 999 μl washing solution, then mix them well.

3

Methods

3.1 Hand Emasculation and Pollination

1. Choose flower buds before anthesis (Fig. 2a) and carefully check the anther to confirm that it does not dehisce yet. Remove five anthers thoroughly using a fine tweezer (Fig. 2b) (see Note 1). 2. Pollinate emasculated mature pistil with opened anthers after 24 h (Fig. 2c). Label the flower and record the exact time of pollination.

3.2 Seed Collection and Embryo Isolation (Before 32-Cell Embryo Stage)

1. According to the time course of embryo development (Fig. 3), seeds at corresponding stages were collected for embryo isolation.

Fig. 2 Hand pollination and seed collection for enzyme treatment. (a, b). Choose flower buds before anthesis for emasculation; (c) pollinate emasculated mature pistil with an opened anther; (d, e) seed collection for enzyme treatment; (f) enzyme treatment of seeds in a 1.5-ml Eppendorf tube

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Fig. 3 Time course of tobacco embryo development. HAP, hours after pollination; DAP days after pollination. Bars, 10 μm

2. Collect the seeds quickly from the placenta into a 1.5-ml Eppendorf tube containing 1 ml enzyme solution (Fig. 2d, e) (see Note 2). 3. Place the 1.5-ml Eppendorf tube horizontally into the 25  C incubator for 30 min (Fig. 2f). Gently shake the Eppendorf tube every 10 min during the process of enzyme treatment (see Note 3). 4. Place the tube on the 1.5 ml centrifuge tube rack for 5 min to sink the seed to the bottom of the tube. 5. Carefully remove the solution and add 1 ml washing solution quickly (see Note 4). 6. Repeat steps 4 and 5 twice. 7. Mix well the washing solution containing the seeds, pipet 100 μl solution containing seeds with a 1 ml pipette tip onto the glass slide (Fig. 4a) (see Note 5). 8. Grind the seeds gently on the glass slide with a small glass pestle (Fig. 4b); check the seeds under the inverted microscope and screen the embryo sac as soon as possible (Fig. 4c) (see Notes 6–8). 9. Prepare the solution for short-term store. 10. Add 2 ml mineral oil onto the petri dish, and then add 20 μl washing solution on the bottom the mineral oil to form a drop of washing solution for embryo sac store. About 20 droplets could be made in the same petri dish (see Note 9). 11. Transfer isolated embryo sacs with a handmade micropipette into the droplet of washing solution covered by mineral oil (Fig. 4d).

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Fig. 4 Procedure for embryo isolation. (a) Pipet 100 μl solution containing seeds with a 1 ml pipette tip onto the glass slide. (b) Grind the seeds gently on the glass slide with a small glass pestle. (c) Seek for isolated embryo sac under the inverted microscope. (d) Transfer isolated embryo sac with a handmade micropipette. (e, f) Isolated embryo sac containing a zygote. (g) Isolated zygote from the embryo sac. (h) Embryos after 32-cell embryo stage could be directly released form the seeds by gentle grinding. Bars for (c) and (d), 20 μm; (e–h), 10 μm

12. Wash the isolated embryo sacs in 50 μl washing solution. Clean embryo sacs containing embryos could be observed under microscope after extensive washing (Fig. 4e, f).

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13. Prepare a droplet of enzyme solution II covered by mineral oil in the petri dish. 14. Transfer isolated embryo sacs into the enzyme solution II for the second enzyme treatment in a 25  C incubator for 10 min. 15. Transfer embryo sacs into the droplet of washing solution, and embryos could be released from the embryo sacs by gently pipetting up and down a few times with the handmade micropipette. 16. After extensive wash, isolated embryos could be used for subsequent analyses (Fig. 4g) (see Note 10). 3.3 Embryo Isolation (After 32-Cell Embryo Stage)

1. Seed collection, enzyme treatment and washing were performed according to the methods described above (Subheading 3.2, steps 1–7). 2. Pipet 200 μl washing solution containing seeds with a 1 ml pipette tip onto a glass slide (see Note 5). 3. Grind the seeds on the glass slide gently with a small glass pestle (see Notes 6–8). 4. Seek for embryos on the glass slide after grinding under inverted microscope (Fig. 4h). Embryos could be directly released from the seeds by gentle grinding as mentioned above. 5. Transfer isolated embryos with a handmade micropipette into the washing solution. 6. After washing, isolated embryos could be directly used for GFP observation and transcriptome analysis (Fig. 5a, b) (see Note 10).

3.4 Estimation of Cell Viability

1. Prepare FDA staining solution by adding 1 μl of FDA stock solution to 999 μl of washing solution (see Note 11). 2. Isolated embryos were transferred into a droplet of 50 μl FDA staining solution covered by mineral oil for 15 min at room temperature (see Note 11). 3. Wash the embryo with 50 μl washing solution before observation. 4. Living embryos with green fluorescence were observed under confocal microscope (Fig. 6).

3.5 mRNA Extraction and cDNA Amplification

1. Collect about 20 embryos in a 200 μl PCR tube for mRNA isolation using the Dynabeads® mRNA DIRECT™ Micro Purification Kit. Isolated embryos in lysis buffer could be used for direct mRNA isolation or storage in 80  C up to 1 month.

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Fig. 5 Isolated embryos could be used for GFP observation and transcriptome analysis. (a) GFP fluorescence of embryos isolated from pNtCP14::H2B-GFP transgenic plants. Bars, 10 μm. (b) Amplified cDNA of 1-Cell and 32-Cell embryo. (c) Relative expression levels of NtWOX2 and NtWOX9 in early embryos. The expression level of GAPDH in 1-cell embryo was set to 1000. Error bars represent mean
standard error (n ¼ 3)

2. cDNA synthesis and PCR amplification were carried out using the SMART-seq™ v4 Ultra™ Low Input RNA Kit for Sequencing (Clontech, USA). 3. PCR-amplified cDNA was purified using an Agencourt AMPure purification kit (Fig. 5b). 4. Purified cDNA could be used for gene expression analysis (Fig. 5c) or RNA-seq.

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Fig. 6 Cell viability of isolated embryos checked by FDA staining. (a) Zygote; (b) 1-cell embryo; (c) 2-cell embryo; (d) 8-cell embryo. Ac apical cell, Bc basal cell, Ep embryo proper, Sus suspensor. Bars, 10 μm

4

Notes 1. Avoid damaging the stigma during the process of emasculation and pollination. 2. To avoid browning of seeds, it is necessary to transfer seeds from the placenta into the enzyme solution as soon as possible. The brown seeds could not be used for embryo isolation in the later stages. 3. To maintain the vitality of isolated embryos, it is necessary to control the time of enzymatic treatment. 4. To avoid removing the seeds during the process of washing, it is necessary to take the solution away carefully from the top to bottom of the Eppendorf tube. 5. The tip of pipette should be cut off when used for aspirating solution containing seeds. Seeds could not be easily pipetted using a normal 1 ml pipette tip. It is better to prepare a glass pipette for this purpose. 6. The power to grind the seeds should be appropriate. If the force for grinding is not enough, the embryo sac and embryos

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may not be easily squeezed out from the seed. If the force for grinding is too strong, the embryo sacs and embryos can be crushed by the grinding. 7. Seek for embryo sacs from the glass slide and transfer them into oil-covered droplet of solution as soon as possible since the mannitol is easy to crystallize from the washing solution. 8. The seeds could be used for embryo sac isolation within 30 min. 9. To avoid precipitation of mannitol, the washing solution should be covered by mineral oil. 10. Wash the isolated embryo sacs or embryos with 50 μl washing solution (1:100 dilution), repeat washing step twice. It is necessary to perform extensive wash to remove seed coat tissues and endosperm contamination with the help of the pipettes. 11. Preparation of the FDA solution and staining of embryos with FDA should be performed in a dark environment.

Acknowledgments This work was supported by National Natural Science Fund of China (31970340, 31600244). References 1. Zhao P, Begcy K, Dresselhaus T, Sun MX (2017) Does early embryogenesis in Eudicots and monocots involve the same mechanism and molecular players? Plant Physiol 173 (1):130–142. https://doi.org/10.1104/pp. 16.01406 2. Chen J, Strieder N, Krohn NG, Cyprys P, Sprunck S, Engelmann JC, Dresselhaus T (2017) Zygotic genome activation occurs shortly after fertilization in maize. Plant Cell 29(9):2106–2125. https://doi.org/10.1105/ tpc.17.00099 3. Meyer S, Scholten S (2007) Equivalent parental contribution to early plant zygotic development. Curr Biol 17(19):1686–1691. https:// doi.org/10.1016/j.cub.2007.08.046 4. Leduc N, MatthysRochon E, Rougier M, Mogensen L, Holm P, Magnard JL, Dumas C (1996) Isolated maize zygotes mimic in vivo embryonic development and express microinjected genes when cultured in vitro. Dev Biol 177(1):190–203. https://doi.org/10.1006/ dbio.1996.0155 5. Abiko M, Maeda H, Tamura K, HaraNishimura I, Okamoto T (2013) Gene

expression profiles in rice gametes and zygotes: identification of gamete-enriched genes and up- or down-regulated genes in zygotes after fertilization. J Exp Bot 64(7):1927–1940. https://doi.org/10.1093/jxb/ert054 6. Zhang J, Dong WH, Galli A, Potrykus I (1999) Regeneration of fertile plants from isolated zygotes of rice (Oryza sativa). Plant Cell Rep 19(2):128–132. https://doi.org/10. 1007/s002990050722 7. Zhao J, Zhou C, Yang HY (2000) Isolation and in vitro culture of zygotes and central cells of Oryza sativa L. Plant Cell Rep 19 (3):321–326. https://doi.org/10.1007/ s002990050020 8. Kumlehn J, Lorz H, Kranz E (1998) Differentiation of isolated wheat zygotes into embryos and normal plants. Planta 205(3):327–333. https://doi.org/10.1007/s004250050327 9. Sprunck S, Baumann U, Edwards K, Langridge P, Dresselhaus T (2005) The transcript composition of egg cells changes significantly following fertilization in wheat (Triticum aestivum L.). Plant J 41

Isolation of Tobacco Embryos (5):660–672. https://doi.org/10.1111/j. 1365-313X.2005.02332.x 10. He YC, He YQ, Qu LH, Sun MX, Yang HY (2007) Tobacco zygotic embryogenesis in vitro: the original cell wall of the zygote is essential for maintenance of cell polarity, the apical-basal axis and typical suspensor formation. Plant J 49(3):515–527. https://doi.org/ 10.1111/j.1365-313X.2006.02970.x 11. Xin HP, Peng XB, Ning J, Yan TT, Ma LG, Sun MX (2011) Expressed sequence-tag analysis of tobacco sperm cells reveals a unique transcriptional profile and selective persistence of paternal transcripts after fertilization. Sex Plant Reprod 24(1):37–46. https://doi.org/10. 1007/s00497-010-0151-y 12. Zhao J, Xin H, Qu L, Ning J, Peng X, Yan T, Ma L, Li S, Sun MX (2011) Dynamic changes of transcript profiles after fertilization are

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associated with de novo transcription and maternal elimination in tobacco zygote, and mark the onset of the maternal-to-zygotic transition. Plant J 65(1):131–145. https://doi. org/10.1111/j.1365-313X.2010.04403.x 13. Zhou XM, Guo YY, Zhao P, Sun MX (2018) Comparative analysis of WUSCHEL-related Homeobox genes revealed their parent-of-origin and cell type-specific expression pattern during early embryogenesis in tobacco. Front Plant Sci 9:311. https://doi.org/10.3389/ fpls.2018.00311 14. Zhao P, Zhou XM, Zhang LY, Wang W, Ma LG, Yang LB, Peng XB, Bozhkov PV, Sun MX (2013) A bipartite molecular module controls cell death activation in the basal cell lineage of plant embryos. PLoS Biol 11(9):e1001655. https://doi.org/10.1371/journal.pbio. 1001655

Chapter 9 Profiling Transcriptomes of Manually Dissected Arabidopsis Embryos Ping Kao and Michael D. Nodine Abstract Genome-wide characterization of RNA populations in early flowering plant embryos can yield insights into the gene regulatory processes functioning during this formative phase of development. However, early embryonic transcriptomes are technically challenging to profile because of the low amount of RNA obtainable and potential RNA contamination from surrounding nonembryonic tissues. Here we provide a detailed protocol for collecting early Arabidopsis thaliana (Arabidopsis) embryos, generating mRNA sequencing (mRNA-seq) libraries, and basic data processing and quality controls of the resulting mRNAseq data. Key words Arabidopsis, Embryo, Transcriptome, RNA-seq

1

Introduction Genome-wide profiling of transcript populations in early flowering plant embryos can provide insights into the gene regulatory processes operating after gamete fusion. These fundamental processes include the transition from gametophytic to sporophytic gene expression programs, and the generation of the basic plant body plan. However, the low amounts of RNA obtainable from early embryos together with potential RNA contamination from surrounding endosperm and maternal seed coat tissues are a technical challenge to profiling transcriptomes from early plant embryos [1, 2]. Here, we present a comprehensive protocol for acquiring embryonic transcriptomes from Arabidopsis embryos. First, we introduce two protocols for acquiring embryos: hand-dissecting embryos from ovules [1] (i.e., the hand dissection method) and bulk rupturing ovules and picking embryos [3] (i.e., the bulk rupture method). We also provide a protocol for total RNA isolation from dissected embryos [1, 4] followed by a standard Smartseq2 protocol for mRNA-seq library construction [5, 6]. Last, we

Martin Bayer (ed.), Plant Embryogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2122, https://doi.org/10.1007/978-1-0716-0342-0_9, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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present a bioinformatic quality control workflow featuring the tissue enrichment test [2] to determine the level of RNA contamination from surrounding seed tissues in the acquired mRNA-seq dataset.

2

Materials

2.1 Early Embryo Isolation

1. Dissection microscope (recommend 10–25
for eyepiece, 100–200
total magnification). 2. Tungsten needles (0.25 mm). 3. Cavity slides. 4. RNAlater Stabilization Solution. 5. RNaseZAP. 6. Nuclease-free water. 7. Glass capillaries (with inner diameter 70–100 μm for early embryos). 8. 30 μm filters. 9. Micro pestles.

2.2 RNA Extraction, cDNA Construction, and mRNA-Seq Library Generation

1. Refrigerated centrifuge. 2. Heat block. 3. External RNA Controls Consortium (ERCC, ThermoFisher) (see Note 1). 4. TRIzol. 5. Chloroform. 6. Isopropanol. 7. Silanized or low-binding test tubes (e.g., LoBind tubes). 8. GlycoBlue (ThermoFisher). 9. Ethanol. 10. Thermocycler. 11. PCR tubes. 12. 10 mM dNTP. 13. 10 μM anchored oligo-dT (50 -/5Biosg/AAGCAGTGGTATCAACGCAGAGTACT30VN-30 ). 14. 10 μM template switching oligo (TSO, 50 -/5Biosg/AAGCAGTGGTATCAACGCAGAGTACrGrG+G-30 ). 15. SuperScript II reverse transcriptase and 5
first-strand buffer. 16. 40 U/μL RNase inhibitor, murine. 17. 100 mM DTT. 18. 5 M betaine.

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19. 100 mM MgCl2. 20. 10 μM isothermal PCR (ISPCR) primers (50 AAGCAGTGG TATCAACGCAGAGT-30 ). 21. KAPA HiFi HotStart ReadyMix (2
, KAPA Biosystems). 22. AMPure XP beads. 23. Elution buffer: 10 mM Tris–HCl, pH 8.5. 24. Fragment Analyzer. 25. High Sensitivity NGS Fragment Analysis Kit (Advanced Analytical). 26. 96-Well ABI sequencer plate (Biozym). 27. Nextera DNA library preparation kit (Illumina). 28. Nextera Index kit (Illumina). 29. DNA Clean & Concentrator-5 kit (Zymo Research). 2.3 mRNA-Seq Data Quality Control Workflow with Tissue Enrichment Test

3

1. R version 3.1 or later. 2. Python 2.7 or later.

Methods

3.1 Early Embryo Isolation (See Note 2) 3.1.1 Hand Dissection (Modified from [1])

1. Clean all equipment with RNaseZap. 2. Add 200 μL 10% RNAlater in the wells of a cavity slide (see Note 3). 3. Remove a silique from plant and slice open the valves longitudinally with tungsten needles. 4. Immerse ovules in 10% RNAlater on a glass slide for dissection. 5. Pin an ovule with a tungsten needle at chalazal end and make an incision at about one-third in length to the top of ovule with another tungsten needle (Fig. 1). 6. Press micropylar end gently with tungsten needle to push the embryo out of embryo sac. 7. Transfer the isolated embryo with a glass capillary to another well on cavity slide with 200 μL 10% RNAlater for washing. 8. Repeat step 7 wash 2
with different glass capillaries (see Note 4). 9. Transfer the washed embryo with a glass capillary into 30 μL of 100% RNAlater in a 1.5 mL LoBind tube. 10. Collect 20 embryos per replicate for mRNA-seq (see Note 5).

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Fig. 1 Schematic representation of embryo dissection. (a) Illustration of an ovule with key features. (b–d) Schematics corresponding to steps 5–7 in hand dissection 3.1.2 Bulk Rupture Method (Modified from [3])

1. Clean all equipment with RNaseZap. 2. Remove a silique from plant and slice open the valves longitudinally with tungsten needles. 3. Transfer ovules into 50 μL 10% RNAlater in a 1.5 mL LoBind tube. 4. Collect ovules from 10 to 15 siliques. 5. Gently rupture the ovules with a micro pestle to release the embryos (see Note 6). 6. Rinse the micro pestle with 100 μL 10% RNAlater. 7. Pipet ten times up and down to help release embryos. 8. Rinse 30 μm filter with 100 μL 10% RNAlater. 9. Filter the extract through rinsed 30 μm filter and collect in a 1.5 mL LoBind tube. 10. Rinse the filter with another 100 μL 10% RNAlater. 11. Centrifuge at 2000
g for 10 min at 4  C. 12. Remove supernatant and resuspend pellet with 50–100 μL 10% RNAlater. 13. Transfer the extract onto a cavity slide and screen for released embryos under microscope.

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14. Wash embryos by transferring them with a capillary to another well containing 200 μL 10% RNAlater in a cavity slide. 15. Repeat step 14 wash 2
with different glass capillaries (see Note 4). 16. Transfer the washed embryos with a capillary into 30 μL 100% RNAlater in a 1.5 mL LoBind tube. 17. Collect 20 embryos per replicate for mRNA-seq (see Note 5). 3.2 RNA Extraction, cDNA Construction, and mRNA-Seq Library Generation

1. Clean all equipment with RNaseZap.

3.2.1 RNA Extraction (See Note 7)

4. Vortex briefly and spin down.

2. Mix collected embryos with 500 μL TRIzol by vortexing (2
for 2 s each). 3. Incubate at 60  C for 30 min. 5. (Optional) Add ERCC spike-ins to help quantify transcript abundances (see Note 1). 6. Add 100 μL chloroform, vortex 2
for 2 s each, and incubate at RT for 3 min. 7. Centrifuge at 12,000
g for 15 min at 4  C to separate the phases. 8. Transfer the upper aqueous phase to a new 1.5 mL LoBind tube (see Note 8). 9. Add 1 volume of isopropanol and 1.5 μL GlycoBlue (22.5 μg), and mix by vortexing briefly. 10. Incubate at 20  C for 15–18 h (e.g., overnight). 11. Centrifuge at >20,000
g for 30 min at 4  C to pellet RNA. 12. Remove supernatant and wash the pellet by adding 500 μL of 75% ethanol and briefly vortex to mix. 13. Centrifuge at >20,000
g for 15 min at 4  C. 14. Repeat steps 12 and 13 1
. 15. Remove as much ethanol as possible, place tube on ice with lid open and air-dry for 10 min (see Note 9). 16. Resuspend with nuclease-free water (see Note 10). 17. Incubate at 60  C for 5–10 min to fully dissolve RNA. 18. Store RNA samples at 80  C or immediately proceed to cDNA synthesis.

3.2.2 cDNA Construction and NGS Library Preparation (Modified from Smart-seq2 Protocol [5, 6])

1. Mix 1 μL RNA sample, 1 μL 10 mM dNTP, and 1 μL 10 μM anchored oligo-dT in PCR tubes. 2. Denature at 72  C for 3 min and cool down on ice immediately. 3. Prepare reverse transcription master mix (RT mix) according to Table 1.

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Table 1 Reverse transcription master mix Component

Volume (μL)

Final concentration

SuperScript II reverse transcriptase (200 U/μL)

0.5

100 U

RNase inhibitor (40 U/μL)

0.25

10 U

Superscript II first-strand buffer (5
)

2

1


DTT (100 mM)

0.5

5 mM

Betaine (5 M)

2

1M

MgCl2 (100 mM)

0.6

6 mM

TSO (10 μM)

1

1 μM

Nuclease-free water

0.15

–

Total volume

7

–

Table 2 Thermocycler program for reverse transcription (RT) Step

Temperature ( C)

Time

Purpose

1

42

90 min

RT and template-switching

2

50

2 min

Denaturation of RNA secondary structures

3

42

2 min

Completion of RT

10 cycles of steps 2 and 3

4 5

70

6

4

15 min

Enzyme inactivation

Hold

Table 3 Preamplification PCR mix Component

Volume (μL)

Final concentration

KAPA HiFi HotStart ReadyMix (2
)

25

1


ISPCR primers (10 μM)

1

0.2 μM

Nuclease-free water

14

–

Total volume

40

–

4. Mix 7 μL RT mix with denatured sample by pipetting and spin down. Incubate in thermocycler with program described in Table 2. 5. Prepare preamplification PCR mix according to Table 3.

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Table 4 Thermocycler program for preamplification Step

Temperature ( C)

Time

1

98

3 min

2

98

15 s

3

67

20 s

4

72

6 min

5

N cycles of steps 2–4 (see Note 11)

6

72

5 min

7

4

Hold

6. Incubate in thermocycler with program according to Table 4. 7. Purify preamplification products with 1:1 (v/v) AMPure beads (see Note 12). (a) Mix 50 μL AMPure beads with PCR product by pipetting. (b) Incubate at room temperature for 10 min. (c) Place on magnetic stand for 5 min. (d) Carefully remove liquid without disturbing the beads. (e) Wash beads with 200 μL 80% ethanol and wait for 30 s. (f) Repeat the wash step 7e. (g) Remove all liquid and air-dry the beads for 5–10 min (see Note 13). (h) Take off magnetic stand and resuspend beads with 17 μL elution buffer. (i) Place on magnetic stand for 2 min. (j) Transfer 16 μL clear supernatant to a new PCR tube. 8. Take 2 μL and run Fragment Analyzer to check the quality and concentration of cDNA. (a) Follow the manufacturer’s instructions. (b) Good cDNA libraries should have a peak at 1–2 kb (Fig. 2). 9. For tagmentation, use Illumina Nextera DNA sample preparation kit or equivalent. 10. Prepare the tagmentation reaction according to Table 5. 11. Incubate at 55  C for 5 min and cool on ice. 12. Purify reaction with DNA Clean & Concentrator kit, following the manufacturer’s instructions. 13. Elute with 20 μL Resuspension buffer from Nextera kit.

Fig. 2 Examples of Fragment Analyzer cDNA profiles. (a) An example of a good cDNA profile. Most transcripts are >1000 bp. (b) An example of a degraded cDNA profile. Most transcripts are 800 bp fragments are not ideal especially for single-end sequencing. For detail optimization, please check [6] 3.3 mRNA-Seq Data Quality Control Workflow with Tissue Enrichment Test 3.3.1 Alignment and Quantification (All Resources Are Listed in Table 8)

1. Use pseudoaligner Kallisto v0.44.0 [7] to quantify resulting mRNA-seq datasets. 2. Build kallisto index for pseudoalignment. (a) Build FASTA file for each TAIR10 transcript model by running BEDTools getfasta v2.17.0 [8] with the TAIR10 GFF3 file and the TAIR10 genome sequence FASTA file (see Note 15). (b) Input the FASTA file with all transcript models and run kallisto index to generate index for pseudoalignment.

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Table 8 Bioinformatic resources Resource

Link

Kallisto [7]

https://pachterlab.github.io/kallisto/download

TAIR10 annotation release version 40 (GFF3)

ftp://ftp.ensemblgenomes.org/pub/plants/release40/gff3/ arabidopsis_thaliana/Arabidopsis_thaliana.TAIR10.40.gff3.gz

TAIR10 genome sequence (FASTA)

ftp://ftp.ensemblgenomes.org/pub/plants/release40/fasta/ arabidopsis_thaliana/dna/Arabidopsis_thaliana.TAIR10.dna. toplevel.fa.gz

TAIR10 cDNA (FASTA)

https://www.arabidopsis.org/download_files/Sequences/TAIR10_ blastsets/TAIR10_cdna_20101214_updated

BEDTools [8]

https://code.google.com/archive/p/bedtools/

SAMtools [9]

http://samtools.sourceforge.net/

Cutadapt [10]

https://cutadapt.readthedocs.io/en/v1.9/installation.html

Tissue enrichment test [2]

https://github.com/Gregor-Mendel-Institute/tissue-enrichment-test

3. Convert sequencing BAM files to FASTQ files with SAMtools v1.4 [9]. 4. Trim adapter sequences with Cutadapt v1.9.1 [10] on FASTQ files with a minimum match length of five bases. Also use Cutadapt to trim all oligo-A or oligo-T sequences 5 bases long from the ends of reads. All trimmed sequencing reads >18 bases long can be used as input for Kallisto quantification. 5. Align trimmed FASTQ file by executing kallisto quant program using the index generated in step 2. kallisto quant returns a TSV file of transcript isoform levels in raw counts and transcripts per million (TPM). (a) Run paired-end samples with default settings. (b) Run single-end samples with argument --single and estimate the library fragment size distribution with argument --fragment-length 200 --sd 100 (see Note 16). 6. Sum TPM of all isoforms to estimate TPM for each gene (see Note 17). 3.3.2 Tissue Enrichment Test

1. Examine purity of embryonic RNA sample with the tissueenrichment-test from GitHub [2] (see Note 18). (a) Define time points of sequencing samples in the . description file. (b) Input the gene-level expression .tsv file and the .description file and run tissue-enrichment-test.

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(c) The tissue-enrichment-test will return a cumulative frequency distribution plot for each sample, a p-values table for the dataset, and an enrichment score heatmap for the dataset. 2. Exclude samples showing enrichment for nonembryonic tissue from subsequent analyses.

4

Notes 1. ERCC poly(A) spike-ins provide references to enable absolute quantification of mRNA molecules. The ERCC reads should be 2–5% of all aligned reads, and the amount of ERCC added to the sample should be calculated accordingly. Please find the recommended amounts of ERCC for each stage in Table 9. 2. We present two strategies for isolating early embryos: hand dissection and bulk rupture methods. While the hand dissection is more consistent, the bulk rupture method allows more efficient embryo collection. 3. Replace the 10% RNAlater every 30 min or after five series of washes. 4. Be careful not to break or lose embryos during washes. A minimum of three washes is recommended to minimize transcript contamination from surrounding tissues. 5. It is preferable to proceed to RNA isolation immediately after collection. If the RNA cannot be isolated immediately after collection, then the collected embryos can be stored in 100% RNAlater at 4  C for a few days. Table 9 Recommendations for amounts of ERCC spike-ins to add to embryonic samples Stage

ERCC dilution

μL per sample (20 embryos)

Preglobular

1:40000

2

Globular

1:2000

2

Early heart

1:2000

5

Late heart

1:1000

4

Early torpedo

1:400

1.3

Late torpedo

1:400

1.8

Bent cotyledon

1:400

5.3

Mature green

1:400

3.7

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6. The force and duration required to rupture the ovules and release intact embryos must be determined by the user. In general, stop and examine the homogenate after rupturing the ovules every 5
and repeat until no ovules remain intact. 7. The whole procedure is done on ice unless otherwise indicated. 8. Do not disturb or transfer the interphase. One should recover about 330 μL. 9. Do not overdry the pellet or it will be difficult to resuspend. 10. In general, 5–12 μL should be sufficient for resuspension. 11. In our experience, we use 11 PCR cycles for 5 ng, 13 PCR cycles for 1 ng, 14 PCR cycles for 0.5 ng and 16 PCR cycles for 0.1 ng total RNA input. Keep the cycle number as low as possible to prevent cDNA overamplification. 12. Make sure AMPure beads are at room temperature before use. If necessary, use 1:0.8 (sample–AMPure beads ratio) to reduce the amount of primer dimer isolated. 13. Air-dry the beads until small cracks appear on the pellet. Do not overdry the beads, or else they will be difficult to resuspend. 14. The cycle number can be increased slightly but should be minimized to prevent library overamplification. 15. Alternatively, the TAIR10 cDNA FASTA file (Table 8) can be downloaded and used to build a kallisto index. 16. The example here assumes that the fragment distribution centers at 200 bp with a 100 bp standard deviation. These parameters can be adjusted according to the actual parameters of each library. 17. Use the R package tximport or any other tools to combine transcript-level expression to gene-level expression. 18. Imported TPM expression levels are converted to percentile ranks in the test. The null hypothesis that tissue type A is not enrich in sample X is tested by Wilcoxon rank-sum test with wilcox.test() in R. For further details and examples please check [2] or the corresponding Github website (https://github. com/Gregor-Mendel-Institute/tissue-enrichment-test).

Acknowledgments ¨ sterreichische Akademie der This work was supported by the O Wissenschaften (Austrian Academy of Sciences). We thank Michael Schon and Falko Hofmann for advice regarding data analyses. Ping Kao also personally thanks Kotoha Tanaka, Kaori Sakuramori, and Uzuki Shimamura for their support.

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References 1. Nodine MD, Bartel DP (2012) Maternal and paternal genomes contribute equally to the transcriptome of early plant embryos. Nature 482:94–97. https://doi.org/10.1038/ nature10756 2. Schon MA, Nodine MD (2017) Widespread contamination of Arabidopsis embryo and endosperm transcriptome data sets. Plant Cell 29:608–617. https://doi.org/10.1105/tpc. 16.00845 3. Raissig MT, Gagliardini V, Jaenisch J et al (2013) Efficient and rapid isolation of earlystage embryos from Arabidopsis thaliana seeds. J Vis Exp. 76:e50371. https://doi.org/ 10.3791/50371 4. Nodine MD, Bartel DP (2010) MicroRNAs prevent precocious gene expression and enable pattern formation during plant embryogenesis. Genes Dev 24:2678–2692. https://doi.org/ 10.1101/gad.1986710 5. Picelli S, Bjo¨rklund A˚K, Faridani OR et al (2013) Smart-seq2 for sensitive full-length transcriptome profiling in single cells. Nat Methods 10:1096–1098. https://doi.org/10. 1038/nmeth.2639

6. Picelli S, Faridani OR, Bjo¨rklund A˚K et al (2014) Full-length RNA-seq from single cells using smart-seq2. Nat Protoc 9:171–181. https://doi.org/10.1038/nprot.2014.006 7. Bray NL, Pimentel H, Melsted P, Pachter L (2016) Near-optimal probabilistic RNA-seq quantification. Nat Biotechnol 34:525–527. https://doi.org/10.1038/nbt.3519 8. Quinlan AR, Hall IM (2010) BEDTools: a flexible suite of utilities for comparing genomic features. Bioinformatics 26:841–842. https:// doi.org/10.1093/bioinformatics/btq033 9. Li H, Handsaker B, Wysoker A, Fennell T, Ruan J, Homer N, Marth G, Abecasis G, Durbin R, 1000 Genome Project Data Processing Subgroup (2009) The sequence alignment/map format and SAMtools. Bioinformatics 25:2078–2079. https://doi. org/10.1093/bioinformatics/btp352 10. Marcel M (2011) Cutadapt removes adapter sequences from high-throughput sequencing reads. EMBnet J 17:10–12

Chapter 10 Laser-Assisted Microdissection of Plant Embryos for Transcriptional Profiling Ana Marcela Florez-Rueda, Lucas Waser, and Ueli Grossniklaus Abstract Transcriptomic studies have proven powerful and effective as a tool to study the molecular underpinnings of plant development. Still, it remains challenging to disentangle cell- or tissue-specific transcriptomes in complex structures like the plant seed. In particular, the embryo of flowering plants is embedded in the endosperm, a nurturing tissue, which, in turn, is enclosed by the maternal seed coat. Here, we describe laser-assisted microdissection (LAM) to isolate highly pure embryo tissue from whole seeds. This technique is applicable to virtually any plant seed, and we illustrate the use of LAM to isolate embryos from species of the Boechera and Solanum genera. LAM is a tool that will greatly help to increase the repertoires of tissuespecific transcriptomes, including those of embryos and parts thereof, in nonmodel plants. Key words Boechera, Embryo, Laser-assisted microdissection, RNA-Seq, Solanum, Tissue specificity, Transcriptome

1

Introduction Studies of the transcriptional basis of plant embryogenesis have so far been largely restricted to the model plant Arabidopsis thaliana [1–12], but some analyses have also been performed in maize [13– 15] and rice [16, 17]. Most of these studies described expression patterns in different regions of the embryo or at different stages of embryogenesis [1–7, 13, 14, 16] or focused on the regulation and dynamics of allele-specific parental contributions to gene expression in the early embryo [8–12, 15, 17]. The seed is composed of tissues that differ in ploidy as well as parental contributions [18, 19]. Thus, to properly study the embryonic transcriptome, the embryo must be removed from its accompanying tissues in the seed, the endosperm and the seed coat. Methods that involved manual dissection of embryos from their surrounding tissues have been developed [20] and variations thereof are widely used (e.g., [4, 5]). Other methods involve the use of transgenic lines allowing for the profiling of individual cell and tissue types, such as

Martin Bayer (ed.), Plant Embryogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2122, https://doi.org/10.1007/978-1-0716-0342-0_10, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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fluorescence-activated cell or nuclei sorting (FACS, FANS) [3, 21, 22]. Labeling and affinity purification of nuclei (INTACT) [23, 24] or polysomes (TRAP) [25, 26] also allow for the analysis of gene expression at a finer scale. Because all these methods require the expression of transgenes under highly specific embryonic promoters, their use has, so far, largely been restricted to Arabidopsis. Concerns have been raised about potential contamination of embryonic samples with debris from surrounding maternal tissues [27]. Given that many genes are expressed in both embryos and seed coat [28], it is not easy to ascertain contamination without independent confirmation of tissue-specific expression. Nonetheless, many transcriptome datasets obtained from manually dissected embryos or embryonic nuclei recovered by FANS detected genes that were also found expressed in the seed coat, while embryo and endosperm datasets obtained by LAM showed less expression of such genes [27]. This highlights that, in comparison to other methods, LAM allows for the isolation of very precise and highly pure samples for transcriptome studies of reproductive tissues [29– 32]. Another important advantage of LAM is that, in contrast to other methods allowing for the profiling of individual cell and tissue types, such as INTACT and TRAP, LAM does not require the generation of transgenic marker lines for cell capture. This makes LAM applicable to any plant embryo or tissue of interest without prior molecular knowledge and is, thus, a powerful method to study nonmodel organisms. Here, we present an LAM methodology adapted to plant embryos. We have implemented this protocol in species of the genus Boechera, a member of the Brassicaceae closely related to Arabidopsis, and in wild tomatoes, belonging to the genus Solanum, in order to exemplify the versatility and broad applicability of LAM for obtaining transcriptomic data of high purity and quality from virtually any plant embryo.

2

Materials

2.1 Removal of RNAses from Working Material and Equipment

1. Cleaning agent for the removal of RNAses (e.g., RNaseZAP™ R2020).

2.2 Tissue Collection and Fixation

1. Boechera and wild tomato plants with fruits and siliques at the desired stage.

2. RNase-free water. 3. Ethanol 70%.

2. Farmer’s fixation solution: 90% (v/v) ethanol, 10% (v/v) acetic acid. 3. Microcentrifuge tubes. 4. Ice and insulated box.

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5. Forceps. 6. Scissors. 7. Double-sided tape. 8. Glass slides. 9. Razor blade. 10. Excicator/vacuum chamber. 2.3 Tissue Embedding, Blocking, and Slide Preparation

1. Histology cassettes Cassettes).

(e.g.,

Tissue-Loc™

HistoScreen™

2. Pencil. 3. Ice and insulated box. 4. RNase-free water. 5. 70% (v/v) ethanol (EtOH). 6. Large glass staining container. 7. Embedding machine (e.g., HistoCore Pearl). 8. Blocking station (e.g., HistoCore Arcadia H EG1150C). 9. Flat bottom plastic weighing dish. 10. Preparation needle. 11. Forceps. 12. Spatula. 13. Razor blade. 14. Sample holder for microtome. 15. Paraffin. 16. Bottom illuminated table. 17. Ethanol torch. 18. Rotary microtome (e.g., Leica RM2255). 19. Microtome steel blades. 20. Black cardboard. 21. Brush. 22. LCM-slides: nuclease-free PET membrane slides 1.4 μm. 23. Heating plate. 24. Sterile pipette. 25. Staining dish. 26. Xylol (or Histo-Clear).

2.4 Laser-Assisted Microdissection (LAM)

1. Compressed air spray (e.g., DUST OFF 67). 2. Laser capture microscope (LCM) Cellcut IX71, consisting of an Olympus IX81 inverted microscope with an Olympus light box IX2-UCB.

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3. MMI Cellcut® laser—MMI AG (Glattbrugg, Switzerland). 4. Software package MMI CellTools version 4.4.9.—MMI AG (Glattbrugg, Switzerland). 5. MMI Isolation caps transparent 0.5 mL Prod. No. 50204. 6. Glass slides for microscopy.

3

Methods

3.1 Removal of RNAses from Working Material and Equipment

1. To ensure an RNAse-free environment, bake any glassware for 8 h at 180  C.

3.2 Tissue Collection and Fixation

1. Prepare microcentrifuge tubes filled with farmer’s fixation solution on ice (see Note 1).

2. Treat any working material and equipment to be used with RNAse removal cleaning agent. Remove the cleaning agent with RNAse-free water and, subsequently, clean the surfaces further with 70% EtOH.

2. Remove the tissue (wild tomato fruits or developing Boechera siliques) containing embryos at the targeted developmental stage with forceps and scissors (see Note 2). 3. For Boechera embryos, place the silique on a glass slide with double-sided tape and carefully cut open the valves along the replum [33] (see Note 3). For wild tomato embryos cut the fruits in half with a razor blade. 4. Immerse the tissue rapidly in the fixation solution on the microcentrifuge tube placed on ice. 5. Place the opened tubes with the samples in a desiccator filled with ice and vacuum infiltrate for 30 min with a release after 15 min. 6. Store overnight at 4  C. 3.3 Tissue Embedding, Blocking, and Slide Preparation

1. Transfer fixed tissue to ice-cold, nuclease-free 70% EtOH. Proceed to embedding on the same day. 2. Prepare a large staining container glass filled with ice-cold, nuclease-free 70% EtOH that accommodates all samples to process. Transfer tissue to embedding cassettes, firmly close the cassettes, and label them appropriately with a pencil. Deposit the cassettes with the samples in the ice-cold ethanol glass container. Work fast to avoid drying of the tissue (see Note 4). 3. Transfer cassettes to sample basket and run embedding machine overnight following the manufacturer’s instructions. Recommended standard settings are 1 h 70% EtOH, three times 1 h 90% EtOH, three times 1 h 100% EtOH, two times

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1 h 100% Xylol, and one time 1 h 15 min 100% Xylol at room temperature, followed by infiltration with paraffin wax two times for 1 h and one time for 3 h at 56  C. 4. Set blocking station to preheat several hours before the planned start of use. 5. Transfer the sample basket to the preheated blocking station. 6. Using the blocking station, fill a plastic weighing dish with paraffin wax at 56  C. Lift the lid of the paraffin bath, pick up only one cassette at a time, and transfer the samples from the cassettes to the liquid paraffin wax at 56  C in the balancing tray using a pair of tweezers. Work fast to avoid solidification of the wax around the samples before placing them in the weighing dishes (see Note 5). 7. Position samples well interspaced among each other and in a position that maximizes embryo recovery when preparing thin sections (see Note 6). 8. After letting the paraffin completely solidify at room temperature, store at 4  C until further processing. 9. Remove paraffin block from weighing dish, place on a bottom illuminated table, and cut the paraffin block into pieces containing the target sample. 10. Coat the grid surface of a labeled sample holder by adding melted paraffin using a spatula and ethanol torch. While the paraffin is still soft, adhere the previously prepared block of paraffin with samples onto the sample holder. 11. Trim the edges of the paraffin block and take care to maintain parallel edges in order to have straight ribbons when preparing thin sections (see Note 7). Before proceeding with sectioning, place samples at 4  C to harden paraffin for at least 10 min. 12. Set heating plate to 42  C. 13. Safely clamp in the sample holder into a microtome equipped with a fresh microtome steel blade. Set the microtome to 7–8 μm and cut the block into ribbons (see Note 8). Refer to manufacturer’s instructions for using the microtome. Place the ribbons onto a box with black cardboard using brush and forceps. 14. Using the brush, forceps, and razor blade, divide the ribbons into pieces containing embryos. Observe ribbons under stereoscope if needed to identify the desired samples. 15. Place an adequate number of LCM slides onto the hotplate (42  C) with the flat surface covered by a PET membrane facing up. 16. Using the pipette, cover the membrane area with nuclease-free water.

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17. Place ribbon segments of interest onto the nuclease-free water droplet and, if necessary, arrange them using the brush. Let the paraffin and the sliced sample expand for proper microscopic visualization for a couple of minutes. Rapidly, discard the nuclease free water droplet in a fluid motion, leaving only the paraffin ribbon segments on the membrane (see Note 9). 18. Place LCM slides back onto the hotplate and cover them to reduce the risk of contamination. Keep the slides on the hotplate until completely dry (see Note 10). 19. Prepare two staining dishes filled with fresh Xylol (or HistoClear) under a fume hood. Remove wax from the prepared slides by two consecutive washes of 15 min each. Let the slides dry under the hood for another 15–20 min and proceed to the LCM immediately (see Note 11). 3.4 Laser-Assisted Microdissection (LAM) on the Laser Dissection Microscope (LCM)

1. Remove dust and small particles from the LCM using the compressed air spray. 2. Turn on the devices for the LCM (computer, microscope, laser control box) and start the MMI software (see Note 12). 3. Calibrate objectives to be used and laser settings, refer to the manufacturer’s instructions for details. 4. Cover the ready to laser LCM slides by placing a nuclease-free glass microscopy slide on the flat surface with the PET membrane containing the sectioned samples. Place the assembled slides on the microscope with the microscopy slide facing down and the LCM slide facing up. 5. Equip the cap holder with a collection cap and position it on the magnetic lift mechanism of the microscope. 6. Select the 4 Objective. On the MMI software, make an overview scan of the slide. 7. On low magnification (4), identify embryos on the slide and define location pins; this will allow you to move back to the exact position for the following cutting step. 8. Change to a higher magnification (10–20) and go back to the previously defined and pinned embryo locations (Fig. 1a–f). 9. Lower the collection cap, making sure that it makes contact on an empty space with the target embryonic tissue; this will ensure proper adherence of the tissue to the cap. Do not collect tissue over previously collected tissue as this will cause detaching from the collection cap. Use the MMI software tools to draw the desired shape for the laser to cut (see Note 13) (Fig. 1g–i). 10. Press the cut function, observe and control that the laser precisely cuts the area of interest.

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Fig. 1 Microscopic observations of embryos under the laser capture microscope (LCM). (a, d, g and j) Solanum peruvianum, (b, e, h, and k) Solanum chilense, (c, f, i, and l) Boechera holboelli. Embryos are clearly discernible in dry, thin sections of seeds (a–c) and their outlines nicely are demarcated at higher magnification (d–e). Accurate drawing of the embryo boundaries for laser cutting secures a high specificity of transcriptomic data (g–h). Embryonic sections are precisely excised after laser-assisted microdissection (LAM) (j–l)

11. Raise the collection cap and make sure that the embryo slice has been collected (see Note 14) (Fig. 1j–l). 12. Repeat from step 8 until all embryos are collected, for each defined pin on a given LCM slide. 13. Continue with the rest of the LCM slides prepared for a given session. Repeat from step 4 until all prepared LCM slides are processed (see Note 15). 14. Select the 4 Objective. Examine the collection cap for an overview of the collection and to identify possible dirt; if dirt is found, remove it (see Note 16). 15. Close the cap and store at Note 17).

80  C until RNA extraction (see

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Fig. 2 Quality control through Agilent 2100 Bioanalyzer assays. (a) Representative trace of 47 pg LAM RNA in a Plant RNA Pico Chip. (b) Trace of amplified cDNA from B. holboelli embryo RNA run in a High sensitivity DNA Chip. (c) Trace of a B. holboelli embryo Nextera XT library run in a High sensitivity DNA Chip

3.5 RNA Extraction, cDNA Synthesis, and Library Preparation for Illumina RNA Sequencing

1. Extract RNA using a kit designed to optimize recovery from low input samples. We have been successful in the described systems with a minimum of 50 slices of embryos. We routinely use Arcturus™ PicoPure™ RNA isolation kit (Applied Biosystems by Thermo Fisher Scientific, USA) with consistently good results. Follow the manufacturer’s instructions (see Note 18). 2. Validate RNA extraction by running Agilent 2100 Bioanalyzer Plant RNA Pico Chip, representative traces of picogram quantities of RNA can be detected (Fig. 2a) (but see Note 19). 3. Amplify cDNA from the extracted RNA using as a baseline the Smart-Seq2 protocol [34]. We routinely use the SMART-Seq® v4 Ultra® Low Input RNA Kit for Sequencing (Clontech Laboratories Inc., USA) with consistently good results. Follow the manufacturer’s instructions.

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4. Validate cDNA amplification output and quality by running Agilent 2100 Bioanalyzer High sensitivity DNA Chip (Fig. 2b) or an Agilent High Sensitivity D1000 ScreenTape. 5. Prepare libraries for sequencing using the Nextera® XT (Illumina Inc., USA) library preparation kit. Follow the manufacturer’s instructions. 6. Validate library output and quality by running Agilent 2100 Bioanalyzer High sensitivity DNA Chip (Fig. 2c) or an Agilent High Sensitivity D1000 ScreenTape. 7. Sequence your transcriptomic libraries on an Illumina Sequencing machine. We have obtained good quality sequencing data on the Illumina HiSeq2500 and Illumina HiSeq4000 platforms.

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Notes 1. It is critical to maintain the samples from collection upon embedding at Import ASCII function (Fig. 12).

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Fig. 6 Loading and size adjustment of an image file

3. Fluorescence lifetime calculation: Choose your folder from the margarita directory and import the ht3 file. Import the yellow channel first, then the red channel. Select the image of interest, set the display as X pixel in field X and Y pixel in field Y, then a window containing your sample will open). To change the displayed heat-map data, select photon number for YFP fluorescence intensity or tau for YFP fluorescence lifetime in weighting field (Fig. 13). 4. After the 100-photon-per-pixel background subtraction, nuclear areas were selected based on their morphology. Select the nuclei of interest by dragging the blue and red crosshairs to crop to the nucleus (Fig. 14, see Note 14). 5. Save the data in a new folder and open the channel (Fig. 15).

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Fig. 7 Fit settings

6. On the X/Y icons assign X ¼ tau (green) and Y ¼ Sg/Sr (i.e., signal ratio between green and red). Change the bin values as marked in the above image with 100 maximum at the Y axis and 1.5–4 at the X axis and save the histogram. 7. When finished measuring, clear all data, and start measuring the next sample. 8. For each measurement/per nuclear, the data obtained in pixels represent cellular fluorescence lifetimes. The obtained pixels were computed by least-square fitting the Gaussian peaks of each cells’ lifetime distributions. 9. Copy the lifetime values into an excel file. 10. For each cell type, lifetimes at the same cell position were pooled from independent measurements.

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Fig. 8 IRF and background file

11. Calculate the means of the obtained values from the donoronly and the FRET samples. 12. Calculate the reduction of the lifetime (Δτ) between donor only and the FRET samples corresponding to each cell position (Δτ ¼ τD  τDA). 13. Preform a Student’s t-test with critical value of p < 0.01 to evaluate the significances, between donor-only and FRET samples at specific cell positions. 3.7.3 Fluorescence Intensity Image

Data can be represented as a fluorescence intensity image where color distribution of the FLIM image is applied automatically. The resulting image then represents the fluorescence lifetime values of all individual pixels. For both donor and FRET samples, images are displayed as a false color-coded fluorescence lifetime image (Fig. 2f and d). The color range of the fluorescence lifetime can be set, in

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Fig. 9 Pop-up window

this example the limits were ranging from red (long lifetime, 3.2 ns) to dark blue (low lifetime, 2.4 ns). In the case of strong interaction, this representation allows a visual comparison between donor-alone versus donor–acceptor samples. However, this representation is not quantitative. 3.7.4 Representation of Lifetime Reduction

To visually assess the distributions of the measured fluorescence lifetime, τD and τDA can be plotted in box plots (Fig. 2h). Box plots can be readily generated by open source software like RStudio (https://www.rstudio.com/) or free online box plot generators like BoxPlotR (http://shiny.chemgrid.org/boxplotr/). The following procedure is dedicated to BoxPlotR (see Note 15). 1. In the Data Upload tab, choose Upload file or Paste data. Make sure the first row of data depicts column headers, and choose the correct Delimiter (e.g., pasted Excel spreadsheet often uses

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Fig. 10 Setting the decay model

Tab delimiter). A spreadsheet will be generated on the webpage upon successful data upload (see Note 16). 2. In the Data visualization tab, a Tukey box plot is displayed by default, where the box represents the distribution between the first and third quartiles (Q1 and Q3) of the data and the line in the box marks the median. The Tukey whiskers marks the closest data points within the 1.5
interquartile range (IQR ¼ Q3  Q1) from Q1 and Q3, and the circles depict Tukey’s outliers. 3. Modify the plot display (data representations like adding average, whisker definition, number of data points and aesthetics like dimension, color, label, etc.) using the toolbox on the left of the webpage until satisfied.

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Fig. 11 Fluorescence lifetime calculation

4. Download the boxplot and save the Box plot statistics spreadsheet. 5. The mean Δτ (and FRET efficiency E ¼ Δτ/τD) can also be plotted in bar charts (Fig. 2i). Error bars representing propagated standard error of mean (SEM or σ x ) can be calculated as: qffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi σ Δτ ¼ ðσ τD Þ2 þ ðσ τDA Þ2 or

s ffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi 2  2 σ τDA σ τD σ E ¼ E þ τD τDA

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Fig. 12 File import in Margarita

where SEM of τD and τDA is retrieved from the standard deviation s divided by the square root of the number of data points n: pffiffiffiffiffi στ ¼ s τ = nτ 6. The p-value from Student’s t-test (or other tests that do not require near-normal distribution, like Wilcoxon–Mann–Whitney test or Kolmogorov–Smirnov test) between τD and τDA can also be plotted on the secondary axis of the bar chart (Fig. 2i).

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Fig. 13 Changing the display heat map data

4

Notes 1. Cover glass thickness should match the requirement of the objective lenses for optimal photon collection. 2. Make sure that the plants are healthy as stress can affect embryo development and influence fluorescence levels as well as PPI. 3. Double-sided tape can be put either on the back of an empty petri dish or on top of a slide (Fig. 1a). 4. It is important to use double-sided tape to avoid movement of the silique. This will facilitate silique opening and embryo isolation. 5. The use of a pap pen is to keep the embryos within a limited area on the slide and to avoid embryo dispersal to facilitate finding them. 6. If using 5%glucose, embryos should be visualized immediately after isolation, optimally within 20 min and cannot be kept for hours or overnight for imaging. Experiments that would require longer acquisition time and time lapse imaging, modifications should be implemented as embryos should be then mounted in special media [21].

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Fig. 14 Selection of nuclei

7. It is important to check the signal intensity before starting the experiment. If possible, use a fluorescent (stereo) microscope with good sensitivity to check fluorescence levels per embryo. We found that this can significantly increase the FLIM measurement efficiency. Make sure that you have homozygous lines to ensure homogenous expression. 8. Solutions can be kept up to 1 month at 4  C. 9. The instrument must be calibrated for every measuring day and the IRF should be measured before initiating embryo preparation. 10. Create a document file where all the measurements including the calibration should be recorded in order to retrieve the measurement settings including laser power, number of frames, and objective used during the measurements. This will ensure consistency between experiments. 11. For FLIM acquisition we used a confocal laser scanning microscope (Zeiss LSM 780) equipped with a single-photon counting device with picosecond time resolution.

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Fig. 15 Fluorescence lifetime calculation

12. For reliable fitting of fluorescence lifetime, photon count per pixel should be at least ~1000. Adjust excitation intensity and time series length to harvest sufficient photons with minimal photobleaching and focal drift. 13. Measure at least 40 samples per experiment and repeat the experiment at least three times. During each experiments the setting should not be altered; the image size, laser power, number of frame, and pixel time should be kept the same for all the samples and between donor and FLIM samples. 14. We noticed strong effects of shorter lifetimes at the periphery of embryonic nuclei, for example, in the form of “green halo” as shown in the above screenshot. Since this “halo” is also prominent in donor-only samples where no FRET-induced lifetime reduction occurs, we concluded that the rings

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surrounding embryonic nuclei are largely contributed by low-lifetime background inteference, therefore they should not be included for analysis. 15. BoxPlotR considers the first row of the spreadsheet as column header by default, even if data points were entered (in such a case, a letter X is added in front of the number on the plot). 16. Box plots can also be created by commonly used spreadsheet software like Microsoft Excel by following online tutorials (e.g., https://support.office.com/en-us/article/create-abox-plot-10204530-8cdf-40fe-a711-2eb9785e510f or http://www.real-statistics.com/descriptive-statistics/boxplots-with-outliers/). Problems and considerations to take into account: (a) Avoid bleaching by using low laser power. Laser power at the objective should not exceed 1–2 μW. (b) Samples may move during image acquisition making it difficult to acquire focused samples, this can be caused either by having samples mounted in excessive liquid. Focal drift can also cause this problem. Using an isolation table and an incubation chamber can help alleviating this problem. (c) Low expressed proteins have low photon counts which will increase the background contribution. The low lifetime caused by the high autofluorescene in the embryo will cause false positive. An increase of frame number to accumulate more photon could be a solution. References 1. Chakrabortty B, Willemsen V, de Zeeuw T, Liao CY, Weijers D, Mulder B, Scheres B (2018) A plausible microtubule-based mechanism for cell division orientation in plant embryogenesis. Curr Biol 28(19):3031. https://doi.org/10.1016/j.cub.2018.07.025 2. Budnik B, Levy E, Harmange G, Slavov N (2018) SCoPE-MS: mass spectrometry of single mammalian cells quantifies proteome heterogeneity during cell differentiation. Genome Biol 19:161. https://doi.org/10.1186/ s13059-018-1547-5 3. Lee HW, Kyung T, Yoo J, Kim T, Chung C, Ryu JY, Lee H, Park K, Lee S, Jones WD, Lim DS, Hyeon C, Heo WD, Yoon TY (2013) Real-time single-molecule co-immunoprecipitation analyses reveal cancer-specific Ras signalling dynamics. Nat Commun 4:1505. https://doi.org/10.1038/ ncomms2507

4. Long YC, Stahl Y, Weidtkamp-Peters S, Smet W, Du YJ, Gadella TWJ, Goedhart J, Scheres B, Blilou I (2018) Optimizing FRETFLIM Labeling conditions to detect nuclear protein interactions at native expression levels in living Arabidopsis roots. Front Plant Sci 9:639. https://doi.org/10.3389/fpls.2018. 00639 5. Bucherl CA, Bader A, Westphal AH, Laptenok SP, Borst JW (2014) FRET-FLIM applications in plant systems. Protoplasma 251 (2):383–394. https://doi.org/10.1007/ s00709-013-0595-7 6. Bhat RA (2009) FRET and FLIM applications in plants. Lab Tech Biochem Mol 33:413–445. https://doi.org/10.1016/S0075-7535(08) 00010-7 7. Bucherl CA, van Esse GW, Kruis A, Luchtenberg J, Westphal AH, Aker J, van Hoek A, Albrecht C, Borst JW, de Vries SC

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(2013) Visualization of BRI1 and BAK1 (SERK3) membrane receptor Heterooligomers during Brassinosteroid Signaling. Plant Physiol 162(4):1911–1925. https://doi.org/ 10.1104/pp.113.220152 8. Crosby KC, Pietraszewska-Bogiel A, Gadella TWJ, Winkel BSJ (2011) Forster resonance energy transfer demonstrates a flavonoid metabolon in living plant cells that displays competitive interactions between enzymes. FEBS Lett 585(14):2193–2198. https://doi.org/10. 1016/j.febslet.2011.05.066 9. Long YC, Goedhart J, Schneijderberg M, Terpstra I, Shimotohno A, Bouchet BP, Akhmanova A, Gadella TWJ, Heidstra R, Scheres B, Blilou I (2015) SCARECROWLIKE23 and SCARECROW jointly specify endodermal cell fate but distinctly control SHORT-ROOT movement. Plant J 84 (4):773–784. https://doi.org/10.1111/tpj. 13038 10. Smaczniak C, Immink RGH, Muino JM, Blanvillain R, Busscher M, Busscher-Lange J, Dinh QD, Liu SJ, Westphal AH, Boeren S, Parcy F, Xu L, Carles CC, Angenent GC, Kaufmann K (2012) Characterization of MADSdomain transcription factor complexes in Arabidopsis flower development. Proc Natl Acad Sci U S A 109(5):1560–1565. https://doi. org/10.1073/pnas.1112871109 11. Stahl Y, Grabowski S, Bleckmann A, Kuhnemuth R, Weidtkamp-Peters S, Pinto KG, Kirschner GK, Schmid JB, Wink RH, Hulsewede A, Felekyan S, Seidel CA, Simon R (2013) Moderation of Arabidopsis root stemness by CLAVATA1 and ARABIDOPSIS CRINKLY4 receptor kinase complexes. Curr Biol 23(5):362–371. https://doi.org/10. 1016/j.cub.2013.01.045 12. Weidtkamp-Peters S, Stahl Y (2017) The use of FRET/FLIM to study proteins interacting with plant receptor kinases. Methods Mol Biol 1621:163–175. https://doi.org/10.1007/ 978-1-4939-7063-6_16 13. Vogel SS, Blank PS, Koushik SV, Thaler C (2009) Spectral imaging and its use in the measurement of Forster resonance energy transfer in living cells. Lab Tech Biochem Mol 33:351–394. https://doi.org/10.1016/ S0075-7535(08)00008-9

14. Xing SP, Wallmeroth N, Berendzen KW, Grefen C (2016) Techniques for the analysis of protein-protein interactions in vivo. Plant Physiol 171(2):727–758. https://doi.org/10. 1104/pp.16.00470 15. Cui HC, Levesque MP, Vernoux T, Jung JW, Paquette AJ, Gallagher KL, Wang JY, Blilou I, Scheres B, Benfey PN (2007) An evolutionarily conserved mechanism delimiting SHR movement defines a single layer of endodermis in plants. Science 316(5823):421–425. https:// doi.org/10.1126/science.1139531 16. Cui HC, Kong DY, Liu XW, Hao YL (2014) SCARECROW, SCR-LIKE 23 and SHORTROOT control bundle sheath cell fate and function in Arabidopsis thaliana. Plant J 78 (2):319–327. https://doi.org/10.1111/tpj. 12470 17. Nakajima K, Sena G, Nawy T, Benfey PN (2001) Intercellular movement of the putative transcription factor SHR in root patterning. Nature 413(6853):307–311. https://doi. org/10.1038/35095061 18. Long YC, Smet W, Cruz-Ramirez A, Castelijns B, de Jonge W, Mahonen AP, Bouchet BP, Perez GS, Akhmanova A, Scheres B, Blilou I (2015) Arabidopsis BIRD zinc finger proteins jointly stabilize tissue boundaries by confining the cell fate regulator SHORTROOT and contributing to fate specification. Plant Cell 27(4):1185–1199. https://doi.org/ 10.1105/tpc.114.132407 19. Long Y, Tahl YS, Weidtkamp-Peters S, Postma M, Zhou W, Oedhart JG, SanchezPerez MI, Adella TWJG, Simon R, Scheres B, Blilou I (2017) In vivo FRET-FLIMr reveals cell-type-specific protein interactions in Arabidopsis roots. Nature 548(7665):97. https:// doi.org/10.1038/nature23317 20. Sanderson MJ, Smith I, Parker I, Bootman MD (2014) Fluorescence microscopy. Cold Spring Harb Protoc 2014(10):pdb top071795. https://doi.org/10.1101/pdb.top071795 21. Kurihara D, Kimata Y, Higashiyama T, Ueda M (2017) In Vitro Ovule Cultivation for Live-cell Imaging of Zygote Polarization and Embryo P https://www.jove.com/video/55975 https:// doi.org/10.3791/55975

Part V Embryogenesis in Other Model and Non-model Species

Chapter 14 Imaging of Embryo Sac and Early Seed Development in Maize after Feulgen Staining Kamila Kalinowska, Junyi Chen, and Thomas Dresselhaus Abstract Compared with small model plants like Arabidopsis containing ovules with few cell layers, embryo sac and embryo development of model crop plants such as maize and other grasses are difficult to image. Multiple layers of tissue usually surround the deeply embedded embryo sac and developing embryo. Moreover, reliable cell biological marker lines labeling, for example, nuclei, plasma membrane, cell walls, or cells of a specific identity are often not available. The introduction of markers to study mutants is difficult and timeconsuming and may require several generations of backcrosses. In this chapter, we therefore present an easy protocol to image maize ovaries and developing embryo sacs before and after fertilization allowing also high-throughput mutant analysis. The laborious embedding of samples and preparation of thin sections are omitted in this fixing-Feulgen staining-clearing (FFC) method. Optical sectioning through multiple layers of tissue is possible allowing 3D reconstructions of the whole embryo sac if necessary. The advantage of staining cell nuclei using the FFC method described here compared, for example, with DAPI staining is a wide range of Schiff’s type reagents available for the Feulgen reaction. Depending on the reagent of choice, various conditions such as different excitation/emission filters or even white light can be applied for imaging. Moreover, in order to better visualize cell division, nuclei polarity as well as cell extent and integrity, periodic acid staining (PAS) of cell walls can be combined with Feulgen staining. Key words Ovary, Embryo sac, Fertilization, Embryogenesis, Maize, Zea mays, Acriflavine, Schiff’s reagent, Feulgen staining

1

Introduction Embryo sac and embryo development are difficult to visualize in plant species containing multiple surrounding ovular cell layers consisting of hundreds and thousands of cells, respectively. This is the case for maize (Zea mays) and many other plant species especially those containing large ovaries and ovules, a desired yield trait, which is directly correlated to fruit and seed size, respectively. However, this makes microscopic studies about the detailed morphology and mutant analyses difficult. In maize, a relatively large number of cell biological markers have been developed in the past 10 years [1], but their introduction into mutants defective in

Martin Bayer (ed.), Plant Embryogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2122, https://doi.org/10.1007/978-1-0716-0342-0_14, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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embryo sac and/or embryo development would require a number of crossings and backcrossings to establish homozygous lines. For other agronomically important plant species, efficient transformation protocols are even more problematic and cell biological markers are not yet available. In this chapter, we therefore describe an easy method to fix and clear (FC) maize ovaries and developing cobs, which can likely also be applied in other plant species. The FC method can be extended as FFC method by Feulgen staining (fixing-Feulgen staining-clearing) to visualize cell nuclei and by periodic acid staining (PAS) to dye cell walls. The FC and FFC methods omit the laborious and time-consuming embedding of samples as well as preparation of thin sections using a vibratome and microtome, respectively. In addition, in contrast to traditional histological preparations, optical sectioning of ovaries allows 3D reconstruction of the whole embryo sac if desired. Cell walls and nuclei can be visualized using PAS and Feulgen staining, which is important to study cellular details of embryo sac and embryo (seed) development, respectively [2– 5]. We recommend adding the two dyes, as it allows to study mutants showing abnormalities in female gametophyte development, during fertilization and/or early embryogenesis associated, for example, with defects in nuclei positioning, mitosis/meiosis, cell polarity and/or division as well as cell size [6–9]. A simplified workflow of the FFC method is shown in Fig. 1. Feulgen staining is based on Schiff’s reaction, which is traditionally used in detection of organic aldehydes. Feulgen staining is also widely applied in histology to identify and visualize chromosomal material or DNA in cell specimens. Staining involves depurination of DNA (hydrolytic removal of purine bases such as adenine or guanine) through treatment with a concentrated acid and subsequent reaction of an aldehyde bond formed in this way with a Schiff’s type reagent [10–12]. The resulting product can be detected colorimetrically or fluorescently, depending on the kind of Schiff’s reagent used for staining. This is also an advantage of this method compared, for example, with 40 ,6-diamidine-2-phenylindole staining (DAPI), which depends on UV light or -lasers. The method described here involves dissection of ovaries, fixing in FAA solution (formalin–ethanol–acetic acid) and storage in 70% (v/v) ethanol. If staining of nuclei using Feulgen reagents is needed, samples are rehydrated by an ethanol series, DNA hydrolysis conducted through treatment with a concentrated hydrochloric acid, and followed by staining with a Schiff’s reagent. Samples are then dehydrated by an ethanol series and cleared through treatment with methyl salicylate–ethanol solutions and pure methyl salicylate. Finally, samples are analyzed microscopically mounted in methyl salicylate. It is possible to perform only FC without Feulgen staining of maize embryo sacs, which has also been used, for example, during the analysis of maize embryo sac mutants [13]. For

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Fig. 1 Simplified workflow of the FFC method (fixing-Feulgen staining-clearing method) of maize ovaries. The different steps of the procedure are indicated as well as the time that should be calculated. Maize ovaries can be sectioned either longitudinally or diagonally

comparison, the different steps concerning FC and FFC method, respectively, are marked in the protocol and in Table 1. Two strategies are used in our group: the first strategy involves putting whole maize ears or large parts of ears into fixing solution and sectioning ovules after completion of fixing, staining and clearing. The second method, which we recommend for stages a few days after fertilization, starts with sectioning and then proceeds with the protocol.

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Table 1 Summary of steps required for fixing, facultative Feulgen staining, and clearing of maize ovaries and developing seeds

FFC-method FC-method

Solution

Whole maize ears

Dissected maize ovaries

Dissection

Dissection

As in Fig. 1

Fixing

Fixing

FAA-1 [10% (v/v) formalin, 50% (v/v) ethanol (for analysis), 5% (v/v) glacial acetic acid] or FAA-2 [3.7% (v/v) formalin, 50% (v/v) ethanol (for analysis), 5% (v/v) glacial acetic acid]

FAA-1, 15 min weak vacuum, then FAA-1, 16–24 h

FAA-2, 10 min weak vacuum, 16 h to 1 month

Storage

Storage

70% (v/v) ethanol

At least 16 h

At least 16 h

Rehydration

50% (v/v) ethanol 30% (v/v) ethanol 15% (v/v) ethanol ddH2O

30 min 30 min 30 min 2 30 min

30 min 30 min 30 min 2 30 min

Feulgen staining

6 M HCl Water

25 min 25 min Thorough rinsing 4 ddH2O with copious distilled water for 1 h, then 2 ddH2O 20 min 20 min

0.5% (w/v) periodic acid solution (PAS) Water

0.1% (w/v) acriflavine Water

Thorough rinsing 4 ddH2O with copious distilled water for 1 h, then 2 ddH2O 20 min 20 min Thorough rinsing 4ddH2O with copious distilled water for 1 h, then 2 ddH2O

Dehydration

15% (v/v) ethanol 30% (v/v) ethanol 50% (v/v) ethanol 70% (v/v) ethanol Dehydration 85% (v/v) ethanol 95% (v/v) ethanol Pure ethanol

30 min 30 min 30 min 30 min to 16 h 30 min 30 min 2 30 min

30 min 30 min 30 min 30 min to 16 h 30 min 30 min 2 30 min

Clearing

Clearing

1–16 h 1–16 h 1–16 h 2 1–16 h

1–16 h 1–16 h 1–16 h 2 1–16 h

Dissection

Dissection

1:3 (v/v) methyl salicylate–ethanol 1:1 (v/v) methyl salicylate–ethanol 3:1 (v/v) methyl salicylate–ethanol Pure methyl salicylate

As in Fig. 1

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Materials

2.1 Fixing and Storage of Whole Maize Ears

1. Fixing solution (FAA-1) for whole maize ears: 10% (v/v) formalin, 50% (v/v) ethanol (for analysis), 5% (v/v) glacial acetic acid (see Note 1). Store at 4  C. 2. 70% (v/v) ethanol (for analysis). 3. Vacuum chamber (see Fig. 2) and a pump.

Fig. 2 Materials and devices necessary for FC/FFC methods to analyze maize ovaries. (a) Whole maize ears were placed into 50-ml tubes inside a glass vacuum chamber, covered with a lid and connected to a pump. (b) If maize ovaries were dissected before fixing, 2 ml tubes containing dissected ovaries are placed in a vacuum chamber. (c) Microscope slides suitable for analysis of dissected maize ovaries. On the left: commercially available concave slides; in the middle: slides cut out from thin aluminum plate. The bottom coverslip was attached using hot wax; on the right side: slides generated using a 3D printer. The bottom coverslip was attached using double-sided adhesive tape

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2.2 Fixing and Storage of Dissected Maize Ovaries

1. Fixing solution (FAA-2) for dissected maize ovaries: 3.7% (v/v) formalin, 50% (v/v) ethanol (for analysis), 5% (v/v) glacial acetic acid (see Note 2). Store at 4  C. 2. 70% (v/v) ethanol (for analysis). 3. Vacuum chamber and a pump.

2.3

Rehydration

1. Ethanol (for analysis) dilutions in double-distilled water: 50% (v/v), 30% (v/v), and 15% (v/v). 2. Double-distilled water. 3. Horizontal shaker.

2.4

Feulgen Staining

1. 6 M hydrochloric acid: Mix 50 ml fuming (12 M) hydrochloric acid with 50 ml double-distilled water. Store at room temperature. 2. 0.5% (w/v) periodic acid solution (PAS) (see Note 3): Weigh 0.5 g periodic acid and dissolve in 100 ml double-distilled water. Store at room temperature and protect from light (using a brown bottle or covering a bottle with aluminum foil). Alternatively, a commercially available ready 0.5% (w/v) PAS can be used. 3. Schiff’s type reagent, for example 0.1% (w/v) acriflavine (see Note 4): Weigh 0.1 g acriflavine and dissolve in 100 ml doubledistilled water. Store at 4  C and protect from light.

2.5

Dehydration

1. Absolute ethanol (for analysis). 2. Ethanol (for analysis) dilutions in double-distilled water: 15% (v/v), 30% (v/v), 50% (v/v), 70% (v/v), 85% (v/v), and 95% (v/v). Store at room temperature.

2.6

Clearing

1. Methyl salicylate (synthetic) (see Note 5). Store at room temperature and protect from light. 2. Methyl salicylate–ethanol solutions, 1:3, 1:1, and 3:1: Measure 25, 50, and 75 ml of methyl salicylate, respectively, and fill with absolute ethanol for analysis up to 100 ml. Store at room temperature and protect from light.

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Methods

3.1 Fixing and Storage of Whole Maize Ears

1. Grow maize at standard greenhouse conditions. Maize female flowers are formed about 8–12 weeks after germination, depending on the genotype. In order to control pollination, cover ears with a paper bag before silk emergence. When silks emerge from husk leaves of the ear to a length of 3–4 cm, pollinate them with fresh pollen and harvest ears at stages of

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interest. If immature ovaries will be studied, ears have to be collected earlier. We advise to correlate developmental stages with ear size, which is genotype dependent. 2. Collect whole maize cobs and remove husk leaves. Place the whole ear in a 50 ml plastic tube (e.g., Falcon™ tube) and add FAA-1 solution till the ear is covered. If the whole ear does not fit into the 50 ml tube, you may cut stripes from the ear containing 3–4 rows of ovaries and place them into the tube. 3. Place open 50 ml tubes containing whole maize ears or ear stripes in a glass vacuum chamber as shown in Fig. 2a. Close the chamber. 4. Attach the vacuum chamber with a rubber or plastic tube to a pump. Switch the pump on. Open a cork to the glass chamber. Generate a vacuum for 15 min. After 15 min close the cork and switch the pump off. 5. Discard old FAA-1 solution and pour fresh FAA-1 solution into the tube, till the whole ear is immersed. 6. Place tubes to 4  C overnight. Do not store them for longer than 24 h in fixing solution (see Note 6). 7. Wash ears twice with 70% (v/v) ethanol: add 70% (v/v) ethanol till the ear is covered, close the tube, mix by inverting the tube, and discard the ethanol. Repeat this step. 8. Add 70% (v/v) ethanol till the ear is covered and place tubes to 4  C for at least overnight. Samples can be stored in 70% (v/v) ethanol for up to several months before proceeding with further steps. Continue with Subheading 3.3 for the full protocol. If only fixing and clearing without staining shall be performed, immediately continue with Subheading 3.5, step 5. 3.2 Fixing and Storage of Dissected Maize Ovaries

1. Grow maize in standard greenhouse conditions as described in Subheading 3.1, step 1. 2. Dissect maize ovaries (longitudinal or diagonal sections) using a sharp razor blade as shown in Fig. 1. For high resolution images, generate about 0.5–0.8 mm thick sections. For a quick observation, sections of 1.5 mm and thicker can be used (see Note 7). 3. Place dissected maize ovaries in 2 ml reaction tubes filled with 2 ml FAA-2. 4. Place samples in a glass vacuum chamber and treat them for 10 min with gentle vacuum as described in Subheading 3.1, step 4 (Fig. 2b). Store samples at 4  C overnight (see Note 8). 5. Wash samples twice with 2 ml 70% (v/v) ethanol.

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6. Add 2 ml 70% (v/v) ethanol and place samples to 4  C for at least overnight. Samples can be stored in 70% (v/v) ethanol for up to several months before proceeding with further steps. Continue with Subheading 3.3 for the full protocol. If only fixing and clearing without staining shall be performed, you can immediately continue with Subheading 3.5, step 5. 3.3

Rehydration

1. Discard old solution and add 50% (v/v) ethanol to cover samples. Incubate for 30 min at room temperature. For whole maize ears, gentle agitation (50 rpm, on a horizontal shaker) is recommended. 2. Discard the solution and add 30% (v/v) ethanol to cover samples. Incubate for 30 min at room temperature. For whole maize ears, gentle agitation is recommended. 3. Discard the solution and add 15% (v/v) ethanol to cover samples. Incubate for 30 min at room temperature. For whole maize ears, gentle agitation is recommended. 4. Discard the solution and add double-distilled water to cover samples. Incubate for 30 min at room temperature. For whole maize ears, gentle agitation is recommended. Repeat this step.

3.4

Feulgen Staining

1. Discard the old solution. For dissected maize ovaries, add 2 ml of 6 M HCl. For whole maize ears, add 6 M HCl till the ear is completely covered. Incubate samples for 20 min at room temperature with gentle agitation. 2. Discard the solution. For dissected maize ovaries, wash samples four times with 2 ml double-distilled water. For whole maize ears, rinse ears thoroughly in copious distilled water for at least 1 h, replace water at least twice. Samples can be incubated in water overnight. Afterwards, wash samples twice with doubledistilled water. 3. Discard water. For dissected maize ovaries, add 2 ml of 0.5% (w/v) periodic acid. For whole maize ears, add 0.5% (w/v) periodic acid till the ear is completely covered. Incubate samples for 20 min at room temperature with gentle agitation. 4. Discard the old solution. For dissected maize ovaries, wash samples four times with 2 ml double-distilled water. For whole maize ears, rinse ears thoroughly in copious distilled water for at least 1 h, replace water at least twice. Samples can be incubated in water overnight. Afterwards, wash samples twice with double-distilled water. 5. Discard water. For dissected maize ovaries, add 2 ml of 0.1% (w/v) acriflavine. For whole maize ears, add 0.1% (w/v) acriflavine till the ear is completely covered. As acriflavine is light sensitive, protect tubes with aluminum foil. Incubate samples for 20 min at room temperature with gentle agitation.

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6. Discard the solution. For dissected maize ovaries, wash samples four times with 2 ml double-distilled water. For whole maize ears, rinse ears thoroughly in copious distilled water for at least 1 h, replace water at least twice. Samples can be incubated in water overnight. Afterward, wash samples twice with doubledistilled water. 3.5

Dehydration

1. Discard the solution and add 15% (v/v) ethanol to cover samples. Incubate for 30 min at room temperature. 2. Discard the solution and add 30% (v/v) ethanol to cover samples. Incubate for 30 min at room temperature. 3. Discard the solution and add 50% (v/v) ethanol to cover samples. Incubate for 30 min at room temperature. 4. Discard the solution and add 70% (v/v) ethanol to cover samples. Incubate for at least 30 min at room temperature. This step can be prolonged up to overnight to leach out excess of acriflavine. 5. Discard the solution and add 85% (v/v) ethanol to cover samples. Incubate for 30 min at room temperature. 6. Discard the solution and add 95% (v/v) ethanol to cover samples. Incubate for 30 min at room temperature. 7. Discard the solution and add absolute ethanol to cover samples. Incubate for 30 min at room temperature. Repeat this step.

3.6

Clearing

1. Discard the old solution. For dissected maize ovaries, add 2 ml of 1:3 methyl salicylate–ethanol solution. For whole maize ears, add 1:3 methyl salicylate–ethanol solution till the ear is completely covered. Incubate for at least 1 h at room temperature with gentle agitation. 2. Discard the solution. For dissected maize ovaries, add 2 ml of 1:1 methyl salicylate–ethanol solution. For whole maize ears, add 1:1 methyl salicylate–ethanol solution till the ear is completely covered. Incubate for at least 1 h at room temperature with gentle agitation. This step can be prolonged up to overnight to remove excess of acriflavine (see Note 9). 3. Discard the solution. For dissected maize ovaries, add 2 ml of 3:1 methyl salicylate–ethanol solution. For whole maize ears, add 3:1 methyl salicylate–ethanol solution till the ear is completely covered. Incubate for at least 1 h at room temperature with gentle agitation. 4. Discard the solution. For dissected maize ovaries, add 2 ml of pure methyl salicylate solution. For whole maize ears, add pure methyl salicylate till the ear is completely covered. Incubate for

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at least 1 h at room temperature with gentle agitation. Repeat this step. For a better overview, all steps of the protocol are collected as in Fig. 1 and Table 1. 3.7

Microscopy

1. For whole maize ears, dissect ovaries as shown in Fig. 1 and described in Subheading 3.2, step 2. For dissected maize ovaries, no further sample preparation is required. 2. Prepare concave microscope slides (Fig. 2c, see also Note 10). Place dissected ovaries in the concavity using forceps and mount samples in methyl salicylate (see Note 11). Place a coverslip over samples. 3. Use a confocal laser scanning microscope to analyze the samples. For acriflavine use a GFP (488 nm) filter to obtain pictures. Examples of application of the above described method to visualize different stages of ovule and seed development in maize are presented in Fig. 3.

4

Notes 1. As an alternative to FAA-1, Chemberlin’s fix [5% (v/v) formalin, 45% (v/v) ethanol, and 5% (v/v) glacial acetic acid] can be used [14]. 2. It is also possible to fix samples with FAA-1 or Chemberlin’s fix [5% (v/v) formalin, 45% (v/v) ethanol, and 5% (v/v) glacial acetic acid]. Take care that sample are not left in fixing solution for longer than 24 h. 3. 0.5% (w/v) periodic acid (PAS) increases unspecific staining of cell walls, which allows a better visualization of single cells and can be advantageous especially for the analysis of cell division mutants. For more specific staining of cell nuclei, staining with PAS should be excluded from the protocol. 4. It is possible to use other Schiff’s type reagents instead of acriflavine. Popular alternatives are, for example, commercial Schiff’s reagents or other DNA-staining reagents including toluidine blue. 5. A number of alternative clearing reagents is available. Popular among plant researchers are xylene, Hoyer’s clearing/mounting solution and Histo-Clear (National Diagnostics). Though methyl salicylate is widely used for maize, it might not be optimal for other monocot plants and needs to be tested for each species. 6. Samples should never be kept for longer than 24 h in fixing solution FAA-1 as tissues could harden and damage.

Feulgen Staining of Maize Embryo Sacs

A

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II II

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C *

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NU

dNU

NU

APs

EN

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D

E

F

BETL ESR

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ESR

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EMB PE

EN

Fig. 3 Application of the FFC method to visualize different ovule and seed developmental stages of maize. (a) Developing ovule during meiosis. After Meiosis I, the upper cells (open arrowhead) arrests and disintegrates, while the second cell (closed arrowhead) undergoes a second meiotic division. (b) Mature ovule showing a female gametophyte at stage FG7. (c) Two days after pollination (DAP), the small apical cell is dividing. The uncellularized endosperm contains a number of peripheral nuclei (open arrowheads) separated by a large central vacuole. (d) 4 DAP endosperm starts to cellularize. The embryo consists of small apical and larger vacuolated suspensor cells. (e) 6 DAP showing cellularized endosperm, a transition stage embryo and degenerating nucellus tissue. The area shown in the inset is enlarged in (f). AC apical cell, APs antipodal cells, BC basal cell, BETL basal endosperm transfer layer, CC central cell, dNU degenerating nucellus, EMB embryo, EN endosperm, ESR embryo surrounding region, II inner integument, NU nucellus, OI outer integument, OV ovary tissue, PE pericarp. Scale bars are 100 μm

7. Correct sectioning of ovaries/developing seeds is crucial for obtaining high quality pictures of embryo sacs and embryos. Always use fresh, sharp razor blades and keep sections as thin as possible. This will probably require some training till satisfactory results will be reached. Both longitudinal and diagonal sectioning methods of maize ovaries are possible. Longitudinal sections are the first method of choice and are advantageous as whole embryo sacs can be observed. We usually use diagonal sections for manual isolation of embryo sac cells and early embryos. 8. We had a good experience to store maize ovaries in FAA-2 for up to 1 month; however, we recommend storage in 70% (v/v) ethanol.

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9. During clearing in methyl salicylate–ethanol, it is possible to prolong steps to 5 h or even overnight (one to two steps per day). Prolongation of the clearing procedure will help to achieve a better clearing of deeper cell layers and removal of residues of unbound staining solution. If the clearing procedure is too long, it usually weakens nuclei staining depending on the reagent used. 10. Microscope slides with 1–3 concavities can be ordered from distributors such as Thermo Fisher Scientific or SigmaAldrich/Merck (Fig. 2c). A slide with a hole inside can also be cut out from a thin aluminum or synthetic material plate. Alternatively, we recommend generating it using a 3D printer (Fig. 2c). A coverslip can be attached with wax, glue, or a two-sided tape on one side of the slide. Samples should be placed on the other side of the slide in the concave, mounted with methyl salicylate and covered with a coverslip. 11. We always use methyl salicylate to mount our samples during microscopic analysis. This method has, however, two major disadvantages: if (1) commercial concave microscope slides are used and sections are relatively thick, samples cannot be inverted and analyzed using an inverted microscope due to possible leakage of the mounting solution; and (2) methyl salicylate has a strong smell. There are several alternatives, for example, mixtures of ethanol and oils such as immersion oil or Canadian oil.

Acknowledgments We acknowledge Na´dia Graciele Krohn, who initially established the FFC method, and further lab members for optimizing the method and sharing their experience. We would like to thank our colleagues Mihaela-Luiza Ma´rton for advice on clearing procedures and Ivan Kulich for assistance with the 3D printer. We thank Armin Hildebrand and Noureddine Djella for plant care. This research was funded by the German Federal Ministry of Education and Research (BMBF), Plant2030 grant 031B0192 to T.D. References 1. Maize CellGenomics Database (2018) J. Craig Venter Institute. http://maize.jcvi.org/ cellgenomics. Accessed 16 Dec 2018 2. Barrell PJ, Grossniklaus U (2005) Confocal microscopy of whole ovules for analysis of reproductive development: the elongate1 mutant affects meiosis II. Plant J 43:309–320

3. Chettoor AM, Evans MM (2015) Correlation between a loss of auxin signaling and a loss of proliferation in maize antipodal cells. Front Plant Sci 6:187 4. Wu CC, Diggle PK, Friedman WE (2011) Female gametophyte development and double fertilization in Balsas teosinte, Zea mays subsp.

Feulgen Staining of Maize Embryo Sacs parviglumis (Poaceae). Sex Plant Reprod 24:219–229 5. Guo F, Huang BQ, Han Y, Zee SY (2004) Fertilization in maize indeterminate gametophyte1 mutant. Protoplasma 223:111–120 6. Chen J, Kalinowska K, Mu¨ller B, Mergner J, Deutzmann R, Schwechheimer C, Hammes UZ, Dresselhaus T (2018) DiSUMO-lIKE interacts with RNA-binding proteins and affects cell-cycle progression during maize embryogenesis. Curr Biol 28:1548–1560 7. Juranic M, Srilunchang KO, Krohn NG, Leljak-Levanic D, Sprunck S, Dresselhaus T (2012) Germline-specific MATH-BTB substrate adaptor MAB1 regulates spindle length and nuclei identity in maize. Plant Cell 24:4974–4991 8. Krohn NG, Lausser A, Juranic M, Dresselhaus T (2012) Egg cell signaling by the secreted peptide ZmEAL1 controls antipodal cell fate. Dev Cell 23:219–225 9. Srilunchang KO, Krohn NG, Dresselhaus T (2010) DiSUMO-like DSUL is required for

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nuclei positioning, cell specification and viability during female gametophyte maturation in maize. Development 137:333–345 10. Kasten FH, Burton V, Lofland S (1962) Schifftype reagents in cytochemistry. 2. Detection of primary amine dye impurities in pyronin B and pyronin Y(G). Stain Technol 37:277–291 11. Kasten FH (1959) Schiff-type reagents in cytochemistry. I. Theoretical and practical considerations. Z Zellforch Microsk Anat Histochem 1:466–509 12. Kasten FH (1958) Additional Schiff-type reagents for use in cytochemistry. Stain Technol 33:39–45 13. Gutierrez-Marcos JF, Costa LM, Evans MM (2006) Maternal gametophytic baseless1 is required for development of the central cell and early endosperm patterning in maize (Zea mays). Genetics 174:317–329 14. Vollbrecht E, Hake S (1995) Deficiency analysis of female gametogenesis in maize. Dev Genet 16:44–63

Chapter 15 Using Giant Scarlet Runner Bean (Phaseolus coccineus) Embryos to Dissect the Early Events in Plant Embryogenesis Min Chen, Anhthu Q. Bui, and Robert B. Goldberg Abstract The giant embryo of the scarlet runner bean (Phaseolus coccineus) has been used historically to investigate the molecular and developmental processes that control the early events of plant embryo development. In more recent years, our laboratory has been using scarlet runner bean embryos to uncover the genes and regulatory events that control embryo proper and suspensor region differentiation shortly after fertilization. In this chapter we describe methods that we have developed to isolate scarlet runner bean embryos at the globular stage of development, and capture embryo proper and suspensor regions by either hand dissection or laser capture microdissection (LCM) for use in downstream genomic analysis. These methods are also applicable for use in investigating the early events of common bean (Phaseolus vulgaris) embryo development, a close relative of scarlet runner bean, which also has a giant embryo in addition to a sequenced genome. Key words Scarlet runner bean, Pollination, Seed development, Embryo, Globular stage, Embryo proper, Suspensor, Embryo isolation, Microscopy, Laser capture microdissection

1

Introduction One of the major challenges in plant biology is to uncover the mechanisms that program embryo differentiation into specialized regions, tissues, and cell types shortly after fertilization. This is a difficult task because the earliest events in embryo development occur deep within the flower and are difficult to access and investigate using molecular and genomic techniques. In recent years, the development of novel methods such as laser capture microdissection (LCM) [1] and isolation of nuclei tagged in specific cell types (INTACT) [2] have overcome some of this difficulty by making it possible to capture either cells or nuclei from postfertilization embryo proper and suspensor regions in order to carry out highthroughput transcriptome analysis. Each of these methods, however, has their strengths and limitations. For example, LCM requires expensive, highly specialized microscopy equipment, and

Martin Bayer (ed.), Plant Embryogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2122, https://doi.org/10.1007/978-1-0716-0342-0_15, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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a significant amount of time and effort is required to capture adequate amounts of embryonic regions for genomic studies— especially from tiny Arabidopsis embryos. In addition, methods for isolating chromatin from paraffin-embedded tissues on slides are just beginning to be developed [3]. On the other hand, LCM can be applied universally to all plants regardless of whether the plant can be transformed or not. INTACT, by contrast, requires generating transgenic plants with chimeric cell-specific genes but permits the capture of nuclei from a variety of embryo regions and cell types for genomic studies—as long as cell-and region-specific promoters are available. Thus, it cannot be used with plants that do not have established transformation protocols. Nevertheless, applying both LCM and INTACT techniques to the investigation of postfertilization plant embryos has uncovered novel insights into the genes and processes controlling early plant embryo development [2, 4–6]. The scarlet runner bean (Phaseolus coccineus) (Fig. 1) provides a novel opportunity for dissecting the molecular processes controlling plant embryo development. At the globular stage, the scarlet runner bean embryo is ~100 times larger than that of Arabidopsis, contains a suspensor with 200 cells that is highly polyploid, and can be isolated directly from developing seeds within the flower [7–11]. Because of its large embryonic size, both embryo proper and suspensor regions can be separated from each other manually and used directly for biochemical and molecular studies [11]. Almost 50 years ago, the late Ian Sussex and his collaborators pioneered the use of giant scarlet runner bean embryos and provided the first insights into the molecular processes controlling early embryogenesis; for example, the suspensor produces signals that are required for embryo proper development [7, 8, 12–15]. During this same period, others demonstrated that hormones, such as gibberellic acid, are synthesized within the giant scarlet runner bean suspensor and contribute to embryo proper formation [16]. The ability to manually isolate large numbers of

Fig. 1 Scarlet runner bean plant, flower cluster, open flower, 5-DAP (days after pollination) pod, globular stage seed, and embryo

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globular stage embryo proper and suspensor regions, in addition to using state-of-the-art LCM techniques, provides a unique opportunity to use the scarlet runner bean to gain entry into the earliest events in plant embryogenesis—complementing the elegant studies that can be carried out with Arabidopsis [2, 4–6] and maize [17]. Our laboratory has resurrected the use of scarlet runner bean for the study of early plant embryo development. We have sequenced thousands of expressed sequence tags (ESTs) from embryo proper and suspensor regions (GenBank Accessions CA896559-916678; GD289845-GD660862) [11], used in situ hybridization [11, 18, 19] and RNA-Seq (GenBank Accession GSE57537) to identify embryo proper- and suspensor-specific mRNAs, uncovered a suspensor cis-regulatory module that activates region-specific transcription of genes within the suspensor shortly after fertilization [19–21], and generated a rough draft of the scarlet runner bean genome (GenBank Accession QBDZ00000000). In this chapter, we outline the methods that we have developed to use giant scarlet runner bean embryos to identify processes controlling the differentiation and functions of the embryo proper and suspensor regions at the globular stage of development. We specifically focus on methods used to grow scarlet runner bean plants, harvest embryos, and manually dissect embryo proper and suspensor regions for direct use in genomic studies, as well as techniques required to fix and embed young seed tissues for LCM and in situ hybridization experiments.

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Materials

2.1 Planting Scarlet Runner Bean Seeds

1. Scarlet runner bean cultivar “Hammond’s Dwarf Red Flower” (Vermont Bean Seed Company). 2. Soil mix (equal parts of ground Canadian sphagnum, horticultural grade sand, white pumice, and redwood compost). 3. Water-soluble fertilizer.

20-20-20

Nitrogen-Phosphorus-Potassium

4. 1-gallon pot. 2.2 Hand-Pollinating Scarlet Runner Bean Flowers

1. Fine-tipped watercolor brush.

2.3 Collecting Globular Stage Scarlet Runner Bean Seeds

1. Dissecting microscope.

2. White merchandise tag with white strings (7/8 in.
1¼ in.). 3. Permanent marker.

2. Aluminum foil. 3. Ice bucket. 4. Ruler.

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5. Plain glass microscope slide. 6. Disposable scalpel with No. 11 blade. 2.4 Dissecting Whole-Mount Globular Stage Scarlet Runner Bean Embryos

1. Dissecting microscope. 2. Concavity slide with depressed well (0.6–0.8 mm deep). 3. Concavity slide with depressed well (1.5 mm deep). 4. Plain glass microscope slide. 5. Disposable scalpel with No. 11 blade. 6. 1 ml syringe with BD PrecisionGlide needle (No. 30.5 gauge). 7. 1.5 ml microcentrifuge tube (Nuclease-free). 8. Tie wire. 9. Dewar flask. 10. Permanent marker. 11. Liquid nitrogen. 12. Micropipette (1 ml). 13. 1 ml pipette tips (Nuclease-free). 14. Fine-tipped forceps. 15. Nuclease-free water.

2.5 Fixing Globular Stage Scarlet Runner Bean Seeds for LCM or in situ Hybridization

1. Ethanol/acetic acid fixative solution for LCM [(ethanol/acetic acid, 3:1 (v/v)]: 15 ml 100% ethanol; 5 ml acetic acid. Prepare freshly and keep on ice. 2. 1% glutaraldehyde fixative solution for in situ hybridization (1% glutaraldehyde, 0.1 M sodium phosphate buffer, 0.1% Triton X-100): 0.8 ml 25% glutaraldehyde solution; 0.2 ml 100% Triton X-100; 10 ml 0.2 M sodium phosphate buffer, pH 7.0; add nuclease-free water to 20 ml. Prepare freshly and keep on ice. 3. Scintillation glass vial. 4. Dissecting microscope. 5. Plain glass microscope slide. 6. Disposable scalpel with No. 11 blade. 7. Forceps. 8. Vacuum oven. 9. Rotator.

2.6 Dehydration for LCM

1. A series of ethanol concentrations in nuclease-free water for dehydration: 75%, 85%, 95%, and 100%. 2. Rotator.

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1. 0.1 M sodium phosphate buffer, pH 7.0. 2. A series of ethanol concentrations in nuclease-free water for dehydration: 5%, 10%, 15%, 20%, 25%, 30%, 40%, 50%, 60%, 70%, 80%, 90%, and 100%. 3. Rotator.

2.8 Infiltration and Embedding for LCM or in situ Hybridization

1. Xylene. 2. 100% ethanol. 3. Rotator. 4. Paraplast X-tra Tissue Embedding Medium (VWR, cat. no. 15159-486). 5. 1-l beaker. 6. Incubator (42  C and 58  C). 7. Disposable aluminum crinkle dishes with tabs (e.g., VWR, cat. no. 25433-008). 8. Hot plate with divided hot/cold sections. 9. Spatula. 10. Alcohol lamp.

2.9 Sectioning for LCM or in situ Hybridization

1. Microtome. 2. Standard razor blade. 3. Plastic block holder. 4. Aluminum foil. 5. Slide warmer. 6. Fine-tipped watercolor brushes. 7. Nuclease-free water. 8. Micropipettes (1 ml and 100 μl). 9. 1 ml and 100 μl filtered pipette tips (Nuclease-free). 10. Ice bucket. 11. Polyethylene naphthalate (PEN) membrane slides for LCM (Leica, cat. no. 11505158 for DNA work or no. 11505189 for RNA work). 12. Superfrost Plus Slides for in situ hybridization.

2.10

LCM

1. Leica LMD 6000 system or similar LCM microscope system. 2. 0.5 ml microcentrifuge tubes (Nuclease-free). 3. Extraction buffer: Buffer XB from PicoPure RNA isolation kit (Thermo Fisher, cat no. KIT0204) can be used for RNA isolation, and Buffer ATL from QIAamp DNA FFPE Tissue Kit (QIAGEN, cat. no. 56404) can be used for DNA isolation.

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Methods

3.1 Planting Scarlet Runner Bean Seeds

1. Sterilize the soil mix by autoclaving at 121  C for 90 min. Let the soil cool before planting seeds. 2. Wash pots with soapy water, rinse, and dry fully. 3. Fill a 1-gallon pot with sterilized soil mix up to 2 in. from the top, and water the soil mix completely. 4. Spread 10 scarlet runner bean seeds evenly on the wet soil mix and lightly press the seeds into the soil. 5. Cover the seeds with a thin layer (~ 1 in.) sterilized soil mix. 6. Place pots with scarlet runner bean seeds in a greenhouse, or growth chamber, with 16 h-light–8 h-dark cycle at 22  C (see Note 1). 7. Transfer seedlings to a second 1-gallon pot 10 days after sowing (two seedlings per pot). 8. Water as needed with water containing 250 ppm of watersoluble 20-20-20 Nitrogen-Phosphorus-Potassium fertilizer.

3.2 Hand-Pollination of Scarlet Runner Bean Flowers

1. Take a flower that opened in the morning, such as flowers #3 and #4 (Fig. 2). Push the standard and wing petals (Fig. 2) away from the coiled keel that contains the style, stigma, and anthers to expose dehiscent anthers (see Note 2). 2. Collect pollen by gently brushing against the anthers using a fine-tipped watercolor brush and make sure the pollen grains, like greyish yellow powder, are visible on the brush tip.

Fig. 2 Scarlet runner bean flowers. All flowers were collected at the same time. In flower #1, the pollen has not been released and the stigma is dull. In flower #2, the anther is dehiscing and the stigma is shiny and sticky. In flower #3, the anther is fully dehisced and stigma is shiny and sticky. In flower #4, the pollen is completely released, but the stigma is senescing and withered. In addition, the wing petal has started to wither. In flower #5, the petal is withered and the pollen is not viable. Flowers #3 and #4 are good for collecting pollen. Flowers #2 and #3 are excellent for pollination

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3. Take a second flower that opened in the morning, such as flowers #2 and #3 (Fig. 2). Push the standard and the wing petals away from the coiled keel until the stigma is exposed. 4. Touch the pollen-covered brush to receptive stigma. Pollen grains will be visible sticking to the stigma if the pollen was successfully transferred. 5. Label the pollinated flower with a white merchandise tag attached to a string. 6. Repeat steps 1–5 and pollinate more flowers (see Note 3). 3.3 Collecting Scarlet Runner Bean Seeds Containing Embryos at the Globular Stage

1. Collect pods 5–7 days after pollination (DAP) and wrap pods in aluminum foil. Keep collected pods on ice at all times (see Note 4). The stage of the embryo development within the expanding pods depends upon both the pollination effectiveness and time of the year. Therefore, pod length and days after pollination provide only a rough estimate of embryonic stage. Precise staging should be done just before the real experiment starts by hand-dissecting embryos from seeds within young expanding pods and examining under the microscope to calibrate the timing of embryo development. 2. Under the dissecting microscope, use the blade to slice open the pod, remove the seeds, and place seeds on a plain microscope slide (Fig. 1). 3. Measure the seed length (Fig. 1). The length of scarlet runner bean seeds containing globular stage embryos is ~2.0–2.5 mm [18]. 4. If an entire whole-mount embryo, or a specific embryonic region (e.g., embryo proper or suspensor), is needed for the experiment [11], follow steps in Subheading 3.4 immediately to isolate and dissect whole-mount embryos. If LCM [1, 4], or in situ hybridization [18, 19, 22], is planned, immediately follow steps in Subheading 3.5 to fix the seeds and perform the downstream procedures.

3.4 Dissecting Scarlet Runner Bean Globular Stage Whole-Mount Embryos

1. Pour liquid nitrogen into a Dewar flask before opening the pods. 2. Label a 1.5 ml microcentrifuge tube and loop around with the tie wire. Shape the extra piece of tie wire into a hook and hang the 1.5 ml tube on the edge of the Dewar flask. It is important that the 1.5 ml tube is suspended in the liquid nitrogen. 3. Fill the 1.5 ml tube with liquid nitrogen. 4. On a plain microscope slide, using the disposable scalpel with a No. 11 blade, cut the seed in half transversely (Fig. 1). Place the micropylar half of the seed on the plain microscope slide

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upright on its cut side, and carefully slice the seed coat from the left and right sides of the standing micropylar-half seed to expose the embryo. 5. Trim the tissues around the embryo as much as possible. 6. Move the trimmed micropylar-half seed onto the concavity slide with the 1.5 mm deep well. 7. Add ~100 μl nuclease-free water into the well. 8. Using the beveled edge of the needle, carefully remove the remaining seed coat and endosperm and isolate the globular stage embryo (see Note 5). 9. After removing all seed tissues surrounding the dissected embryo, the cleaned embryo is transferred into another clean concavity slide with a small amount of nuclease-free water using a micropipette with a cut off 1 ml tip. 10. If you need to collect different embryo regions, separate the embryo proper from the suspensor using a new needle (Fig. 1) (see Note 6). 11. Using a fine point forceps, transfer the embryo, embryo proper, or suspensor directly into a microfuge tube suspended within liquid nitrogen. Inspect the forceps under a dissecting microscope to ensure that the embryo, or specific embryo region, was removed successfully. 12. We are able to dissect globular stage embryos at the rate of ~10 per hour. Be patient, as this step takes practice. Store the collected embryos and embryo regions at 70  C until use in downstream experiments. Each collection should last no more than 3 h to ensure that the DNA and RNA remain intact because the pods will be on ice during this entire period. 13. Based on our experience, an average of 5 ng and 80 ng of total RNA can be isolated from one globular-stage embryo proper and suspensor, respectively (see Note 7). 3.5 Fixing Seeds with Globular-Stage Embryos for LCM or in situ Hybridization

1. Based on Subheading 2.5, prepare fresh fixative solution in a glass scintillation vial right before collecting pods (see Note 8) and keep the fixative on ice at all times. 2. Using the disposable scalpel with a No. 11 blade, cut the seed in half transversely on a plain glass slide and transfer the micropylar-half seed into the fixative solution. 3. Once all seeds have been collected, vacuum-infiltrate the seeds for 30 min in a vacuum oven at maximum vacuum (~25 in. Hg) with no heat to obtain complete fixation. The seeds should be immersed in the solution. 4. Transfer the vial containing seeds to 4  C and store overnight.

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1. Perform dehydration steps in the glass vial with serial ethanol concentrations in the following order: 75%, 85%, 95%, 100%, 100%, and 100%. At each step, the glass vial should be on the rotator at room temperature for 2–3 h. 2. Seeds can be stored in 100% ethanol at 4  C overnight (see Note 9).

3.7 Seed Dehydration for in situ Hybridization

1. Replace the fixative solution with 0.1 M Sodium Phosphate Buffer (pH 7.0), and incubate for 30 min on a rotator at room temperature. 2. Perform dehydration steps in the glass vial with serial ethanol concentrations in the following order: 5%, 10%, 15%, 20%, 25%, 30%, 40%, 50%, 60%, 70%, 80%, 90%, 100%, 100%, and 100%. At each step, the glass vial should be on the rotator at room temperature for 30 min. 3. Seeds can be stored in 100% ethanol at 4  C overnight.

3.8 Infiltration and Embedding of Seeds for LCM or in situ Hybridization

1. Prepare liquid Paraplast by filling a 1-l beaker with solid Paraplast chips, and place the Paraplast chip-filled beaker in a 58  C incubator until all chips have melted (see Note 10). Keep the liquid Paraplast in the 58  C incubator. 2. Perform seed clearing steps in a glass vial with serial xylene solutions in ethanol using the following order: 25%, 50%, 75%, 100%, and 100% (see Note 11). At each step, the glass vial should be on the rotator at room temperature for 2–3 h. At the last step, fill the glass vial only half way with 100% xylene because the solid Paraplast chips will gradually be added directly into the vial in the following steps and the solid Paraplast chips will take up space within the vial. 3. Perform seed infiltration steps by adding 10 solid Paraplast chips to the cleared seeds in 100% xylene within the glass vial and incubate at room temperature for 1 h (see Note 12). 4. Repeat step 3 three times, adding 10 more chips each time. 5. Add 10 more solid Paraplast chips to the cleared seeds within the glass vial and incubate at 42  C for 1 h (see Note 13). 6. Repeat step 5 two times, adding 10 more chips each time. 7. Discard the xylene and paraffin mixed solution carefully, making sure to retain the infiltrated seeds within the vial. Add the liquid Paraplast prepared earlier to the glass vial with the cleared seeds, and incubate at 58  C for 2–3 h. 8. Change the liquid Paraplast solution at 2–3 h intervals until the xylene is removed and xylene odor is gone. Usually six to eight changes are needed to accomplish this prior to embedding the seeds.

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9. Turn on the hot plate. Place an aluminum crinkle dish on the hottest part of the plate. 10. Pour the infiltrated seeds within the liquid Paraplast into the aluminum dish. 11. Top off the dish with liquid Paraplast. 12. Heat up one end of the spatula under the alcohol lamp, and use the heated spatula to arrange the seeds within the dish until the seeds are at least 5 mm away from each other (see Note 14). 13. Carefully move the dish to the coolest part of the hot plate without disturbing the seeds. 14. Label each dish with a marked tag. 15. Let the liquid Paraplast containing fixed seeds within the dish harden completely to form a block. Store the seed-containing paraffin block at 4  C for least 4 h or until ready to section (see Note 15). 3.9 Sectioning Paraffin-Embedded Seeds for LCM or in situ Hybridization

1. Turn on slide warmer and set at 42  C. 2. Cut the paraffin block from the aluminum dish into small blocks containing only one seed. 3. Heat up one end of the spatula under the alcohol lamp. Apply the heated end of the spatula to the bottom of the paraffin block in order to melt the paraffin, and quickly mount the paraffin block onto the plastic block holder. Add extra paraffin to the side of the plastic block holder, if necessary, to secure the paraffin block on the holder. Place the block holder at 4  C to harden the paraffin for at least 4 h. 4. Before sectioning, place the mounted paraffin blocks on ice. 5. Trim the tissue block such that the face of the tissue block is shaped as a trapezoid (see Note 16). 6. Insert the plastic holder with the trimmed block into the microtome stage. Orient the parallel sides of the trapezoid parallel to the blade’s edge. 7. Bring the stage closer to the blade and determine how to angle the block in relation to the blade. Adjust the block as necessary (see Note 17). 8. Once the block is at the correct angle, begin cutting at 10 μm until the blade reaches the seed. 9. Set the section thickness (usually 5–10 μm). Maximize the thickness, while still allowing for precise microscopic cell recognition. In our experience, the optimal thickness for scarlet runner bean globular stage embryo longitudinal sections is 6 μm.

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10. Slowly begin to turn the wheel of the microtome. When the paraffin sections start to form a ribbon, gently pick up the end of the ribbon with a watercolor brush and lift the ribbon from the blade. 11. When a 10-section ribbon is generated, lock the microtome. Use the watercolor brush to transfer the ribbon from the microtome to aluminum foil covering the top of the lab bench until enough ribbons have accumulated to make a slide (see Note 18). 12. Place a labeled (e.g., date, person sectioning) slide on the slide warmer. Use PEN membrane slides for LCM and Superfrost Plus slides for in situ hybridization. 13. Put 1.5 ml nuclease-free water on the slide. Gently place the ribbons with the desired length onto the water, and arrange ribbons containing serial seed sections on the slide in order of sectioning with a watercolor brush (see Note 19). 14. Leave the slide on the slide warmer for 15–30 min until the sections become transparent and flatten out. 15. Drain excess water off the slide and let the slide dry for ~1 h. Save the slide in a slide box and store at room temperature. 16. A day before in situ hybridization or capturing embryo regions using LCM, deparaffinize the slides by dipping slides in 100% xylene for 2 min twice. Let the slides air dry in a fume hood overnight. The deparaffinized slides are stored in a slide box at room temperature until used for LCM (see Subheading 3.10) or in situ hybridization (see Note 20). 3.10 LCM of Scarlet Runner Bean Globular Stage Embryos

1. LCM usually is performed in a 2–3 h session (see Note 21). 2. We use a Leica LMD 6000 system [4, 23], but other microscopes capable of carrying out LCM can be used (e.g., Pix-Cell II [1]). It is best to read the relevant operation manual before proceeding with LCM. 3. Turn the Leica LMD 6000 power on in the following order: CTR-MIC control box, computer, and laser. During laser warm up, one green light (left) will be displayed. When the second green light (right) turns on, the laser is ready for use. 4. Start the LMD application software when the microscope is fully initialized. 5. Click the “LOAD” slide button from the software menu. The slide holder stage will move forward. Slide the slide holder with the PEN membrane slide containing mounted seed sections onto the slide holder stage until it snaps into place. Make sure the sections on PEN slide are facing down. Return the slide holder stage back by clicking the “Continue” button on the dialog box.

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6. Click the “LOAD” collector button from the software menu. The tube holder stage will move forward. Place the tube tray with loaded PCR tubes into tube holder stage. The tube caps should face up. Add 30 μl extraction buffer into the caps of PCR tubes. The captured embryo regions will fall into the caps after cutting. Depending on the type of experiment, extraction buffer XB from the PicoPure RNA isolation kit can be used for RNA isolation, whereas Buffer ATL from the QIAamp DNA FFPE Tissue Kit can be used for DNA isolation. Return the tube holder stage back by clicking the “OK” button on the dialog box. 7. Test different laser settings (power, aperture, and speed) to obtain the optimized cutting condition for each embryo region (e.g., embryo proper and suspensor) and/or seed part (e.g., seed coat) that you are interested in capturing. The ideal condition is one with relatively quick cutting speed using the lowest power possible, which will allow the cut region to fall easily into the PCR cap (see Note 22). 8. Select “Draw and Cut” mode from the software interface. Use the touch screen pen to outline the specific embryo region of interest for capture directly on the touchscreen. Click the “Start Cut” button to perform the laser cutting. If the embryo or seed region is not completely released after cutting, manually move the laser to cut the attached part by clicking the “Move and Cut” button to release the cut section. Because the extraction buffer in the PCR cap will slowly evaporate due to the heat generated by the microscope lamp, we recommend adding 10 μl nuclease-free water to the cap halfway through the session if the session lasts more than 1 h. 9. When capturing is complete, take out the PCR tube, briefly spin it in a microfuge, then store it at 70  C until RNA or DNA isolation. RNA can be isolated from the captured seed regions for transcriptome analysis [4] or DNA for methylome analysis [24] (see Note 23). 10. Close LMD software and turn off power in the following order: laser, CTR MIC control box, and computer. 11. We have used this LCM procedure successfully for both transcriptome [4] and methylome [24] analysis of specific seed regions and subregions.

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Notes 1. Do not water between sowing and transferring seedlings. The water in the soil mix is enough for germination. The common greenhouse insect pests that we observe are two-spotted spider mites and thrips. Two-spotted spider mites can be treated with

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Forbid 4F Ornamental Insecticide/Miticide (1 ml per gallon water). Thrips can be treated with Hachi-Hachi SC Insecticide (7.5 ml per gallon water). 2. Scarlet runner bean plants start flowering 30–40 days after sowing, and produce clusters of typical legume red flowers (Fig. 1). The two lowermost petals combine to form the “keel,” the uppermost petal is modified into the hood-like “standard,” and side petals spread into the “wings” (Fig. 2). Flowers open at sunrise and close at sunset. Usually, there are two open flowers in a flower cluster (Fig. 1). The cultivar that we use is a dwarf variety, which grows to about 18 in. in height. Because scarlet runner bean is an open pollinator that utilizes bees for pollination under natural field conditions [25, 26], hand-pollination is required in the greenhouse. In our experience, this takes practice and patience in order to be successful and obtain large numbers of developing pods and seeds. Pollinate flowers early in the morning because the heat of the day can affect pollen viability, and the stigma remains receptive to pollen for 1 day only [25]. Peeling, or cutting back the petals of the flowers carefully, makes it easier to identify flower parts at the beginning of hand-pollination. The receptive stigma is shiny and sticky. The flower with the receptive stigma has bright red petals (flowers #2 and #3 in Fig. 2). Flowers that are open for more than 1 day have dark red petals and withered stigmas, and cannot be used for pollen collection or pollination (flower #5 in Fig. 2). Flowers similar to those like flowers #3 and #4 in Fig. 2 are a good source of viable pollen; however, flowers similar to flower #4 in Fig. 2 cannot be used for pollination because their stigmas are withered and senescing. Unopened flowers such as flower #1 in Fig. 2 cannot be used for either collecting pollen or pollination because pollen has not been released from the anthers and their stigmas are dull and unreceptive to pollination. Common bean (Phaseolus vulgaris) can be a useful alternative for studying early embryo development because (1) it has a large embryo and giant suspensor morphologically similar to those of scarlet runner bean [15]; (2) it self-pollinates, making hand-pollination unnecessary [27]; and (3) an annotated complete genome sequence is available [28]. We typically allow a subset of pods on both scarlet runner bean and common bean to develop to maturity, collect the dry seeds, and establish our own seed stocks for both bean varieties. 3. Pollen can be collected from the flowers of the same plant or from different plants. Hand-pollination using different plants will encourage a higher pollination rate. In our laboratory, the pollination rate varies from 10% to 80% depending upon the experience of the person pollinating.

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4. If RNA is to be isolated from whole-mount embryos, or from LCM captured embryo regions, maintaining an RNase-free environment is critical. Clean work areas and equipment extensively—including aluminum foil, gloves, dissecting microscope, scalpels, forceps, ice bucket, pipettes, microtome, slide warmer, brushes, razor blades, plastic blocks, spatulas, and aluminum dishes, among others—with freshly made 0.1% DEPC water or commercially available cleaning reagents, such as RNase Zap. 5. Because a globular stage embryo is embedded within the seed coat, it is a little tricky to remove surrounding seed coat tissues. We do this by stirring the water to move the embryo around so that tissue debris attached to the embryo falls off during the water movement. 6. To avoid cross-contamination of embryo proper and suspensor regions, use different forceps to transfer each embryo region into separate tubes containing liquid nitrogen. 7. Isolated RNA can be used for classical EST sequencing [10, 11] or transcriptome analysis [4]. Because scarlet runner bean suspensor cells are highly polyploid [7], a significant amount of genomic DNA should be able to be isolated for use in chromatin immunoprecipitation (ChIP) sequencing and chromatin studies. 8. Depending on your sample, the fixation time should be adjusted. In our experience, the fixation time for scarlet runner bean seeds is ~12–16 h. Therefore, always collect pods in the afternoon to avoid excessive fixation during the night prior to processing the sample through a series of increasing alcohol concentrations for sample dehydration. 9. Fixed seeds can be stored in 100% ethanol for several months, and both RNA and DNA remain intact. 10. This step will take at least 5 h. The liquid Paraplast will be used in filtration and embedding and should be ready before infiltration and embedding. Therefore, this step needs to be done ahead of time. 11. Xylene is toxic. Handle with care in the fume hood. 12. Wax infiltration must be done gradually in order to preserve seed internal structures. Thus, solid Paraplast chips are dissolved in the xylene-containing vial with the seeds, and more chips are added and dissolved in xylene sequentially. This allows the concentration of Paraplast in xylene to gradually increase within the seed-containing vial. 13. At room temperature, solid Paraplast cannot be dissolved in xylene when the concentration of Paraplast within the xylene reaches a critical point. In order to complete the infiltration steps it is necessary to increase the temperature.

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14. Think in advance how you want to position seeds in the Paraplast so that you can cut the Paraplast into small blocks containing one seed. Usually, approximately 10 globular stage seeds can be placed in one aluminum dish. Seeds should lay flat on the bottom of the aluminum dish so that the seed can be sectioned longitudinally on the microtome. 15. The total tissue preparation time for LCM or in situ hybridization up to this step takes at least a week. We have also used the rapid microwave paraffin method for LCM experiments, which cut the total preparation time to 5 h and reduced RNA degradation [29]. 16. The face of the tissue block should be as small as possible so that as many sections as possible can be placed onto one slide. The edges of the trapezoid should be as parallel as possible; otherwise the ribbon will be curved and will take up too much space on the slide. Do not trim too close to the seed, as that will prevent the sections from forming a ribbon. 17. In a longitudinal seed section, the serial sections that contain embryo proper and suspensor within the same section are called “middle” or “medial” sections (Fig. 3). These are the

Fig. 3 Scarlet runner bean seed sections captured using LCM. The bottom row shows medial sections, which contain complete embryo proper and suspensor regions within the same section. Sections #1, #2, and #5 do not contain a complete suspensor

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only serial sections we use to collect embryo proper and suspensor regions using LCM because they (1) contain morphologically intact embryo proper and suspensor regions and (2) minimize contamination with surrounding seed tissue (e.g., endosperm and seed coat). To maximize the number of middle or medial sections, it is important to orient the face of the tissue block parallel to the blade. This does not mean that you will always get medial sections because it depends on the orientation of the embedded seed and embryo within the seed. However, most of the time this step will help to ensure that you have as many medial sections as possible for use in downstream experiments. In our experience, two to four medial sections (6 μm) out of ~16–20 serial sections through a globular stage embryo can be expected per globular stage seed. 18. Usually only the sections with an embryo will be mounted on the slides. The first 20–30 sections will not contain any part of the embryo. With experience, it will become obvious during sectioning when you will begin to obtain sections with the embryo. 19. Think in advance how many sections you want to put on one slide. Because the ribbon will expand by ~25%, make sure you leave enough space around the ribbon. 20. For in situ hybridization using scarlet runner bean seeds, we follow procedures developed extensively in our laboratory [18, 19, 22]. Our detailed in situ hybridization protocol using radioactive probes is contained within refs. 18 and 22. Our recent method using nonradioactive probes is contained within ref. 19, although the fixing and sectioning steps are the same for both types of probes. 21. To capture as many embryo regions as possible in one LCM session, examine your slides in advance, and take a picture of the slides using the “overview function” of the LMD 6000 system. Print out the slide “overviews” and mark all of the medial sections. This will allow you to “target” specific sections on each slide for embryo region capture. To avoid contamination of embryo regions with seed coat and endosperm tissues, only capture the embryo proper and suspensor from medial sections (Fig. 3). To prevent cross-contamination between embryo proper and suspensor regions (1) the junction of the embryo proper and the suspensor should not be captured, and (2) only one embryo region should be captured during the same LCM session. 22. The settings that we use to capture globular stage embryo proper and suspensor regions with the Leica LMD 6000 system are as follows: magnification ¼ 20
; power ¼ 31; aperture ¼ 12; speed ¼ 12.

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23. The number of captures required depends on the amount of RNA or DNA needed for downstream experiments. Based on our experience, an average of 0.3 ng and 0.8 ng RNA can be obtained per scarlet runner bean embryo proper and suspensor capture, respectively. To maximize RNA or DNA quality, we recommend that all captures should be done within 1 month after fixation for RNA, and 3 months after fixation for DNA. Using an embedding station can speed up the infiltration and embedding steps, which could maximize RNA or DNA quality. Total RNA from LCM captured embryo regions can be isolated using the PicoPure RNA Isolation Kit (Thermo Scientific) according to the manufacturer’s instructions. RNA-Seq libraries can be prepared using as little as 5 ng of total RNA from LCM-captured embryo regions. Double-stranded cDNA can be synthesized and then amplified using Ovation RNA-Seq System V1 (NuGen) according to the manufacturer’s instructions. RNA-Seq libraries can be generated from the doublestranded cDNA using any commercially available kits. DNA from LCM-captured embryo regions can be isolated using FFPE DNA isolation kit (QIAGEN), and isolated DNA can be used to construct Bisulfite-Seq libraries for methylome studies [24].

Acknowledgments Our work with scarlet runner bean embryos was supported by grants from the US Department of Agriculture, Ceres, Inc., and the National Science Foundation. References 1. Kerk NM et al (2003) Laser capture microdissection of cells from plant tissues. Plant Physiol 132(1):27–35 2. Deal RB, Henikoff S (2010) A simple method for gene expression and chromatin profiling of individual cell types within a tissue. Dev Cell 18 (6):1030–1040 3. Amatori S et al (2014) PAT-ChIP coupled with laser microdissection allows the study of chromatin in selected cell populations from paraffin-embedded patient samples. Epigenetics Chromatin 7:18 4. Belmonte MF et al (2013) Comprehensive developmental profiles of gene activity in regions and subregions of the Arabidopsis seed. Proc Natl Acad Sci U S A 110(5): E435–E444

5. Palovaara J et al (2017) Transcriptome dynamics revealed by a gene expression atlas of the early Arabidopsis embryo. Nat Plants 3 (11):894–904 6. Slane D et al (2014) Cell type-specific transcriptome analysis in the early Arabidopsis thaliana embryo. Development 141 (24):4831–4840 7. Brady T (1973) Feulgen cytophotometric setermination of the DNA content of the embryo proper and suspensor cells of Phaseolus coccineus. Cell Differ 2(2):65–75 8. Clutter M et al (1974) Macromolecular synthesis during plant embryogeny. Cellular rates of RNA synthesis in diploid and polytene cells in bean embryos. J Cell Biol 63(3):1097–1102 9. Henry KF, Goldberg RB (2015) Using giant scarlet runner bean embryos to uncover

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regulatory networks controlling suspensor gene activity. Front Plant Sci 6:44 10. Kawashima T, Goldberg RB (2010) The suspensor: not just suspending the embryo. Trends Plant Sci 15(1):23–30 11. Le BH et al (2007) Using genomics to study legume seed development. Plant Physiol 144 (2):562–574 12. Sussex I et al (1973) Biosynthetic activity of the suspensor of Phaseolus coccineus. Caryologia 25:261–272 13. Yeung EC, Sussex I (1979) Embryogeny of Phaseolus coccineus: the suspensor and the growth of the embryo-proper in vitro. Z. Pflanzenphysiol 91(5):423–433 14. Walbot V et al (1972) Macromolecular synthesis during plant embryogeny: rates of RNA synthesis in Phaseolus coccineus embryos and suspensors. Dev Biol 29(1):104–111 15. Walbot V, Clutter M, Sussex I (1972) Reproductive development and embryogeny in Phaseolus. Phytomorphology 22:59–78 16. Lorenzi R et al (1978) Embryo-suspensor relations in Phaseolus coccineus: Cytokinins during seed development. Planta 143:59–62 17. Zhan J et al (2015) RNA sequencing of lasercapture microdissected compartments of the maize kernel identifies regulatory modules associated with endosperm cell differentiation. Plant Cell 27(3):513–531 18. Weterings K et al (2001) Regional localization of suspensor mRNAs during early embryo development. Plant Cell 13(11):2409–2425 19. Henry KF et al (2018) A shared cis-regulatory module activates transcription in the suspensor of plant embryos. Proc Natl Acad Sci U S A 115(25):E5824–E5833 20. Henry KF, Kawashima T, Goldberg RB (2015) A cis-regulatory module activating

transcription in the suspensor contains five cis-regulatory elements. Plant Mol Biol 88 (3):207–217 21. Kawashima T et al (2009) Identification of cis-regulatory sequences that activate transcription in the suspensor of plant embryos. Proc Natl Acad Sci U S A 106(9):3627–3632 22. Cox KH, Goldberg RB (1988) In: Shaw CH (ed) Analysis of plant gene expression. Plant molecular biology: a practical approach. IRL Press, Oxford, United Kingdom 23. Sanders PM (2005) Differentiation and degeneration of cells that play a major role in tobacco anther dehiscence. Sex Plant Reprod 17:219–241 24. Lin JY et al (2017) Similarity between soybean and Arabidopsis seed methylomes and loss of non-CG methylation does not affect seed development. Proc Natl Acad Sci U S A 114 (45):E9730–E9739 25. Blackwall RLC (1971) A study of the plant/ insect relationships and pod-setting in the runner bean (Phaseolus multiflorus). J Hort Sci 46:365–379 26. Quagliotti L, Marletto F (1987) Research on the pollination of runner bean (Phaseolus coccineus) for dry grain production. Adv Hortic Sci 1(1):43–49 27. Singh SP, Gepts P, Debouck DG (1991) Races of common bean (Phaseolus vulgaris, Fabaceae). Econ Bot 45(3):379–396 28. Schmutz J et al (2014) A reference genome for common bean and genome-wide analysis of dual domestications. Nat Genet 46 (7):707–713 29. Inada N, Wildermuth MC (2005) Novel tissue preparation method and cell-specific marker for laser microdissection of Arabidopsis mature leaf. Planta 221(1):9–16

Chapter 16 Microscopical Detection of Cell Death Processes During Scots Pine Zygotic Embryogenesis Jaana Vuosku and Suvi Sutela Abstract Programmed cell death (PCD) processes are essential in the plant embryogenesis. To understand how PCD operates in a developing seed, the dying cells need to be identified in relation to their surviving neighbors. This can be accomplished by the means of in situ visualization of fragmented DNA—a well-known hallmark of PCD. In the developing Scots pine (Pinus sylvestris L.) seed, several tissues die via morphologically different PCD processes during the embryogenesis. Here, we describe the protocols for the characterization of Scots pine seeds at the early and late developmental stages and, further, the localization of nucleic acids and DNA fragmentation by the acridine orange staining and TUNEL (terminal deoxynucleotidyl transferase (TdT)-mediated deoxyuridine triphosphate (dUTP) nick end labeling) assay in the dying seed tissues. Key words Acridine orange staining, DNA fragmentation, Pinus, Programmed cell death, Scots pine, TUNEL assay, Zygotic embryogenesis

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Introduction In the life cycle of plants, the first signs of programmed cell death (PCD) are seen already during embryogenesis, when certain cells or even entire tissues or organs die for correct embryonic pattern formation [1]. In a cell, the main target of the PCD machinery is the nucleus, and the degradation processes include both chromatin and nuclear envelope [2]. Chromatin degradation is not restricted to the nucleus but may also take place in the cytoplasm [3]. Furthermore, in a PCD process with necrotic-like morphology, cells break down and fragmented nucleic acids can be detected also in the surrounding extracellular space [4]. In situ DNA cleavage can be visualized in individual cells by the TUNEL (terminal deoxynucleotidyl transferase (TdT)-mediated deoxyuridine triphosphate (dUTP) nick end labeling) assay [5] as well as on the basis of acridine orange (AO) fluorescence [6]. Genomic DNA fragmentation generates a multitude of DNA double-strand breaks with free

Martin Bayer (ed.), Plant Embryogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2122, https://doi.org/10.1007/978-1-0716-0342-0_16, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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30 -hydroxyl (30 -OH) groups. This character forms the basis for the TUNEL assay in which labeled dUTPs are added to the accessible 30 -OH ends and subsequently visualized depending on the introduced label. AO is a cell-permeant nucleic acid selective dye that emits green fluorescence when bound to double-stranded DNA (dsDNA) and red fluorescence at when bound to single-stranded DNA (ssDNA) or RNA. Pinus species of the gymnosperms constitute an evolutionarily old group of vascular plants that last shared a common ancestor with angiosperms about 300 mya [7]. Thus, the anatomy as well as cell and molecular biology of embryogenesis in pines differ significantly from those of Arabidopsis, the angiosperm model species [8]. In Scots pine (Pinus sylvestris L.), the zygotic embryo development takes 2 years. Generally, wind pollination in Finland occurs in late May or early June, after which the pollen tube germination slows down and fertilization does not occur until about 1 year later [9]. The time of fertilization and, consequently, embryo development varies between years in the same locality according to the effective temperature sum (d.d.) (i.e., the heat sum unit based on the daily mean temperatures minus the adapted +5  C base temperature) [9–11]. The sequence of embryo development is divided into three phases, which include proembryogeny, early embryogeny, and late embryogeny. Proembryogeny contains the stages before suspensor elongation, whereas early embryogeny initiates with the elongation of the suspensor system and terminates with the appearance of the root meristem. The embryo development culminates in the establishment of root and shoot meristems and in the maturation of the embryo during late embryogeny [12]. The developing Scots pine seed provides a favorable model for PCD because several tissues die via morphologically different PCD processes during the embryogenesis [13]. Initially, multiple embryos arise, but only the dominant embryo survives and completes the development [9]. Subordinate embryos are eliminated via autophagic PCD, which also causes the deletion of cells in suspensors that serve temporary functions during embryo development [14]. Embryos grow within the corrosion cavity of the megagametophyte, a haploid maternally originated tissue that forms the nutrient source of developing embryos [12]. Throughout the embryogenesis, the megagametophyte cells in the narrow embryo surrounding region are destroyed by sudden necrotic-like cell death for nourishing the developing embryos [4, 15]. Furthermore, cells of the nucellar layers that surround the outer surface of the megagametophyte die [4] and the collapsed and phenoliferous nucellar cell walls form a barrier to the passage of water and against fungal infections in a mature seed [16, 17]. Here, we describe the procedures for the preparation (Fig. 1a–c) and characterization of developing Scots pine zygotic embryos (Fig. 1d, e) as well as for the fluorescence microscopical

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Fig. 1 Preparation of immature Scots pine seeds and determination of developmental stages of embryos by toluidine blue-stained anatomical sections. The sections were imagined with a light microscope (Nikon Eclipse E600) using a digital camera (QImaging MicroPublisher 5.0 RTV). (a) The preparation was initiated by cutting off the peduncle head of a cone with the smallest scales using a pruning knife. (b) The cone scales were opened and (c) seeds were placed into sterile water which after the seed coat was carefully removed. (d) A toluidine blue-stained section of Scots pine seed at the early embryogeny. The dominant embryo and subordinate embryos in the corrosion cavity surrounded by the embryo surrounding region of the megagametophyte. (e) The dominant embryo in the corrosion cavity at the late embryogeny. cc corrosion cavity, e embryo, esr embryo surrounding region, m megagametophyte, ms megaspore membranes, nc nucellar cap, nl nucellar layers, sr suspensor remnants. Scale bars: 100 μm

detection of DNA fragmentation and PCD in different seed tissues by AO staining (Fig. 2) and the TUNEL assay (Fig. 3).

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Materials Autoclave solutions when appropriate, use sterile H2O when making dilutions.

2.1 Timing of Cone Collection, Seed Preparation, and Fixation

Climate data is collected in the weather observation stations. 1. 1 PBS buffer: 137 mM NaCl, 2.7 mM KCl, 8 mM Na2HPO4  2H2O, 1.7 mM KH2PO4, pH 7.4.

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Fig. 2 Detection of nucleic acids and cell death during Scots pine embryogenesis by acridine orange (AO) staining. The sections were imagined with an HBO 100 mercury lamp in LSM 5 Pascal (Carl Zeiss) confocal laser scanning microscope. The AO is a dual fluorescence dye: Double-stranded nucleic acid fluoresces green, and single-stranded nucleic acids (RNA or fragmented DNA) fluoresce red. (a) The dominant embryo and dying subordinate embryos in the corrosion cavity at the early embryogeny. (b) The dominant embryo filling the corrosion cavity at the late embryogeny. (c) Megagametophyte cells surrounding the corrosion cavity are destroyed by morphologically necrotic cell death with the release of cell debris and nucleic acids into the corrosion cavity. (d) The inner part of the megagametophyte stays alive with no sign of DNA fragmentation. Nuclei emit green, except for rRNA containing nucleoli, which emits red fluorescence. In the cytoplasmic region, the red color indicates the presence of mRNA and thus active gene expression. (e) In a control section without AO the cell walls of megagametophyte exhibit strong autofluorescence. Scale bars: a and b 100 μm; c-e 10 μm

2. 70% EtOH. 3. 95–98% H2SO4. 4. 1 N NaOH. 5. Paraformaldehyde, reagent grade. 6. Sterile H2O. 7. Chemical balance. 8. Filter papers.

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Fig. 3 Detection of nuclear DNA fragmentation by TUNEL assay in Scots pine seed sections imagined with an HBO 100 mercury lamp in LSM 5 Pascal (Carl Zeiss) confocal laser scanning microscope. The TUNEL method takes advantage of the multiple free 30 hydroxyl termini generated by activated endonucleases to insert rhodamine-labeled dUTPs with red fluorescence emission. (a) TUNEL-positive nuclei in the nucellar layer. (b) TUNEL-positive nuclei in the megagametophyte in the vicinity of the corrosion cavity. (c) DNase treatment induced DNA breaks in the nuclei of megagametophyte cells in a positive control section. (d) Autofluorescence detected in the megagametophyte in a negative control section (omission of TdT). Scale bars: 50 μm

9. Fume hood. 10. Funnel. 11. Glassware and stir bar. 12. Gloves and eye protection. 13. Hot plate with magnetic stirrer. 14. Ice. 15. Pasteur pipettes. 16. Petri dishes. 17. Pruning/grafting knife. 18. Rotating mixer.

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19. Small test tubes with caps. 20. Scalpel. 21. Spatula. 22. Thermometer. 23. Tweezers with thin tips. 2.2 Dehydration and Paraffin Infiltration

1. Alcohol burner/spirit lamp. 2. Cold plate/ice–water bath. 3. Graded series of ethanol in sterile H2O (abs EtOH, 95%, 85%, 70%, 60%, 50%, 40%, 30%). 4. Fume hood. 5. Heat block (suitable for test tubes used). 6. Incubation oven. 7. Paraffin embedding molds. 8. Pasteur pipettes. 9. Paraplast® paraffin pellets (mp 56–57  C) (Sigma-Aldrich) or similar. 10. Petri dishes. 11. Plate heater. 12. Rotating mixer. 13. Sample name tags. 14. Tert-butanol (TBA). 15. Tweezers.

2.3

Sectioning

1. Alcohol burner/spirit lamp. 2. Brush. 3. Dissecting needle. 4. Microtome. 5. Pipet. 6. Plate heater. 7. Protective eyewear. 8. Razor blades. 9. Spatula. 10. Sterile water. 11. Super Frost Ultra Plus slides (Menzel-Gl€azer) or similar. 12. Vacuum chamber. 13. Wooden blocks.

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1. 0.05% Toluidine blue O in water 2. Coverslips. 3. Entellan. 4. Fume hood. 5. Glass staining jars; dish, cover, slide rack, metal handle. 6. Light microscope. 7. Plate heater. 8. Xylene or Histochoice Clearing Agent (Sigma).

2.5 Dewaxing and Rehydration of Slide

1. Fume hood. 2. Glass staining jars; dish, cover, slide rack, metal handle. 3. Graded series of ethanol in sterile H2O (abs EtOH, 90%, 70%, 50%, 30%, 15%). 4. Xylene or Histochoice Clearing Agent (Sigma).

2.6 Acridine Orange (AO) Staining

1. 0.1 M phosphate buffer pH 6. 2. 1.6 mM AO in 0.1 M phosphate buffer pH 6. 3. 32 mM AO stock in distilled H2O (store at +4  C in dark). 4. Coverslips. 5. Glass staining jars; dish, cover, slide rack, metal handle. 6. Fluorescence or laser microscope. 7. Fume hood. 8. Nail polish. 9. Vectashield® Mounting medium (Vector Laboratories) or similar.

2.7

TUNEL Assay

1. 10 mM phosphate buffer, pH 7.5. 2. 10 mM Tris–HCl pH 7.5. 3. 1 PBS buffer: 137 mM NaCl, 2.7 mM KCl, 8 mM Na2HPO4  2H2O, 1.7 mM KH2PO4, pH 7.4. 4. Coverslips. 5. DNase reaction mixture: 3 U/μl DNase I in 50 mM Tris HCl, pH 7.5, 10 mM MgCl2, 50 μg/ml BSA. 6. Fluorescence or laser microscope. 7. Fume hood. 8. Glass staining jars; dish, cover, slide rack, metal handle. 9. Graded series of ethanol in sterile H2O (15%, 30%, 50%, 70%, 90%, abs EtOH). 10. Humidified chambers.

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11. In Situ Cell Death Detection Kit, Fluorescein (Roche) or similar. 12. Nail polish. 13. Pipet. 14. Precut pieces of Parafilm. 15. Proteinase-K. 16. Tweezers. 17. Vectashield® Mounting medium (Vector laboratories) or similar.

3

Methods

3.1 Timing of Cone Collection, Seed Preparation and Fixation 3.1.1 Timing of Cone Collection 3.1.2 Protocol for Making Fixative

The timing of cone collections can be adjusted to the early (Fig. 1d) and late embryogeny (Fig. 1e) on the base of the accumulation of the effective temperature sum (see Note 1). In Finland, real-time climate data is collected in the weather observation stations around the country and provided by the Finnish Meteorological Institute. After collection, keep cones (see Note 2) in paper bags at +4  C and prepare as soon as possible. Bear in mind that formaldehyde is toxic. Read material safety data sheet (MSDS) before working with this chemical. Gloves and safety glasses should be worn and solutions made and handled inside a fume hood (see Note 3). 1. Always use fresh fixative (4% paraformaldehyde in 1 PBS buffer) for seeds. Start the making of fixative by adding about 90 ml of 1 PBS to a glass beaker on a stir plate in a fume hood. Use 1 N NaOH or NaOH pellets to raise pH to 11. 2. Weight 4 g of paraformaldehyde in fume hood and add to the PBS solution. Heat and stir solution at approximately +60  C for 15 min or until the solution clears. Take care that the solution does not boil. Cool solution on ice. 3. Adjust pH to 7.4 with H2SO4. Bring to final volume with 1 PBS and filter, for example, with Whatman filter paper.

3.1.3 Seed Preparation and Fixation

1. Cut the peduncle head with the smallest scales off and open the scales of the cone with the pruning knife (Fig. 1a, b). Use the tip of the knife to place the seeds into a drop of sterile water in a petri dish to keep them moist (Fig. 1c). Rinse knife in 70% EtOH when needed. 2. Hold a seed in tweezers and cut the seed coat open from the narrower side of a seed with a sharp scalpel. Carefully remove the seed coat without damaging the underlying megagametophyte tissue.

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3. Put a prepared seed immediately into a small test tube, which contains fixative (see Note 4). Prepare eight to ten seeds in the same way and add to the same tube. Keep tubes on ice during preparation (see Note 5). 4. Incubate samples overnight (ON) on a rotating mixer at +4  C. 5. Next morning remove fixative with a Pasteur pipette and wash samples twice with 1 PBS for 1 h on a rotating mixer at +4  C. 3.2 Dehydration and Paraffin Infiltration

1. Dehydrate samples through an ascending ethanol series (30%, 40%, 50%, 60%, 70%, 85%, and 95% EtOH). Incubate for 1 h in every solution on a rotating mixer at +4  C. Finally, incubate for 1 h in abs EtOH on a rotating mixer at room temperature (RT) (see Note 6). 2. Remove abs EtOH from the samples and add TBA (see Note 7). Incubate for 1 h at +40  C. Replace old TBA solution with a new one and incubate at +40  C ON. Next morning change TBA once again and incubate for 1 h at +40  C (see Note 8). Work in a fume hood. 3. During next 2 days, add paraffin pellets gradually (only one or two pellets at a time) to the tubes at +58  C (see Note 9). Let the previous pellets melt before adding new ones. 4. Leave the tubes open and let TBA evaporate ON at +58  C. Leave also a portion of paraffin pellets in the incubation oven so that you have melted paraffin next day. 5. Change the paraffin with fresh one several times (use a warm transfer pipette or pour away), for example, twice a day for 2 days. 6. Embed seeds in paraffin. Position mold on a plate heater (+58  C), place a sample name tag to the mold and fill with paraffin. Transfer tubes to heat block (+58  C) and pour the content of a tube to a petri dish on a plate heater. Carefully place a seed into the paraffin using warm tweezers. Move the mold onto a cold plate or ice–water bath to solidify paraffin. When solidified store paraffin blocks at +4  C.

3.3

Sectioning

1. Examine the orientation of the mounted pine seed in the paraffin block. 2. Trim the block with a razor blade to size that it fits a wooden block in the orientation desired. 3. Place some paraffin pieces (derived from the trimming of the block) on the wooden block, heat spatula and melt. Melt the paraffin on the side of paraffin block to be attached to the wood, push blocks together and leave to RT to solidify.

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4. Trim the paraffin block to obtain a rectangle. Leave 2–3 mm of paraffin on the sides of the mounted pine seed. Take also excess paraffin off from the top of the pine seed. Cool blocks at +4  C or ice–water bath before cutting with microtome. 5. Follow the instructions on a microtome user guide. Briefly, attach the block to a microtome, adjust position (block edges must be parallel to knife) and lock. Insert sharp blade to its position and start approaching the block first using thin sections (e.g., 10 μM). Wear safety glasses. 6. When approaching and cutting the first corners from the paraffin block adjust the microtome to 40 μM, then decrease the sectioning thickness to 10 μM to produce the ribbon of sections (see Note 10). Clean blade with a brush when needed. When microtome reaches the pine seed, decrease the section thickness to 5–7 μm and cut the whole seed (see Note 11). 7. Place glass slides on a plate heater (+50  C). Fill the glass slide with water. 8. Pick the section ribbon (of approximately eight sections) with a dissecting needle and place it on top of the water to straighten for 5 min. 9. Pipet excess water from the slide and dry them in a vacuum chamber for 1 h. Place slides into an incubation oven (+40  C) to dry ON. Store slides at +4  C. 3.4 Toluidine Blue Staining

1. Place slides in a slide rack into 0.05% toluidine blue solution for 15–30 min (depending on the section thickness). 2. Wash sections twice in water for 2 min. 3. Dry slides by placing them on a plate heater (+50  C) for at least 15 min. 4. Conduct dewaxing of sections with xylene in a fume hood. Place slides in a slide rack into xylene for 15 min. Repeat dewaxing with another xylene incubation. 5. Mount sections by dropping 1–2 drops of Entellan to the sections and place coverslip. Dry the slides on a fume hood in RT for 2 days.

3.5 Dewaxing and Rehydration

1. Preexamine your sections with a microscope and select slides with intact sections containing embryos. 2. Conduct dewaxing of sections in fume hood. Place slides in a slide rack into xylene for 10 min. Repeat dewaxing step with another xylene incubation. 3. Remove xylene by incubation in abs EtOH for 15 min followed by a second abs EtOH treatment of 3 min.

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4. Rehydrate through descending ethanol series with 1 min incubation in every ethanol solution. 5. Proceed to TUNEL assay or AO staining procedure from 15% EtOH solution. 3.6 Acridine Orange Staining of Nucleic Acids

AO is a mutagen and suspected human carcinogen. Handle AO in fume hood and use gloves. Avoid excess of light. 1. Place rehydrated sections in a slide rack into staining dish containing 0.1 M phosphate buffer for 5 min. 2. Transfer negative control (autofluorescence) (see Note 12) to 0.1 M phosphate buffer and other slides to staining solution (1.6 mM AO in 0.1 M phosphate buffer) for 5 min. 3. Wash slides in 0.1 M phosphate buffer for 2 min and finally in distilled water. 4. Mount coverslips in dim light/the dark. Add one drop of Vectashield to the middle of the slide. Place coverslip onto the slide and attach them with nail polish (see Note 13). Store slides in dark at +4  C. 5. Examine and imagine the sections preferable during the next week. Remember to avoid light when handling the slides and complete the imagining promptly (see Note 14). The excitation maximum of AO-stained RNA (and ssDNA) is 460 nm and the emission 650 nm, whereas when AO is associated with dsDNA the excitation maximum is close to 500 nm and emission 525 nm. Choose appropriate light source and filter combinations depending on the microscope. If possible, examine RNA-AO and dsDNA-AO emissions separately and filter emission >650 nm out to exclude the autofluorescence of cells.

3.7 Detection of DNA Fragmentation with TUNEL Assay

Work in fume hood and use gloves as the labeling solution contains toxic and potentially carcinogenic substances. 1. Select slides and include at least two additional slides for the positive and negative TUNEL labeling controls (see Note 15). 2. Incubate rehydrated sections at +37  C in a staining dish containing 10 μl/ml proteinase K in 10 mM phosphate buffer (pH 7.5) for 20 min. 3. Wash sections in 1 PBS for 2 min. 4. Repeat wash step twice. 5. Produce positive control sections by a DNase I treatment prior the TUNEL assay. Take the slide marked as a positive control and dry it by pressing paper tissues to the sides of the slides. Leave the other slides stay in the in 1 PBS buffer.

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6. Pipet 500 μl DNase reaction mixture (3 U/μl DNase I in 50 mM Tris HCl (pH 7.5), 10 mM MgCl2, 50 μg/ml BSA) to the positive control slide. Incubate for 10 min at RT. Rinse twice in 1 PBS buffer. 7. Prepare the TUNEL reaction mixture by combining the TdT enzyme and the labeling solution as instructed in the TUNEL assay kit in use (see Note 16). 8. Dry all slides by pressing paper tissues to the sides of the slides. 9. Depending on the number of sections on a slide, pipet 50–100 μl TUNEL reaction mixture on the sample and positive control sections. 10. Pipet the same amount of the labeling solution on the negative control sections. 11. Cover sections with pieces of Parafilm and displace air bubbles. 12. Incubate slides in a humidified chamber for 1 h at +37  C in the dark (see Note 17). 13. Remove Parafilm coverslips by pipetting 1 PBS around the edges of a Parafilm piece until it floats, then lift it carefully with tweezers. Do not remove Parafilm pieces by force. 14. Wash sections in 1 PBS for 10 min. 15. Repeat wash step twice. 16. Mount coverslips in dim light/the dark. Add one drop of Vectashield to the middle of the slide. Place coverslip onto the slide and attach them with nail polish (see Note 13). Store slides in dark at +4  C. 17. Examine and imagine the sections preferable during the next week. Remember to avoid light when handling the slides. The TUNEL signal can be examined using 450–500 nm as an excitation wavelength and 515–565 nm for emission. Choose appropriate light source and filter combinations depending on the microscope (see Note 18). Imagine negative and positive controls using the same settings.

4

Notes 1. When the cone collection was repeated four times during the growing season, the toluidine blue-stained anatomical preparates of the developing embryos indicated the following developmental pattern [11]. Around 50% of the embryos were at the proembryogeny stage and 50% at the early embryogeny stage (Fig. 1d), when the effective temperature sum was approximately 500 d.d. Nearly 100% of the embryos were at the early embryogeny stage, when the effective temperature sum was

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between 600 and 700 d.d. Approximately 85% of the embryos had reached the late embryogeny stage (Fig. 1e) while the remaining 15% were still at the early embryogeny stage, when the effective temperature sum was over 800 d.d [11]. 2. Generally, a pine cone contains 20–30 seeds; however, the size, number and quality of seeds are genotype dependent. Collect extra cones when the genotype characteristics are unknown. 3. Paraformaldehyde is a polymer of formaldehyde. Paraformaldehyde itself is not a fixing agent, and thus, needs to be broken down into its basic building blocks, formaldehyde. This can be done by heating or basic conditions until paraformaldehyde becomes solubilized. 4. If you mark sample tubes in fixation step with a drawing pen, cover marks with transparent tape. Otherwise, marks easily dissolve during the EtOH treatments. 5. The seeds should sink to the bottom of the tubes in order to be properly fixated. If seeds float on the surface, the tube can be placed to a vacuum chamber for 10 min to ease the process. 6. The procedure can be interrupted if necessary, and samples can be stored for a longer time in 70% EtOH at +4  C. 7. Keep TBA in a warm but not hot place or incubation oven before use because it freezes below 25  C. 8. When TBA evaporation is started the tubes should contain enough paraffin to cover all the seeds. The estimation of paraffin volume can be alleviated by marking the level of TBA to each tube with a permanent marker before adding the first paraffin pellets. 9. You can put a tiny piece of filter paper between the paraffin pellet(s) and samples into the tubes so that paraffin is not directly in touch with the samples. The slower paraffin is absorbed into the samples the better the result. 10. Generating smooth and straight section ribbon may require some optimizing. Start by checking the angle of the blade, blade sharpness, section thickness, block temperature, and wheel rotation. 11. The section ribbons can be proceeded immediately onto slides or stored at +4  C (e.g., in flat cardboard box). The cut sections can also be examined before putting on slides; if the aim is to study PCD of embryos it would be preferable exclude sections containing only megagametophyte cells. 12. Additional control slides can be produced by breaking RNA and/or DNA with an RNase, DNase, or H2O2 treatment, for instance. For RNase treatment keep slide in producer recommended buffer for 30 min at +37  C and thereafter wash

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RNase off with 0.1% diethyl pyrocarbonate (DEPC, please, read MSDS) in 1 PBS for 15 min following wash with 1 PBS for 15 min. Treat positive slides the same but excluding the RNase. 13. A bright colored nail polish is more convenient than a colorless one because in dim light you can more easily see where you have added it. 14. The AO stain fades relatively rapidly. Try to find suitable sections fast and complete the imagining promptly. The higher the used magnification, the faster will the fading occur. 15. Positive control sections for the TUNEL assay are produced prior to the labeling procedure by a DNase treatment that induces DNA breaks (Fig. 3c). Negative control sections are incubated with the labeling solution only—that is without the TdT enzyme (Fig. 3d)—during the TUNEL labeling procedure. 16. Remember to take the labeling solution from a freezer to thaw beforehand. Always prepare the TUNEL reaction mixture just before an experiment and keep it on ice. Prepare only the amount you need for the slides. Do not store mixed reagents for the next experiment. 17. A humidified chamber can be made using a small box (e.g., plastic or glass food storage container) which is temperature resistant. Put a few layers of wet paper towels on the bottom. You can also put two glass stirring rods on the bottom and arrange slides on them to keep the slides away from the wet papers. Cover box with a lid or aluminum foil. 18. The structures of a Scots pine seed can easily be seen in fluorescence microscopy because they exhibit strong autofluorescence. The nuclear counterstaining can be carried out using common DNA intercalating dyes, such as DAPI or Hoechst, for localizing nuclei if necessary.

Acknowledgments The authors wish to warmly thank Dr. Anne Jokela, Dr. Riina Muilu-M€akel€a, M.Sc. Mira S€a€askilahti, M.Sc. Johanna Kestil€a, and Ms. Eeva Pihlajaviita for their significant contribution to the developing and optimization of the presented procedures.

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References 1. Bozhkov PV, Filonova LH, Suarez MF (2005) Programmed cell death in plant embryogenesis. Curr Top Dev Biol 67:135–179 2. Earnshaw WC (1995) Nuclear changes in apoptosis. Curr Opin Cell Biol 7:337–343 3. Yamada T, Takatsu Y, Kasumi M et al (2006) Nuclear fragmentation and DNA degradation during programmed cell death in petals of morning glory (Ipomoea nil). Planta 224:1279–1290 4. Vuosku J, Sarjala T, Jokela A et al (2009) One tissue, two fates: different roles of megagametophyte cells during scots pine embryogenesis. J Exp Bot 60:1375–1386 5. Gavrieli Y, Sherman Y, Bensasson SA (1992) Identification of programmed cell-death in situ via specific labeling of nuclear DNA fragmentation. J Cell Biol 119:493–501 6. Martins CF, Dode MN, Ba´o SN et al (2007) The use of the acridine orange test and the TUNEL assay to assess the integrity of freezedried bovine spermatozoa DNA. Genet Mol Res 6:94–104 7. Bowe LM, Coat G, de Pamphilis CW (2000) Phylogeny of seed plants based on all three genomic compartments: extant gymnosperms are monophyletic and Gnetales’ closest relatives are conifers. Proc Natl Acad Sci U S A 97:4092–4097 8. Cairney J, Pullman GS (2007) The cellular and molecular biology of conifer embryogenesis. New Phytol 176:511–536 9. Sarvas R (1962) Investigations on the flowering and seed crop of Pinus silvestris. Comm Inst Forest Fenn 53:1–198

10. Sirois L, Begin Y, Parent J (1999) Female gametophyte and embryo development of black spruce along a shore-hinterland climatic gradient of a recently created reservoir, northern Quebec. Can J Bot 77:61–69 11. Vuosku J, Jokela A, L€a€ar€a E et al (2006) Consistency of polyamine profiles and expression of arginine decarboxylase in mitosis during zygotic embryogenesis of scots pine. Plant Physiol 142:1027–1038 12. Singh H (1978) Embryology of gymnosperms. Borntrager, Berlin 13. Vuosku J, Sutela S, Tillman-Sutela E et al (2009) Pine embryogenesis: many licences to kill for a new life. Plant Signal Behav 4:928–932 14. Filonova LH, von Arnold S, Daniel G et al (2002) Programmed cell death eliminates all but one embryo in a polyembryonic plant seed. Cell Death Differ 9:1057–1062 15. Vuosku J, Sutela S, S€a€askilahti M et al (2010) Dealing with the problem of non-specific in situ mRNA hybridization signals associated with plant tissues undergoing programmed cell death. Plant Methods 6:7 16. Tillman-Sutela E, Kauppi A (1995) The morphological background to imbibition in seeds of Pinus sylvestris L. of different provenances. Trees-Struct Funct 9:123–133 17. Tillman-Sutela E, Kauppi A (1995) The significance of structure for imbibition in seeds of the Norway spruce, Picea abies (L.) karst. Trees-Struct Funct 9:269–278

Part VI In Vitro Systems to Study Embryogenesis

Chapter 17 Regulation of Somatic Embryo Development in Norway Spruce Sara von Arnold, Tianqing Zhu, Emma Larsson, Daniel Uddenberg, and David Clapham Abstract Somatic embryogenesis in Norway spruce combined with reverse genetics can be used as a model to study the regulation of embryo development in conifers. The somatic embryo system includes a sequence of developmental stages, which are similar in morphology to their zygotic counterparts. The system can be sufficiently synchronized to enable the collection and study of a large number of somatic embryos at each developmental stage. Here we describe a protocol for establishing transgenic cell lines in which genes of interest are upregulated or downregulated. Furthermore, we present methods for comparing embryo morphology and development in transgenic and control cell lines, including phenotyping the embryos, histological analysis, and tracking embryo development. The expression pattern of different genes is determined by GUS reporter assays. Key words Morphological analysis, Norway spruce, Reporter gene expression, Somatic embryogenesis, Tracking embryo development, Transgenic cell lines

1

Introduction The basic features of the plant body are established during embryogenesis. Thus all of the rudimentary structures and information necessary for the development of a mature tree, which might grow to be several 100 years old, are present in the embryo. Most of our existent knowledge about the regulation of plant embryogenesis is based on studies of the angiosperm model plant Arabidopsis (Arabidopsis thaliana). In contrast, our knowledge concerning the regulation of embryo development in conifers is limited. Angiosperms and gymnosperms, which shared a final common ancestor about 300 million years ago [1], have evolved different patterns during embryo development. Comparative studies of the regulation of embryo development in angiosperms and gymnosperms are therefore interesting from an evolutionary point of view.

Martin Bayer (ed.), Plant Embryogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2122, https://doi.org/10.1007/978-1-0716-0342-0_17, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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The cell division pattern during embryo development is highly regular in Arabidopsis, as compared to that in gymnosperms. In Arabidopsis, embryo-defective mutants have been used to study zygotic embryogenesis. No such mutants have been characterized in gymnosperms. Other limitations when it comes to studying gymnosperm zygotic embryogenesis are the long generation time, irregular flowering, unsynchronized development of embryos within the same cone, and inaccessibility of the embryo inside the ovule. Most of these problems can be overcome by using somatic embryos grown in vitro. The somatic embryo system includes a stereotyped sequence of developmental stages that can be synchronized by specific treatments, making it possible to collect a large number of somatic embryos at a specific developmental stage (see Fig. 1) [2]. The process from differentiation of early somatic embryos to production of mature embryos occurs within 5–7 weeks, much faster than the development of zygotic embryos within the seed, which takes 5 months. The sequence of embryo development in gymnosperms can be divided into three phases [3]: (1) Proembryogeny—all stages before elongation of the suspensor; (2) Early embryogeny—all stages after elongation of the suspensor and before establishment of the root meristem; (3) Late embryogeny—intensive histogenesis including the establishment of stem cell niches in root and shoot meristems. These phases are followed by embryo maturation and germination. The early proembryogeny phase cannot be compared between zygotic and somatic embryos, but early and late embryogeny are similar in zygotic and somatic embryos. Somatic embryogenesis combined with reverse genetics has therefore become a useful tool to study the regulation of embryogenesis in conifers. In addition, somatic embryos can be used for large-scale vegetative propagation of economically important conifers, and have a great potential to capture the genetic gain in breeding programs [4]. Embryogenic cultures of Norway spruce proliferate as proembryogenic masses (PEMs), which are characterized by the presence of small meristematic cells and elongated highly vacuolated cells (see Fig. 1a and e). Early somatic embryos (EEs) differentiate from PEMs. The time for the transition from proliferation to embryo differentiation varies among cell lines from a couple of days up to 2 weeks. EEs have a polar structure with a compact globular embryonal mass in the apical part and vacuolated, expanding suspensor cells in the basal part (see Fig. 1b and e). There is a distinct border between the apical and the basal parts. The embryonal mass consists of densely cytoplasmatic cells delineated by a distinct protoderm [5, 6]. In late embryos (LEs) (see Fig. 1c and e), the embryonal mass has increased in size, and the terminally differentiated suspensor cells are successively eliminated by programmed

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Fig. 1 Somatic embryogenesis in Norway spruce. (a) Proembryogenic masses, PEMs. (b) Early embryo, EE. (c) Late embryo, LE. (d) Mature embryo, ME. EM embryonal mass, S suspensor. Scale bar ¼ 100 μm. (e) Schematic representation of the developmental stages during somatic embryo development. Stage 1, proliferating PEMs in the presence of the plant growth regulators (+PGR) auxin and cytokinin. Stage 2, differentiating EE after 1 week on prematuration medium lacking PGRs (-PGR). Stage 3, EE after 1 week on maturation medium containing abscisic acid. Stage 4, LE after 2 weeks on maturation medium. Stage 5, early maturing embryo after 3–4 weeks on maturation medium, characterized by the initiation of cotyledons. Stage 6, almost fully mature embryo after 4–5 weeks on maturation medium. Stage 7, fully mature cotyledonary embryo after 5–7 weeks on maturation medium. The times given for development of EEs, LEs, and mature embryos represent average times, which vary among cell lines. (Originally published in ref. 9, with the permission of the editor)

cell death (PCD), a process that starts already during differentiation of EEs [7, 8]. Furthermore, the shoot and the root meristems are delineated. During embryo maturation the cells expand and start to accumulate storage reserves. By using the somatic embryo system in Norway spruce it has been possible to gain information about the regulation of embryo development in a conifer. The auxin responses during somatic embryo development in Norway spruce have been shown by analyzing transgenic cell lines expressing the GUS reporter gene under an auxin-responsive promoter [9]. When polar auxin transport (PAT) is blocked during embryo development the endogenous

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auxin content increases and the frequency of cells eliminated by PCD decreases in early embryos. This results in abnormal embryo morphology, and the mature embryos lack an organized shoot apical meristem and have fused cotyledons [10, 11]. The establishment of the apical–basal polarity in early embryos is dependent on a gradient of cells at different stages of PCD along its apical–basal axis, and dysfunctional PCD leads to embryonic aberrations [8, 12]. In a similar way as has been shown in Arabidopsis, members of the WUSCHEL-RELATED HOMEOBOX (WOX) gene family regulate patterning during embryo development in Norway spruce. PaWOX8/9 is important for the correct orientation of the cell division plane and cell fate determination of the stem cells at the basal part of the embryonal mass [13], and PaWOX2 for the specification of the protoderm [6]. Furthermore, PaWOX3 regulates lateral organ formation [14]. Both similarities and differences between Norway spruce and Arabidopsis have been identified in the regulation of the transition from the early morphogenic phase to the late maturation phase [15]. Furthermore, in Arabidopsis, but not in Norway spruce, overexpression of a LEAFY COTYLEDON1 (LEC1)-like gene is sufficient to promote embryonic properties postembryonically [16]. However, a LEC1-like gene (PaHAP3A) in Norway spruce might be an early genetic maker of tissues harboring embryogenic potential. Embryogenic cultures of Scots pine (Pinus sylvestris) proliferate by a cleavage-like process [17], which is different from the proliferation pattern in Norway spruce (see above). These differences are probably related to the fact that zygotic embryos in Pinus species but not Picea species go through a cleavage process [18]. Transcripts that are highly abundant during the cleavage process in zygotic embryos of Scots pine accumulate differently during somatic embryogenesis in Scots pine and Norway spruce [19, 20]. When altering the expression of these transcripts in embryogenic cultures of Norway spruce the proliferation pattern changes and becomes more similar to the cleavage-like proliferation process occurring in embryogenic cultures of Scots pine [20]. In this chapter we present the protocols we have developed for studying the regulation of somatic embryo development in embryogenic cultures of Norway spruce. The methods used for initiating and proliferating embryogenic cultures have been described previously [2]; for recent advances in this area, see ref. 21 and references therein.

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Materials

2.1 Standard Procedures for Stimulating Development and Maturation of Somatic Embryos

1. Embryogenic cultures proliferating in liquid proliferation medium. 2. Proliferation medium: half-strength liquid LP medium containing plant growth regulators (PGRs). 18.8 mM KNO3, 7.5 mM NH4NO3, 1.5 mM MgSO4, 2.5 mM KH2PO4, 3.0 mM CaCl2, 50 μM Fe-EDTA, 10 μm Zn-EDTA, 10 μM MnSO4, 10 μM H3BO3, 0.1 μM Na2MoO4, 0.01 μM CuSO4, 0.01 μM CoCl2, 4.5 μM KI, 4.9 μM pyridoxine–HCl, 16.3 μM nicotinic acid, 26.7 μM glycine, 15 μM thiamine–HCl, 500 μM myoinositol, 1 mM D-glucose, 1 mM D-xylose, 1 mM L-arabinose, 0.56 μM L-alanine, 0.13 μM L-cysteine–HCl, 0.06 μM Larginine, 0.08 μM L-leucine, 0.06 μM L-phenylalanine, 0.06 μM L-tyrosine for full-strength medium. The medium is diluted to half-strength and supplemented with D-sucrose (30 mM), 2,4-dichlorophenoxyacetic acid (2,4-D, 9 μM), and N6-benzyladenine (BA, 4.4 μM). The pH is adjusted to 5.8 prior to autoclaving. L-Glutamine (3 mM) is filter-sterilized and added into autoclaved and cooled medium (see Note 1). 3. Prematuration medium: half-strength liquid LP medium lacking PGRs (see Note 1). 4. Maturation medium: solidified BMI-SI medium containing abscisic acid (ABA). 23 mM KNO3, 3.5 mM NH4NO3, 1.5 mM MgSO4, 0.6 mM KH2PO4, 1.5 mM CaCl2, 100 μM Fe-ETDA, 10 μM Zn-EDTA, 5 μM MnSO4, 10 μM H3BO3, 0.1 μM Na2MoO4, 0.01 μM CuSO4, 0.01 μM CoCl2, 4.5 μM KI, 2.5 μM pyridoxine–HCl, 4 μM nicotinic acid, 26.7 μM glycine, 3 μM thiamine–HCl, 5.5 mM myoinositol, 90 mM Dsucrose, and 500 mg/L casein hydrolysate. The medium is solidified with 0.35% (w/v) Gelrite. The pH value is adjusted to 5.8 before autoclaving. L-Glutamine (3 mM) and ABA (30 μM) are filter-sterilized and added to the autoclaved and cooled medium prior to pouring into sterile petri dishes (see Note 1). The medium is overlaid by two filter papers (Whatman no. 2).

2.2 Establishment of Transgenic Embryogenic Cell Lines

1. Embryogenic cultures proliferating in liquid proliferation medium. 2. Agrobacterium tumefaciens (e.g., strain GV3101 or C58C1) carrying the vector construct to be used (see Note 2). 3. Petri dishes with solidified proliferation medium overlaid by a filter paper (Whatman no. 2). 4. Petri dishes with selection medium I: solidified proliferation medium supplemented with 400 mg/L timentin and 250 mg/ L cefotaxime (see Note 3).

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5. Petri dishes with selection medium II: solidified proliferation medium supplemented with 400 mg/L timentin, 250 mg/L cefotaxime, and an appropriate substance for selection of transformed cells. 2.3

Phenotyping

1. Embryogenic cultures containing EEs and LEs (see stages 3 and 4 in Fig. 1e). 2. SeaPlaque agarose.

2.4 Confocal Microscopy

1. Embryogenic cultures containing EEs and LEs (see stages 3 and 4 in Fig. 1e). 2. SeaPlaque agarose. 3. Laser scanning confocal microscope.

2.5 Histological Analysis of Somatic Embryos

1. Embryogenic cultures containing embryos at the developmental stage to be analyzed. 2. Fixation solution: PBS buffer (pH 7) containing 4% paraformaldehyde, 0.25% glutaraldehyde. 3. Dehydration solutions: 0.85% NaCl; 50% ethanol, 0.85% NaCl; 70% ethanol, 0.85% NaCl; 85% ethanol, 0.85% NaCl; 95% ethanol; 100% ethanol. 4. Embedding agents: Histo-Clear II; Histowax.

2.6 Progression of Somatic Embryo Development

1. Embryogenic cultures incubated in prematuration medium for 1 week. 2. Petri dishes with maturation medium. 3. SeaPlaque agarose.

2.7 Time-Lapse Tracking of Somatic Embryo Development

1. Embryogenic cultures incubated in prematuration medium for 1 week.

2.8 GUS Reporter Gene Expression

1. Transgenic lines harboring the GUS gene fused to the promoter region of the gene of interest.

2. Petri dishes with maturation medium.

2. 90% acetic acid. 3. GUS buffer: 50 mM sodium phosphate buffer (pH 7.0), 0.25% Triton X-100, 1 mM Fe3+/Fe2+ CN, 1 mM Na2EDTA. 4. GUS buffer supplemented with 1 mM 5-bromo-4-chloro-3indole β-glucuronic acid.

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Methods The methods we present in this chapter are used for studying the regulation of somatic embryo development in embryogenic cultures of Norway spruce. The functions of different genes are analyzed in transgenic cell lines in which the genes of interest are upregulated or downregulated (described under Subheading 3.2). The effects of upregulating or downregulating specific genes on embryo morphology and development are analyzed by phenotyping the embryos (described under Subheadings 3.3 and 3.4), histological analysis (described under Subheading 3.5) and tracking embryo development (described under Subheadings 3.6 and 3.7) (see Note 4). The expression patterns of the various genes are determined by analyzing the GUS activity in reporter lines (described under Subheading 3.8). Similar approaches can be taken for studying how different processes affect embryo development by treating somatic embryos at different developmental stages with for example compounds that block PAT, PCD, or histone deacetylases (e.g., see refs. 10, 12, 15).

3.1 Standard Procedures for Stimulating Development and Maturation of Somatic Embryos

The development of somatic embryos in Norway spruce is described in Fig. 1. However, the time it takes for the development of early, late, and mature embryos varies among cell lines and has to be carefully controlled before starting the experiments. All cultures are incubated at 20  C in darkness. 1. Embryogenic cultures are subcultured weekly or biweekly, depending on the growth rate of the cultures, by transferring 3 mL of settled cell aggregates into 97 mL proliferation medium (in 250-mL Erlenmeyer flasks). The cultures are grown on a gyratory shaker (100 rpm). 2. For stimulating differentiation of early embryos, the cultures are first washed twice by transferring 3 mL settled cell aggregates to 15-mL Falcon tubes containing 10 mL prematuration medium. 3 mL washed cell aggregates are transferred into 97 mL prematuration medium in 250-mL Erlenmeyer flasks. The cultures are grown for about 1 week as described in step 1 (see Notes 5 and 6). 3. The cell aggregates are plated in a thin layer on filter papers placed on maturation medium. 4. The filter papers are transferred to fresh maturation medium every second week.

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3.2 Establishment of Transgenic Embryogenic Cell Lines

The Agrobacterium cocultivation method we present here works well for most embryogenic cell lines of Norway spruce (see Note 7). 1. Embryogenic cultures are cocultivated with A. tumefaciens in liquid proliferation medium for 5 h. 2. The cultures are poured onto filter papers placed on solidified proliferation medium. 3. After 48 h of inoculation the filter papers are transferred to selection medium I. 4. After 4 days the filter papers are transferred to selection medium II. 5. The filter papers are weekly transferred to fresh selection medium II. 6. Putative transformants, proliferating after 1–2 months on selection medium are analyzed for the presence of the transgene by PCR.

3.3 Phenotyping Early and Late Somatic Embryos

1. Embryo samples are resuspended in 2 mL of water or prematuration medium and then solidified with 1.2% SeaPlaque agarose in 60 mm petri dishes (see Note 8). 2. The frequency of embryos with various phenotypes is analyzed using an inverted microscope (see Note 9).

3.4 Confocal Microscopy of Early and Late Somatic Embryos

1. Early or late embryos are transferred to 60 mm petri dishes containing 2 mL water or prematuration medium solidified with 1.2% SeaPlaque agarose (see Notes 8 and 10).

3.5 Histological Analysis of Somatic Embryos

The methods for fixation, dehydration, and embedding presented are according to ref. 22. However, other standard protocols can be used.

3.5.1 Fixation

2. The embryos are scanned using a confocal microscope (see Note 11).

1. The samples are placed in ice-cold fixation solution, and vacuum is applied until the fixation solution starts to bubble. 2. Vacuum is released slowly. 3. Samples are transferred to fresh fixation solution and incubated at +4  C for 12–16 h under gentle shaking.

3.5.2 Dehydration

1. The samples are initially dehydrated on ice, first in 0.85% NaCl for 30 min, then in 50% ethanol, 0.85% NaCl for 90 min, and finally in 70% ethanol, 0.85% NaCl for 90 min or overnight. 2. The following dehydration steps are performed at +4  C: 85% ethanol, 0.85% NaCl for 90 min; 95% ethanol for 90 min; 100% ethanol for 90 min; and finally in 100% ethanol for 12–16 h.

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1. The samples are transferred to 100% ethanol at room temperature for 2 h. 2. The samples are sequentially embedded in 50% ethanol, 50% Histo-Clear II at room temperature for 1 h; 100% Histo-Clear II at room temperature for 1 h; 100% Histo-Clear II at room temperature for 1 h; and 50% Histo-Clear II, 50% Histowax at +40 to +50  C for 12–16 h. 3. Finally the samples are soaked in 100% melted Histowax at 60  C for 3 days. Twice a day the samples are transferred to fresh melted Histowax. 4. The embedded embryos are serially sectioned at 4–10 μM. Sections are analyzed in a light microscope (see Note 12). By calculating the average developmental stage of embryos at different time points it is possible to compare the progression of embryo development in various cell lines or to infer possible changes in rates of developmental progression after genetic manipulation or after specific treatments (see ref. 16 and Fig. 2). 5

Average developmental stage

3.6 Progression of Somatic Embryo Development

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Fig. 2 Progression of embryo development in an untransformed control cell line (light grey circles) and in a transgenic cell line, where PaHAP3A (was) upregulated (black squares). After 1 week in prematuration medium embryogenic cultures were plated in a thin layer on maturation medium. From the beginning of the maturation treatment (week 0) until 7 weeks on maturation medium, about 50 embryos were randomly selected and examined each week. The embryos were classified into 5 stages. Stages 1–4 are the same as stages 1–4 presented in Fig. 1. Stage 5 includes both maturing and mature embryos (Stages 5–7 in Fig. 1). Presented data shows the average developmental stage of the selected embryos at each time point based on three biological replicates

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1. Embryogenic cultures, incubated for 1 week in prematuration medium, are plated in a thin layer on maturation medium. 2. Samples are collected weekly, starting from 1 week in prematuration medium to 7 weeks on maturation medium (see Note 13). The samples are first resuspended in 2 mL water or culture medium then mixed with 1.2% SeaPlaque agarose to allow solidification in 60 mm petri dishes (see Note 8). 3. The developmental stage of at least 50 randomly selected embryos in each sample is analyzed using an inverted microscope. The embryo developmental stage is classified as stages 1–5 (see Fig. 1e) (see Note 14). The average developmental stage of the embryos analyzed at each time point is estimated. 3.7 Time-Lapse Tracking of Somatic Embryo Development

By comparing the developmental pattern from early embryos to mature embryos in control and transgenic lines it is possible to identify how different genes regulate embryo development and patterning or to identify how specific treatments influence the developmental pattern (see Fig. 3). 1. After 1 week in prematuration medium the cultures are plated on solidified maturation medium (see Note 13). 2. After 1–2 weeks, 50–100 EEs or LEs (stage 3 or 4 in Fig. 1e) are collected individually and plated on fresh maturation medium (see Note 15). 3. The developmental stage and the phenotype of the selected embryos are scored using an inverted microscope every second day for about 2 weeks (see Note 16).

3.8 GUS Reporter Gene Expression

1. Samples are collected in ice-cold 90% acetic acid and vacuum infiltrated for 10 min at room temperature. 2. The samples are vacuum infiltrated in GUS buffer on ice for 30 min. 3. The GUS buffer is exchanged with fresh GUS buffer supplemented with 5-bromo-4-chloro-3-indole β-glucuronic acid and the samples are vacuum infiltrated on ice for 30 min. 4. Vacuum is released, and the samples are incubated at 37  C in darkness. The incubation time varies depending on the strength of the GUS activity, usually 2–12 h, but in some cases up to 72 h. 5. GUS staining is analyzed under an inverted microscope (see Note 17).

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Fig. 3 Time-lapse tracking of somatic embryo development. LEs were isolated after 2 weeks on maturation medium and transferred to fresh maturation medium. The developmental pathways of the embryos were followed over 10 days, normal embryo in a control cell line (left panel) and abnormal, coneshaped, embryo in a transgenic cell line where PaWOX8/9 was downregulated (see Fig. 3) (right panel) followed two different developmental pathways to maturation. Normal development and degeneration–regeneration development. During the degeneration–regeneration pathway new embryogenic tissue (ET) differentiated from the first selected embryo before maturing embryos (MEs) developed. (Originally published in ref. 13, with the permission of Oxford University Press)

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Notes 1. The compositions of proliferation, prematuration, and maturation media are not critical. The standard media routinely used in the lab for culturing embryogenic cultures of a conifer can be used. The important point is that the proliferation medium should contain auxin and cytokinin, the prematuration medium should not contain plant growth regulators, and the maturation medium should contain abscisic acid. The media we routinely use for proliferation, prematuration, and maturation are presented in ref. 2. 2. Already established methods are used for isolating coding sequences and promoter regions, for vector constructions, and for inserting the vectors into A. tumefaciens (e.g., see refs. 14, 16). 3. Timentin and cefotaxime are used to remove agrobacteria. 4. The transformation procedure can affect embryo morphology and development (see refs. 6, 14). Therefore, it is important to always include both transformed and untransformed controls in the analyses. 5. Depending on the aim of the study and the availability of tissue, larger or smaller Erlenmeyer flasks can be used. In each case it is important to consider the ratio of the volume of the flasks and the volume of the suspension culture. 6. The prematuration treatment should be given for allowing the transition from PEM to somatic embryo, which usually occurs about 1 week after withdrawal of PGRs. However, since the transition time varies among cell lines and also depends on the condition of the cultures, the time has to be adjusted for synchronizing further development of the embryos. 7. New genes can be delivered to cells in embryogenic cultures either by cocultivation with Agrobacterium tumefaciens (e.g., see ref. 13) or by particle bombardment (see ref. 2). In general, Agrobacterium-mediated transformation is preferred since it is faster and simpler than bombardment. In some coniferous species it is not possible to study the regulation of specific processes owing to problems in producing transgenic cell lines. Transgenic cultures of Norway spruce can be used for studying processes present in other coniferous species (see ref. 20). However, before taking such an approach it is crucial to have a basic knowledge about the proliferation pattern of somatic embryos in both species. 8. Since the embryos will be analyzed immediately, the composition of the medium is not important as long as the osmotic potential is not increased or decreased, that is, avoid affecting cellular integrity by higher osmotic potential differences of resuspension medium and culture medium.

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Fig. 4 Phenotyping EEs and LEs. (a) Normal embryo, N, with a distinct, cylindrical, embryonal mass and a suspensor. (b) Twin embryo, T, with two separated embryonal masses and a common suspensor. (c) Coneshaped embryo, C, with supernumerary suspensor cells. The embryo lacks a strict border between the embryonal mass and the suspensor, and suspensor cells can differentiate from the upper part of the embryonal mass. (d) Embryo with meristematic cells in the suspensor, M. (e) Ball-shaped embryo, B, which lacks suspensor cells and has no sign of polarity. (f) Frequency of early LEs with different morphology in a control cell line. Presented data are based on scoring of 500–600 structures in two independent experiments. Bars ¼ 100 μm. (Originally published in ref. 10, with permission of John Wiley & Sons)

9. We routinely classify EEs and LEs in Norway spruce into five phenotypes (see Fig. 4). However, other types of classifications might be needed for particular cell lines (genotypes). In general, it is easiest to phenotype the embryos at the developmental stage corresponding to early LEs. 10. Alternatively, the embryos are sandwiched between two cover glasses. 1–2 drops of water or medium are added between the cover glasses. 11. We have used a Zeiss 780 confocal microscope (Carl Zeiss AG, Germany) equipped with a 488 nm Argon laser under the 20 objective (NA ¼ 0.8) (see Fig. 5). 12. To visualize general anatomical features the sections can be stained by 0.05% aqueous toluidine blue O solution. 13. At least three biological replicates are recommended. 14. It is possible to classify the embryos into seven stages (see Fig. 1). However, it is usually most important to focus on the earlier stages Stage 1–4), and the maturing and mature embryos can be combined into one group (stage 5) (see ref. 16). 15. Make sure that each embryo can be tracked, for example by using a microscope eyepiece graticule (ocular micrometer) or simply by drawing or placing a fine-scaled grid on the bottom of the petri dish.

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Fig. 5 Images of EEs acquired by laser scanning microscopy. Cells are visible owing to their intrinsic autofluorescence, when excited with strong intensity argon laser line at 488 nm. Images are pseudocolored green. The left image shows a normal EE in a control cell line. The right image shows an abnormal EE in a transgenic cell line where PaWOX8/9 had been downregulated. The aberrant morphology is caused by the disturbed orientation of the cell division planes in the basal part of the embryonal mass. Bars ¼ 50 μm. (Originally published in ref. 13, with the permission of Oxford University Press)

16. The selected embryos cultured separately develop faster than embryos in intact cultures (cf Figs. 1 and 3). 17. Depending on the size of the samples and the aim of the analyses, the samples can either be directly analyzed under an inverted microscope or processed further using clearing and/or sectioning protocols. When analyzing GUS activity in mature embryos the samples usually need to be cleared. Various standard methods can be used (e.g., bleaching in hydrogen peroxide (H2O2)–acetic acid (1:1) solution at 90  C for 30 min and then clearing in a 2.5 g/mL chloral hydrate solution at 4  C for at least 12 h after the GUS staining) (see ref. 14). References 1. Smith SA, Beaulieu JM, Donoghue MJ (2010) An uncorrelated relaxed-clock analysis suggests an earlier origin for flowering plants. PNAS 107(13):5897–5902 2. von Arnold S, Clapham D (2008) Spruce embryogenesis. In: Sua´rez MF, Bozhkov PV (eds) Plant embryogenesis. Methods in molecular biology, vol 427. Humana Press, Totowa, New Jersey, pp 31–47 3. Sing H (1978) Embryology of gymnosperms. In: Zimmermann W, Carlquist Z, Ozenda P,

Wulff HD (eds) Handbuch der Pflanzenanatomie. Gebru¨der Borntrager, Berlin, pp 187–241 4. Klimaszewska K, Hargreaves C, Lelu-Walter M-A, Trontin J-F (2016) Advances in conifer somatic embryogenesis since year 2000. In: Germana` MA, Lambardi M (eds) In vitro plant embryogenesis in higher plats. Methods in molecular biology, vol 1359. Humana Press, Totowa, New Jersey, pp 131–166 5. Filonova LH, Bozhkov PV, von Arnold S (2000) Developmental pathway of somatic

Norway Spruce Embryogenesis embryogenesis in Picea abies as revealed by time-laps tracking. J Exp Bot 51 (343):249–264 6. Zhu T, Moschou PN, Alvarez JM, Sohlberg J, von Arnold S (2016) WUSCHEL-RELATED HOMEOBOX 2 is important for protoderm and suspensor development in the gymnosperm Norway spruce. BMC Plant Biol 16:19. https://doi.org/10.1186/s12870016-0706-7 7. Filonova LH, Bozhkov PV, Brukhin VB, Daniel G, Zhivotovsky B, von Arnold S (2000) Two waves of programmed cell death occur during formation and development of somatic embryos in the gymnosperm, Norway spruce. J Cell Sci 113:4399–4411 8. Bozhkov PV, Sua´rez MF, Filonova LH (2005) Programmed cell death in plant embryogenesis. Curr Top Dev Biol 67:135–179 9. von Arnold S, Larsson E, Moschou PN, Zhu T, Uddenberg T, Bozhkov PV (2016) Norway spruce as a model for studying regulation of somatic embryo development in conifers. In: Park Y-S, Bonga JM, Moo H-K (eds) Vegetative propagation of forest trees. National Institute of Forest Science, Seoul, pp 351–372. ISBN 978-89-8176-064-9 10. Larsson E, Sitbon F, Ljung K, von Arnold S (2008) Inhibited polar auxin transport results in aberrant embryo development in Norway spruce. New Phytol 177:356–366 11. Larsson E, Sitbon F, von Arnold S (2008) Polar auxin transport controls suspensor fate. Plant Signal Behav 3:469–470 12. Smertenko A, Bozhkov PV (2014) Somatic embryogenesis: life and death processes during apical-basal patterning. J Exp Bot 55 (1):1343–1360 13. Zhu T, Moschou PN, Alvarez JM, Sohlberg J, von Arnold S (2014) WUSCHEL-RELATED HOMEOBOX 8/9 is important for proper embryo patterning in the gymnosperm Norway spruce. J Exp Bot 65:6543–6552 14. Alvarez J, Sohlberg J, Engstro¨m P, Zhu T, Englund M, Moschou PN, von Arnold S (2015) The WUSCHEL-RELATED

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HOMEOBOX 3 gene PaWOX3 regulates lateral organ formation in Norway spruce. New Phytol 208:1078–1088 15. Uddenberg D, Valladares S, Abrahamsson M, Sundstro¨m J, Sunda˚s-Larsson A, von Arnold S (2011) Embryogenic potential and expression of embryogenesis-related genes in conifers are affected by treatment with a histone deacetylase inhibitor. Planta 234:527–539 16. Uddenberg D, Abrahamsson M, von Arnold S (2016) Overexpression of PaHAP3A stimulates differentiation of ectopic embryos on maturing somatic embryos of Norway spruce. Tree Genet Genomes 12:18. https://doi.org/ 10.1007/s11295-016 17. Abrahamsson M, Valladares S, Merino I, Larsson E, von Arnold S (2017) Degeneration patterning in somatic embryos of Pinus sylvestris L. In Vitro Cell Dev Biol 53:86–96 18. Buchholtz JT (1926) Origin of cleavage polyembryony in conifers. Bot Gaz 8(1):55–71 19. Merino I, Abrahamsson M, Sterck L, CravenBartle B, Canovas F, von Arnold S (2016) Transcript profiling for early stages during embryo development in Scots pine. BMC Plant Biol 16:255. https://doi.org/10.1186/ s12870-016-0939-5 20. Merino I, Abrahamsson M, Larsson E, von Arnold S (2018) Identification of molecular processes that differ among Scots pine somatic embryogenic cell lines leading to the development of normal and abnormal cotyledonary embryos. Tree Genet Genomes 14:34. https://doi.org/10.1007/s11295-018-1247z 21. Pullman GS, Zeng X, Copeland-Kamp B, Crockett J, Lucrezi J, May SW, Bucalo K (2015) Conifer somatic embryogenesis: improvements by supplementation of medium with oxidation-reduction agents. Tree Physiol 35:209–224 22. Karlgren A, Carlsson J, Gyllenstrand N et al (2009) Non-radioactive in situ hybridization protocol applicable for Norway spruce and a range of plant species. J Vis Exp 26:1205. https://doi.org/10.3791/1205

Chapter 18 In Vitro Production of Zygotes by Electrofusion of Rice Gametes Md Hassanur Rahman, Erika Toda, and Takashi Okamoto Abstract In angiosperms, fertilization and embryogenesis occur in the embryo sac, which is deeply embedded in ovular tissue. In vitro fertilization (IVF) systems using isolated gametes have been utilized to dissect postfertilization events in angiosperms, such as egg activation, zygotic development, and early embryogenesis. In addition, using IVF systems, interspecific zygotes and polyploid zygotes have been artificially produced, and their developmental profiles/mechanisms have been analyzed. Taken together, the IVF system can be considered a powerful technique for investigating the fertilization-induced developmental sequences in zygotes and generating new cultivars with desirable characteristics. Here, we describe the procedures for the isolation of rice gametes, electrofusion of gametes, and the culture of the produced zygotes and embryo. Key words Egg cell, Sperm cell, In vitro fertilization, Zygote, Embryo, Rice

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Introduction Fertilization is the central event in the life cycle of most organisms because upon fertilization, parental gametes and genomes are fused and mixed, respectively, to generate progeny. Fertilization and subsequent events in angiosperms, such as gamete fusion, embryogenesis, and endosperm development, occur in the embryo sac, which is deeply embedded in ovular tissue [1–4]. Investigations for the molecular mechanisms in fertilization and early embryogenesis have been impeded by the difficulties in directly researching the biology of the embedded female gamete, zygote, and early embryo. Accordingly, these reproductive and developmental events have been investigated predominantly through analyses of Arabidopsis mutants or transformants coupled with live imaging [5–11]. Alternatively, direct analyses using isolated gametes or zygotes are possible because procedures for isolating viable gametes have been established in a wide range of plant species, including monocotyledonous and dicotyledonous plants [12, 13]. Using viable gametes

Martin Bayer (ed.), Plant Embryogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2122, https://doi.org/10.1007/978-1-0716-0342-0_18, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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isolated from maize, rice, tobacco, and wheat flowers, determination of the precise timing of gamete fusion and utilization of zygotes immediately or shortly after gamete fusion for further cellular and molecular analyses are possible. These IVF systems have been utilized as tools in angiosperm to observe and analyze fertilization and postfertilization processes directly [14]. The IVF system used in angiosperms consists of three basic microtechniques: (1) the isolation and selection of male and female gametes, (2) the fusion of pairs of gametes, and (3) single cell culture [12]. The isolated gametes can be fused electrically [15– 17] or chemically using calcium [18–20], polyethylene glycol [21, 22], or bovine serum albumin [23]. Investigations with calcium-based fusion of maize gametes have shown that an influx of calcium is triggered by gamete fusion and that cell wall formation, an event known as egg activation, is induced by the calcium influx [24]. Moreover, fusion behavior and gamete interaction have been traced by video-enhanced microscopy using PEG-mediated gamete fusion [25]. However, zygotes produced by calcium-, PEG-, BSA-fusion appeared to become arrested in development [19, 22, 23, 25]. Importantly, among these four different procedures, only zygotes produced by electrofusion are known to divide and develop into embryo-like structures and plantlets. A complete IVF system was developed by Kranz and Lo¨rz [26] using maize gametes and electrical fusion. To take advantage of the abundant resources stemming from rice research, a rice IVF system was established by Uchiumi et al. [17]. Recently, Maryenti et al. [27] successfully developed a wheat IVF system to introduce the IVF technology to wheat research. The zygote produced by the electrical fusion of an egg cell with a sperm cell in maize, rice, and wheat developed into two-celled embryo and a globular-like embryo via zygotic embryogenesis in a similar manner to that in planta [17, 26, 27]. Moreover, the IVF produced embryos continued to develop and regenerated into fertile plants. The use of these IVF systems has made it possible to successfully investigate important postfertilization events such as karyogamy [28–30], egg activation and zygotic development [31–33], paternal chromatin decondensation in zygote nucleus [34], the microtubular architecture in egg cells and zygotes [35], positional relationship between gamete fusion site on fused egg cell and first division plane of zygote [36], contribution of parental genome to zygotic development [37, 38], cell cycle progression during zygotic development [39], and epigenetic resetting in early embryos [40]. To develop investigations using zygotes produced in vitro or isolated from pollinated flowers, we recently developed a polyethylene glycol calcium (PEG-Ca2+)-mediated transfection system with zygotes and egg cells [41]. With this procedure, macromolecules such as DNA, RNA, and proteins can be delivered into

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zygotes, and transient expression of genes and proteins of interest in the target cells is also possible. Moreover, Cas9 protein-gRNA ribonucleoproteins or CRISPR/Cas9 vectors were delivered into rice zygotes using the PEG-Ca2+ transfection system, resulting in production of rice plants with targeted mutations [42]. In addition, IVF systems have been utilized to produce interspecific zygotes between maize and wheat, barley or sorghum [31] or to prepare polyploid rice zygotes [38]. These indicate that the IVF system is a powerful technique for generating new cultivars with desirable characteristics as well as for investigating post fertilization events. In our protocol, we describe the procedures for the isolation of rice gametes, electrofusion of gametes, and culture of the produced zygotes and embryos.

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Materials

2.1 Isolation and Transfer of Gametes

1. Rice flowers (see Note 1). 2. Stereomicroscope. 3. Nontreated plastic dishes with a diameter of 3.5 cm. 4. Coverslips (24
40 mm), siliconized at the edges with 5% dichlorodimethylsilane in 1,1,1-trichloroethane (see Note 2). 5. Mineral oil (embryo culture tested grade). 6. Mannitol solution adjusted to 370 mosmol/kg H2O and Autoclaved. 7. Sliding stage for the insertion of a coverslip and a plastic dish. 8. Glass capillaries made from 50 μl aspirator tubes, tip openings 150–250 μl (drawn by hand). 9. Glass needles with fine tips. 10. Manual handling injector.

2.2 Fusion of Gametes

1. Mannitol solution adjusted to 370 mosmol/kg H2O and autoclaved. 2. Mannitol solution adjusted to 520 mosmol/kg H2O and autoclaved. 3. Inverted microscope. 4. Electrofusion apparatus (ECFG21, Nepa Gene, Ichikawa, Chiba, Japan). 5. Manipulator with a double pipette holder. 6. Electrodes (CUY5100Ti100, Nepa Gene) fixed to the pipette holder.

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2.3 Culture of Zygotes into Embryo-Like Structure and Plantlets

1. Nontreated 3.5-cm plastic dishes. 2. Millicell CM inserts, diameter 12 mm (Millipore, Madison, WI, USA). 3. Feeder cells: rice suspension cell culture (Line Oc, provided by RIKEN Bio-Resource Center, Tsukuba, Japan) (see Note 3). 4. Medium for zygote culture: N6Z-medium [43] with modifications: 2 g/l CHU (N6) basal salt mixture, 0.025 mg/l Na2MoO4·2H2O, 0.025 mg/l CoCl2·6H2O, 0.025 mg/l CuSO4·5H2O, 0.01 mg/l retinol, 0.01 mg/l calciferol, 0.01 mg/l biotin, 1 mg/l thiamin–HCl, 1 mg/l nicotinic acid, 1 mg/l pyridoxine–HCl, 1 mg/l choline chloride, 1 mg/l Ca-pantothenate, 0.2 mg/l riboflavin, 0.2 mg/l 2,4-D, 0.02 mg/l cobalamine, 0.02 mg/l p-aminobenzoic acid, 0.4 mg/l folic acid, 2 mg/l ascorbic acid, 40 mg/l malic acid, 40 mg/l citric acid, 40 mg/l fumaric acid, 20 mg/l Na-pyruvate, 1000 mg/l glutamine, 250 mg/l casein hydrolysate, 100 mg/l myoinositol. Osmolality, 450 mosmol/kg H2O adjusted with glucose. pH 5.7 and filter-sterilized. 5. Regeneration media: Solidified MS medium with some modifications [44]: MS salt, MS vitamin, 100 mg/l myoinositol, 2 g/l casamino acids, 30 g/l sucrose, 30 g/l sorbitol, 0.2 mg/l 1-naphthaleneacetic acid (NAA), 1 mg/l kinetin, 0.3% Gelrite. 6. Rooting media: The same as the regeneration media excluding kinetin and NAA.

3

Methods The flow of production of rice zygote by electrofusion and subsequent zygote culture is shown in Fig. 1. Egg cells and sperm cells are isolated from mature rice flowers, and an egg cell and a sperm cell are transferred into fusion droplets on a coverslip. These male and female gametes are electrically fused, and the resulting zygote is cultured on a Millicell membrane insert in N6Z medium with feeder cells.

3.1 Isolation of Gametes

1. Collect the panicles in which some flowers have already opened and others remain unopened. Pick up the unopened flowers from the panicles and dissect them. Isolate the ovaries and anthers, and transfer them separately into 3.5 cm plastic dishes filled with 3 ml of mannitol solution (370 mosmol/kg H2O) for isolating egg and sperm cells, respectively. 2. For egg cell isolation, remove the stigmas from ovaries and transfer them into new 3.5 cm plastic dishes filled with 3 ml of the mannitol solution (see Note 4). Sink the ovaries to the bottom of the dishes and cut them transversely at the middle

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Gamete isolation Egg cell

Sperm cell

Electrode Coverslip Mineral oil Mannitol droplet

Electro-fusion

Fertilized egg cell (Zygote)

Culture of zygote

Millicell insert Feeder cell Zygote

Fig. 1 Schematic illustration of in vitro production of zygotes by electrofusion of rice gametes. An isolated egg cell and a sperm cell are transferred into a fusion droplet of mannitol solution overlaid with mineral oil on a coverslip. Thereafter, gametes are aligned and fixed at one electrode under an AC field and fused by DC pulse. The produced zygotes are cultured on the membrane of a Millicell insert in N6Z medium with feeder cells

with a razor blade (see Note 5). Collect the egg cells released from lower parts of the cut ovaries (see Fig. 2c, d). Approximately 8–12 egg cells are transferred into a mannitol droplet on coverslips using glass capillaries connected with manual handling injector under an inverted microscope (see Fig. 2h, Note 6). 3. For sperm cell isolation, roughly break anthers in mannitol solution with forceps to free the pollen grains (see Fig. 2e). Use the sperm cells released from the burst pollen grains for electrofusion (see Fig. 2f, g, Note 7). 3.2 Fusion of Gametes

1. Overlay a siliconized coverslip with 0.25 ml mineral oil. For tentative storage of isolated egg cells, make one or two μl mannitol droplets (370 mosmol/kg H2O) using glass capillaries connected with manual handling injector. In addition, make 2 μl mannitol droplets (370 mosmol/kg H2O) in two rows, each with six droplets (see Fig. 2h, Note 8). 2. Transfer one egg cell to each of the six mannitol droplets (see Note 9), then transfer one sperm cell to each droplet. 3. Set up the fusion apparatus and adjust the position of electrodes.

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Fig. 2 Isolation of rice egg cells (a–d) and sperm cell (e–g), and electrofusion of isolated gametes (h–l). (a) An ovary harvested from a rice flower before flowering. The red line indicates the incision line on the ovary. (b) Cutting the ovary in mannitol solution. (c) A rice egg cell (arrowhead) being released from basal portion of the dissected ovary. (d) An isolated rice egg cell. (e) A broken anther (arrow) and released pollen grains (arrowheads) in mannitol solution. (f) A pollen grain (arrow) released its content in mannitol solution. Two sperm cells were enclosed with the square. (g) Two rice sperm cells released from pollen grain. (h) An illustration of the fusion droplets on a coverslip covered with mineral oil. The arrowhead indicates a mannitol droplet for tentative storage of isolated egg cells for subsequent fusion. The gray bars indicate electrodes. (i) Alignment of an egg cell with a sperm cell (arrowhead) on one of the electrodes under an alternating current (AC) field in a fusion droplet. (j) Aligned egg and sperm cells after the addition of mannitol solution with a higher osmolality to the fusion drop. The sperm cell becomes oblong (arrowhead). (k) Fusion of gametes following a negative direct current (DC) pulse. The arrowhead indicates the fusion point. (l) A zygote 10 s after fusion. The arrowhead indicates the fusion point. Bars ¼ 50 μm in (d, f, i); 500 μm in e and 10 μm in (g)

4. Align and fix the two gametes at one electrode under an alternating current (AC) field (1 MHz, 5 V rms). By moving the microscope stage, first fix an egg cell to the electrode. Using the same procedure, fix a sperm cell to the female gamete (see Fig. 2i). Adjust the final distance of the electrodes to approximately two to three times the sum of the diameters of the cells. 5. Add approximately 1–2 μl of mannitol solution (520 mosmol/ kg H2O) gently to the fusion droplet using a thin glass capillary (see Fig. 2j, Note 10).

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6. Induce cell fusion by applying a single negative direct current (DC) pulse (40 μs, 12–15 kV/cm) (see Fig. 2k, l, Note 11). 7. Remove the fusion products from the electrode by gently moving the sliding stage. Move the electrodes out of the droplet and conduct the next gamete fusion (see Subheading 3.2, step 4). 3.3 Culture of Zygotes into Embryo-Like Structure and Plantlets

1. Place 0.2 ml zygote culture medium in a Millicell CM insert and put it into a 3.5 cm plastic dish containing 2 ml of the zygote culture medium. Add 40–60 μl of a rice suspension cell culture into the dish as feeder cells. 2. After sterilization of the microcapillary by washing with absolute ethanol and sterilized water, transfer IVF-produced zygotes into fresh mannitol droplets (450 mosmol/kg H2O) twice and then transfer those onto the membranes of a Millicell CM insert (see Fig. 1, Note 12). 3. After overnight culture of zygotes at 26  C in the dark without shaking, continue culture with gentle shaking (30 rpm) (see Fig. 3a–e, Notes 13 and 14). 4. Five days after fusion, remove feeder cells by transferring the Millicell dishes containing the embryos into new 3.5 cm plastic dishes filled with 2 ml of fresh zygote culture medium (see Fig. 3f, Note 15). Continue culturing as above. 5. After 18 days in culture, subculture cell colonies developed from the IVF-produced zygotes onto a regeneration medium by use of a sterilized Pasteur pipette. Incubate under continuous light at 30  C for 12–30 days (see Fig. 3g, Note 16). 6. Transfer the differentiated shoots into a rooting medium and culture them under a 13 h:11 h light–dark cycle at 28  C for 11–13 days (see Fig. 3h, i). 7. Transfer the resulting plantlets to soil pods and grown in environmental chambers as described in Note 1 (see Fig. 3j). If needed, harvest seeds from the regenerated plants (see Fig. 3k) and germinate them.

4

Notes 1. Rice plants (Oryza sativa L. cv Nipponbare) were grown in environmental chamber under condition of 26  C in a 13:11 h light–dark cycle with a photosynthetic photon flux density of 150–300 μmol/m2/s. Under these growth conditions, flowers can be obtained throughout all seasons. 2. Coverslips should be noncoated, as using coated coverslips will result in attachment of the cells to the surface of the coverslip. Coverslips supplied from Fisher Scientific (No. 125485J) are recommended.

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Fig. 3 Development of IVF-produced zygotes into globular-like embryo (a–e) and regeneration of the globularlike embryo into fertile plant (f–k). (a) A zygote 1 h after fusion. (b) A zygote 4 h after fusion. Two nucleoli are indicated by arrowheads. (c) An asymmetric two-celled embryo 18 h after fusion. (d and e) An embryo 48 h after fusion. Panels (d) and (e) present brightfield and DAPI-stained fluorescence images, respectively. (f) A cell mass 5 days after fusion, which developed from the globular-like embryo. (g) A white cell colony 18 days after fusion. (h) A developed cell colony 4 days after transferring the white cell colony (panel g) into regeneration medium (22 days after fusion). Green spots are visible in/on the cell colony. (i) Regenerated shoots. Generation of shoots can be observed after 8 days of subculturing of the white cell colony (26 days after fusion). (j) A plantlet after 12 days of subculturing a regenerated shoot in hormone-free medium (43 days after fusion). (k) A regenerated plant with seed sets (100 days after fusion). Scale bars indicate 50 μm in (a–d, f); 1 mm in (g–i) and 1 cm in (j)

3. Rice suspension cells, Line Oc, were subcultured once weekly according to instructions from RIKEN Bio-Resource Center. No difference has been observed in feeder effects between freshly subcultured cells and 1-week-cultured cells.

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4. Unless the stigmas are removed, ovaries always float on the mannitol solution. To isolate egg cells, sinking ovaries into the mannitol solution is essential. Usually, 15–25 ovaries are put into a dish. 5. Usually, 3–8 egg cells are automatically released from approximately 20 cut ovaries. Gently pushing the middle position of lower part of the cut ovary with a glass needle will produce additional egg cells. 6. Egg cells can be kept in the mannitol droplet at room temperature until 6 h after isolation for conducting IVF with no decrease in fusion efficiency. Alternatively, egg cells can be kept at 4  C overnight for conducting IVF with a slight decrease in fusion efficiency. 7. Sperm cells should be used for IVF within 1 h after isolation. Otherwise, sperm cells appear to degenerate and cannot be fused with egg cells. 8. Take care that the droplets do not spread over the glass surface, but are located inside the oil and have no access to the air. The volume of mannitol droplets can be varied between 0.5 and 2 μl. 9. At each round of fusion procedures, five to six sets of gamete fusions are recommended because conducting many sets of gamete fusion takes time and sperm cell will be degenerate during the course of the experiments. 10. The addition of mannitol solution with a higher osmolality changes the shape of the sperm cell to oblong and makes the attachment of the egg cell to the electrode more stable (see Fig. 2i, j). Without this treatment, egg cells are often released from the electrode upon fusion induced by a DC pulse, and fusion efficiency is greatly reduced. 11. If no cell fusion occurs, reduce the distance between the two electrodes and pulse again. 12. The efficiency of successful electrofusion is approximately 80–90% under optimal conditions. A total of 20–40 egg cells can be isolated from 100 processed ovaries, and 15–30 egg cells can be fused with sperm cells by one experimenter in a day. 13. Gamete fusion usually occurs within 1 s (see Fig. 2k, l). The rice zygotes produced by IVF start to form cell walls (see Fig. 3a) and two nucleoli can be observed in a zygote around 4 h after fusion (see Fig. 3b). At around 12 h after fusion, welldeveloped granular organelles, probably starch granules, are visible in the zygotes and the first asymmetric cell division of the zygotes is observed at 17–22 h after fusion (see Fig. 3c). After the first division, the two-celled embryos continue to develop into early embryos at 40–50 h after fusion (see Fig. 3d, e).

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14. Approximately 90% IVF-produced zygotes divide into two-celled embryos, and 90% of IVF-produced two-celled embryos develop into globular embryos. 15. After 5 days of culture of the IVF-produced zygotes, cocultivation with feeder cells is not needed. 16. Normally, after 4 days of subculture of the cell colony on a solidified-regeneration medium (22 days after fusion), green spots become visible (see Fig. 3h) and the emergence of multiple shoots is observed after 8 days of subculturing (26 days after fusion) (see Fig. 3i). References 1. Guignard ML (1899) Sur les antherozoides et la double copulation sexuelle chez les vegetaux angiosperms. Rev Ge´n Bot 11:129–135 2. Nawaschin S (1898) Revision der Befruchtungsvorgange bei Lilium martagon und Fritillaria tenella. Bull Acad Imp Sci St-Pe´tersbourg 9:377–382 3. Raghavan V (2003) Some reflections on double fertilization, from its discovery to the present. New Phytol 159:565–583 4. Russell SD (1992) Double fertilization. Int Rev Cytol 40:357–390 5. Mori T, Kuroiwa H, Higashiyama T, Kuroiwa T (2006) Generative cell specific 1 is essential for angiosperm fertilization. Nat Cell Biol 8:64–71 6. Berger F (2011) Imaging fertilization in flowering plants, not so abominable after all. J Exp Bot 62:1651–1658 7. Sprunck S, Rademacher S, Vogler F, Gheyselinck J, Grossniklaus U, Dresselhaus T (2012) Egg cell–secreted EC1 triggers sperm cell activation during double fertilization. Science 338:1093–1097 8. Denninger P, Bleckmann A, Lausser A, Vogler F, Ott T, Ehrhardt DW, Frommer WB, Sprunck S, Dresselhaus T, Grossmann G (2014) Male–female communication triggers calcium signatures during fertilization in Arabidopsis. Nat Commun 5:4645 9. Hamamura Y, Nagahara S, Higashiyama T (2012) Double fertilization on the move. Curr Opin Plant Biol 15:70–77 10. Hamamura Y, Nishimaki M, Takeuchi H, Geitmann A, Kurihara D, Higashiyama T (2014) Live imaging of calcium spikes during double fertilization in Arabidopsis. Nat Commun 5:4722 11. Maruyama D, Vo¨lz R, Takeuchi H, Mori T, Igawa T, Kurihara D, Kawashima T, Ueda M, Ito M, Umeda M, Nishikawa S, Groß-Hardt R, Higashiyama T (2015) Rapid elimination of

the persistent synergid through a cell fusion mechanism. Cell 161(4):907–918 12. Kranz E (1999) In vitro fertilization with isolated single gametes. Methods Mol Biol 111:259–267 13. Okamoto T (2011) In vitro fertilization with isolated rice gametes: production of zygotes and zygote and embryo culture. Methods Mol Biol 710:17–27 14. Wang YY, Kuang A, Russell SD, Tian HQ (2006) In vitro fertilization as a tool for investigating sexual reproduction of angiosperm. Sex Plant Reprod 19:103–115 15. Kranz E, Bautor J, Lo¨rz H (1991) In vitro fertilization of single, isolated gametes of maize mediated electrofusion. Sex Plant Reprod 4:12–16 16. Uchiumi T, Komatsu S, Koshiba T, Okamoto T (2006) Isolation of gametes and central cells from Oryza sativa L. Sex Plant Reprod 19:37–45 17. Uchiumi T, Uemura I, Okamoto T (2007) Establishment of an in vitro fertilization system in rice (Oryza sativa L.). Planta 226:581–589 18. Faure JE, Digonnet C, Dumas C (1994) An in vitro system for adhesion and fusion of maize gametes. Science 263:1598–1600 19. Kranz E, Lo¨rz H (1994) In vitro fertilization of maize by single egg and sperm cell protoplast fusion mediated by high calcium and high pH. Zygote 2:125–128 20. Khalequzzaman M, Haq N (2005) Isolation and in vitro fusion of egg and sperm cells in Oryza sativa. Plant Physiol Biochem 43:69–75 21. Sun M, Yang H, Zhou C, Koop H-U (1995) Single-pair fusion of various combinations between female gametoplasts and other protoplasts in Nicotiana tabacum. Acta Bot Sin 37:1–6 22. Tian HQ, Russell SD (1997) Micromanipulation of male and female gametes of Nicotiana

In Vitro Fertilization Using Rice Gametes tabacum: II. Preliminary attempts for in vitro fertilization and egg cell culture. Plant Cell Rep 16:657–661 23. Peng XB, Sun MX, Yang HY (2005) A novel in vitro system for gamete fusion in maize. Cell Res 15:734–738 24. Antoine AF, Faure JE, Dumas C, Feijo JA (2001) Differential contribution of cytoplasmic Ca2+ and Ca2+ influx to gamete fusion and egg activation in maize. Nat Cell Biol 3:1120–1123 25. Sun MX, Moscatelli A, Yang HY, Cresti M (2002) In vitro double fertilization in Nicotiana tabacum (L.): polygamy compared with selected single pair somatic protoplast and chloroplast fusions. Sex Plant Reprod 13:113–117 26. Kranz E, Lo¨rz H (1993) In vitro fertilization with isolated, single gametes results in zygotic embryogenesis and fertile maize plants. Plant Cell 5:739–746 27. Maryenti T, Kato N, Ichikawa M, Okamoto T (2019) Establishment of an in vitro fertilization system in wheat (Triticum aestivum L.). Plant Cell Physiol 60(4):835–843. https://doi. org/10.1093/pcp/pcy250 28. Faure JE, Mogensen HL, Dumas C, Lo¨rz H, Kranz E (1993) Karyogamy after electrofusion of single egg and sperm cell protoplasts from maize: cytological evidence and time course. Plant Cell 5:747–755 29. Ohnishi Y, Hoshino R, Okamoto T (2014) Dynamics of male and female chromatin during karyogamy in rice zygotes. Plant Physiol 165:1533–1543 30. Ohnishi Y, Okamoto T (2017) Nuclear migration during karyogamy in rice zygotes is mediated by continuous convergence of actin meshwork toward the egg nucleus. J Plant Res 130:339–348 31. Kranz E, von Wiegen P, Lo¨rz H (1995) Early cytological events after induction of cell division in egg cells and zygote development following in vitro fertilization with angiosperm gametes. Plant J 8:9–23 32. Sato A, Toyooka K, Okamoto T (2010) Asymmetric cell division of rice zygotes located in embryo sac and produced by in vitro fertilization. Sex Plant Reprod 23:211–217 33. Rahman M, Toda E, Kobayashi M, Kudo T, Koshimizu M, Takahara M, Iwami M, Watanabe Y, Sekimoto H, Yano K, Okamoto

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T (2019) Expression of genes from paternal alleles in rice zygotes and involvement of OsASGR-BBML1 in initiation of zygotic development. Plant Cell Physiol 60:725–737. https://doi.org/10.1093/pcp/pcy030 34. Scholten S, Lo¨rz H, Kranz E (2002) Paternal mRNA and protein synthesis coincides with male chromatin decondensation in maize zygotes. Plant J 32:221–231 35. Hoshino Y, Scholten S, von Wiegen P, Lo¨rz H, Kranz E (2004) Fertilization induced changes in the microtubular architecture of the maize egg cell and zygote—an immunocytochemical approach adapted to single cells. Sex Plant Reprod 17:89–95 36. Nakajima K, Uchiumi T, Okamoto T (2010) Positional relationship between the gamete fusion site and the first division plane in the rice zygote. J Exp Bot 61:3101–3105 37. Toda E, Ohnishi Y, Okamoto T (2016) Development of polyspermic rice zygotes. Plant Physiol 171:206–214 38. Toda E, Ohnishi Y, Okamoto T (2018) Effects of an imbalanced parental genome ratio on development of rice zygotes. J Exp Bot 69:2609–2619 39. Sukawa Y, Okamoto T (2018) Cell cycle in egg cell and its progression during zygotic development in rice. Plant Reprod 31:107–116 40. Jahnke S, Scholten S (2009) Epigenetic resetting of a gene imprinted in plant embryos. Curr Biol 19:1677–1681 41. Koiso N, Toda E, Ichikawa M, Kato N, Okamoto T (2017) Development of gene expression system in egg cells and zygotes isolated from rice and maize. Plant Direct 1:e00010 42. Toda E, Koiso N, Takebayashi A, Ichikawa M, Kiba T, Osakabe K, Osakabe Y, Sakakibara H, Kato N, Okamoto T (2019) An efficient DNAand selectable-marker-free genome-editing system using zygotes in rice. Nat Plants 5:363–368. https://doi.org/10.1038/ s41477-019-0386-z 43. Kumlehn J, Lo¨rz H, Kranz E (1998) Differentiation of isolated wheat zygotes into embryos and normal plants. Planta 205:327–333 44. Hiei Y, Ohta S, Komari T, Kumashiro T (1994) Efficient transformation of rice (Oryza sativa L.) mediated by Agrobacterium and sequence analysis of the boundaries of the T-DNA. Plant J 6:271–282

Chapter 19 Isolated Microspore Culture in Brassica napus Patricia Corral-Martı´nez, Carolina Camacho-Ferna´ndez, and Jose M. Seguı´-Simarro Abstract Isolated microspore culture is the most efficient technique among those used to induce microspore embryogenesis. In the particular case of Brassica napus, it is also the most widely used and optimized. In this chapter, we describe a protocol for microspore culture in B. napus which includes the steps necessary to isolate and culture microspores, to induce microspore-derived embryos, to produce doubled haploid plants from them, as well as to check for the developmental stage of the microspores isolated, their viability, and the ploidy level of regenerated plantlets. Key words Androgenesis, Doubled haploid, Embryogenesis, Haploid, Microspores, Pollen, Rapeseed, Tissue culture

1

Introduction Androgenesis is the process of obtaining an individual with a genome coming exclusively from a male donor. These individuals are usually derived from haploid microspores and young pollen grains. Microspore-derived embryos (MDEs) may remain haploid or become doubled haploid (DH) individuals spontaneously or through the application of some treatments for genome duplication [1]. MDEs are obtained from microspores/pollens upon changing their originally gametophytic developmental program toward embryogenesis [2]. This developmental switch is typically induced through the application of a species-specific in vitro stress treatment [3] to microspores and pollen grains at defined developmental stages [4]. DHs are very useful tools in plant breeding. Among their many different advantages (reviewed in [5, 6]), their most important application is to reduce time and resources to produce pure (100% homozygous) lines for hybrid seed production. Hybrid seeds are typically obtained by crossing two pure, 100% homozygous parental lines, which makes mandatory the previous

Martin Bayer (ed.), Plant Embryogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2122, https://doi.org/10.1007/978-1-0716-0342-0_19, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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production of pure lines. Using conventional approaches, this is a laborious process which requires several years (5–10) of inbreeding and selection. However, DH technology is able to reduce this time (and obviously the resources needed) to some months when the experimental protocols are optimized. From a more basic research perspective, microspore embryogenesis is an inducible morphogenic process convenient for the study of different aspects of plant cell totipotency and embryogenesis itself. There are two main ways to induce microspore embryogenesis: anther culture and isolated microspore culture. Both are in vitro culture techniques whose objective is to generate MDEs from immature pollen or microspores. Anther culture is simpler, since microspore isolation from the anther is not needed, but it is less efficient as well, and the control of culture conditions is more difficult. On the other hand, microspore culture is more technically demanding, but also more efficient and controlled. This makes it the choice for those species where both methods are available, which unfortunately is not the case in many agronomically interesting crops. However, Brassica napus (rapeseed) is a model species for microspore embryogenesis, as well as Nicotiana tabacum (tobacco), Triticum aestivum (wheat), and Hordeum vulgare (barley) [5]. In some lines and cultivars of this species, this process can be easy and efficiently induced with the use of optimized methods. In this chapter we describe a method for isolated microspore culture in B. napus. In addition, we present methods to identify the most suitable developmental stages for microspore/pollen isolation, to determine their viability, and to check the ploidy of the produced MDEs/plantlets.

2

Materials

2.1

Plant Material

2.2

Equipment

This step-by-step protocol was developed using two different Brassica napus double haploid breeding lines: DH12075 (a low-response line widely used in different studies on Brassica breeding and genomics, as well as on DH research) and DH4079 (a high-response, model line for microspore embryogenesis in Brassica species, selected from cultivar Topas). It could be used for other B. napus lines or genotypes as long as the most suitable bud lengths for isolation of microspores at the right developmental stage (see Note 1) are previously determined. 1. Laminar flow hood. 2. Refrigerated centrifuge (see Note 2). 3. Incubators at 32
C and 25
C. 4. Growth chamber at 25
C.

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5. Inverted microscope. 6. Fluorescence microscope equipped with 358/461 nm and 480/520 nm filter combinations for DAPI and FDA observation, respectively. 7. Stereo microscope with micrometer. 8. Flow cytometer. 9. Autoclave. 10. General laboratory equipment: precision balance, pH meter, rotatory shaker, refrigerator, and so on. 2.3 Disinfection, Isolation, Culture, Ploidy Analysis, and Acclimation

1. Sterile glass bottles for liquid media. 2. Tea sieves (see Note 3). 3. Honey jars with 70% (v/v) ethanol at 4
C. 4. Honey jars (approx. 500 ml) with 4 g/l sodium hypochlorite, 0.05 (v/v) Tween at 4
C for bud disinfection. 5. 3 honey jars (approx. 500 ml) with sterile dH2O. 6. Syringe plungers kept in 70% ethanol. 7. Sterile beakers (previously wrapped in aluminum foil and autoclaved) with an internal diameter larger than that of the syringe plungers used. 8. Sterile Erlenmeyer flask with a funnel containing 30 μm-mesh nylon filters (previously wrapped in aluminum foil and autoclaved). 9. Neubauer improved counting chamber (hemacytometer) to calculate microspore plating densities (see Note 4). 10. A Bunsen-type burner, or similar, for flame sterilization. 11. Forceps. 12. Motorized or manual pipette. 13. Microscope slides and coverslips (see Note 5). 14. Micropipette (1–10 μl). 15. Sterile baby food jars (200 ml) with plastic lids. 16. Autoclavable, 90  100 mm plastic pots. 17. 30 μm-mesh filters for flow cytometry. 18. Sterile syringes. 19. Sterile 0.2 μm membrane filters. 20. Ice (see Note 6). 21. Sterile 15 ml centrifuge tubes. 22. Sterile Whatman paper. 23. Sterile 10 and 25 ml disposable pipettes. 24. Pipette tips for the micropipettes used.

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25. Sterile culture dishes (see Note 7). 26. 1.5 ml conical microtubes. 27. Parafilm. 28. 9 cm culture dishes for MDE germination. 29. Vermiculite. 30. Composite soil. 31. Transparent plastic cups. 32. Plastic tubes 3.5 ml (for flow cytometry). 33. Razor blades. 2.4 Culture Media and Chemicals

1. Isolation medium: 13% (w/v) sucrose in dH2O, pH 5.8 (see Note 8). 2. NLN-13 medium for microspore culture: NLN basal medium (Table 1; see Note 9) supplemented with 13% (w/v) sucrose and 0.5 g/l Ca(NO3)2·4H2O. Adjust pH to 5.8 and sterilize by filtration through sterile 0.2 μm membrane filters within a laminar flow hood. Store at 4
C for a maximum of 1 month. 3. 4,6-Diamidino-2-phenylindole (DAPI) stock solution: Dissolve 1 mg DAPI in 1–2 drops of DMSO. Add milliQ water up to 1 ml. 4. Nuclear Isolation Buffer (NIB): 10 mM sperminetetrahydrochloride, 10 mM NaCl, 200 mM hexylene glycol (2,5-hexanediol) in 0.01 M Tris–HCl, pH 7.0. It can be stored at 4
C for up to 1 year. 5. DAPI working solution (1.25 μg/ml) [7]: 5 ml NIB, 12.5 μl DAPI stock solution, 50 μl Triton X-100, and 2 ml glycerol. Mix the components in a beaker and adjust the total volume to 10 ml with glycerol (around 3 ml more). Cover beaker with aluminum foil and shake overnight. The DAPI working solution can then be stored at 4
C for several months. 6. Fluorescein diacetate (FDA) stock solution: 0.2% FDA in acetone (see Note 10). 7. Medium for MDE germination and conversion into plantlets: B5-Gamborg medium (Table 1) supplemented with 1% sucrose and 0.6% Plant Agar. Adjust pH to 5.8, autoclave, and pour in 90  25 mm sterile culture dishes and sterile baby food jars. 8. Lysis buffer (LB01) for flow cytometry: 5 mM Tris (hydroxymethyl) aminomethane, 2 mM Na2EDTA, 0.5 mM spermidine, 80 mM KCl, 20 mM NaCl, 15 mM β-mercaptoethanol, and 0.1% (v/v) Triton X-100. Adjust pH to 7.5. 9. Commercial DAPI-based staining buffer for flow cytometry, such as Partec CyStain UV Precise P.

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Table 1 Macroelements, microelements, and vitamins of NLN-13 and B5 media for B. napus microspore culture, based on the original formulations of NLN [11] and of B5 [12] NLN-13

B5

Ca(NO3)2

500

0

KH2PO4

125

0

KNO3

125

2500.00

MgSO4

61

121.56

NaH2PO4

0

130.44

(NH4)2SO4

0

134.00

CaCl2

0

113.23

CoCl2·6H2O

0.025

0.025

CuSO4·5H2O

0.025

0.025

FeNaEDTA

36.7

36.70

H3BO3

10

3.00

KI

0

0.75

MnSO4·H2O

18.95

10.00

Na2MoO4·2H2O

0.25

0.25

ZnSO4·7H2O

10

2.00

D(+)-biotin

0.05

0

Folic acid

0.5

0

L-glutamine

800

0

Glutathione (reduced)

30

0

Glycine

2

0

Myoinositol

100

100

Nicotinic acid

5

1

Pyridoxine–HCl

0.5

1

L-serine

100

0

Thiamine–HCl

0.5

10

Sucrose

130,000

Macroelements

Microelements

Vitamins and amino acids

Concentrations are expressed in mg/l

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Methods

3.1 Donor Plant Growth Conditions

3.2 In Vitro Culture of Isolated Microspores

Donor plants are grown in a growth chamber at 20
C, light intensity of 200 μmol/m2/s with a 16 h photoperiod and 60–65% relative humidity (see Note 11). When plants start flowering they are transferred to growth chamber with the same conditions but at 15
C to increase the difference between the temperature of normal plant growth and the heat shock-based treatment used for embryogenesis induction, which favors the embryogenic response. Soil is supplemented with long-lasting nitrogen sources for rapid, sustained growth (e.g., Nitrophoska Fertilizer). After a week at 15
C and once the first flowering has occurred, plants can be used as donors for microspore culture. It is important to remove open flowers regularly to increase flower bud production and ensure a regular supply of useful buds at the right developmental stage. 1. Switch on the flow hood, clean and disinfect the working surface, place all the materials in (except for cooled flasks and buds), and turn on the UV light for 30 min. 2. Excise inflorescences from plants (Fig. 1a) and place them in a plastic container within a box with ice (see Note 12). 3. Under a stereo microscope equipped with micrometer (Fig. 1b), sort buds with different sizes (see Note 13) and place them in separate dishes on ice until finishing this step. Once finished, place buds of each length in separate tea sieves (Fig. 1c). 4. Turn on the refrigerated centrifuge to start cooling to 4
C. 5. Take buds to the laminar flow hood to start microspore isolation. All following steps are carried out in a laminar flow hood. 6. Sterilize the selected flower buds. For this, transfer tea sieves through the different precooled jars with the different sterilizing or washing solutions, as follows: (a) 30 s in ethanol 70%. (b) 10 min in 4 g/l sodium hypochlorite while regularly shaking the sieves to remove air bubbles from the bud surface. (c) Wash three times 5 min each by dipping sieves into three different jars with cold sterile dH2O while shaking the sieves. After transfer of sieves to the next washing jar, soak the syringe plungers in the previous jar to remove ethanol (remember that plungers are stored in 70% ethanol) and have them ready for use. 7. After the last wash, remove buds from sieves by gently laying them on sterilized Whatman papers to let them dry.

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Fig. 1 Steps of the microspore culture protocol. (a) B. napus inflorescence with still unopened buds at different stages. It is important to use only the central, small buds (2–4 mm), discarding the rest that are too mature. (b) Rapeseed flower bud under the stereo microscope. Measurement of the bud is done using the millimeter scale of the micrometer and measuring the entire sepal length. (c) Tea sieve with buds, used to sterilize them with bleach while keeping them together. (d) Crushing of buds in culture media using a small beaker and a syringe plunger. (e) A 30 μm nylon filter placed in a funnel and previously sterilized by autoclaving. Erlenmeyer flask serves as a support for the funnel and conical tube used to collect the filtrate. (f) Final 1 ml suspension of microspores after the three centrifugation rounds and before resuspension in the final volume

8. Prepare a small beaker with 5 ml of isolation medium and place buds in with a sterile forceps. Gently crush buds with the washed syringe plunger (Fig. 1d) until all buds have been crushed. Rinse the plunger with 5 ml of isolation medium to maximize recovery of microspores. 9. Filter the microspore suspension through the 30 μm nylon meshes previously fitted in funnels. Collect the filtered suspension in a 15 ml tube (Fig. 1e). Put the tube on ice. The final volume should be around 10 ml in each tube. 10. Centrifuge the resulting suspension for 4 min at 4ºC and 800 rpm in a refrigerated Eppendorf Centrifuge 5804R with a 17.5 cm rotor radius. After each centrifugation round, pour the supernatant off with a fast wrist movement to avoid resuspension of the microspore pellet. 11. Add fresh isolation medium up to 10 ml and resuspend the pellet by gently shaking the tube. Repeat the centrifugationwashing rounds three times.

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Fig. 2 Neubauer Improved counting chamber. (a) Overview. The two red arrows indicate the position of the two counting cells. (b) Single counting cell. Asterisks indicate the five squares of a diagonal to be counted. See text for further details

12. After the third centrifugation, resuspend the purified microspores in 1 ml of isolation medium (Fig. 1f). 13. Calculate the density of the initial (1 ml) microspore suspension (see Note 14), and adjust it to 40,000 microspores/ml (the optimal density established for B. napus microspore cultures [8]), using a hemocytometer counting chamber (Fig. 2). 14. Check microspore viability by FDA staining and, if there are doubts about the developmental stage, double check it by DAPI staining (see Note 15). 15. Pour 1, 3, or 10 ml of the final microspore suspension in 3 cm Ø, 6 cm Ø, or 9 cm Ø culture plates, respectively. 16. Seal with Parafilm (double or better triple-seal). Keep plates in darkness (see Note 16) at 32
C for heat shock treatment (see Note 17). 17. Upon finishing the treatment, keep dishes in darkness at 25
C (see Note 18). To check the efficiency of heat shock, extract a sample from a culture dish 2–6 days after microspore isolation, stain with DAPI (see Note 19), and check for dividing microspores, that is, microspores with more than one DAPI-stained nuclei within the same exine coat (Fig. 3a, b). 18. Two or three weeks after isolation, MDEs are visible under a stereo microscope at low magnification or even with the naked eye (Fig. 3c, d). Final efficiency of the culture is calculated counting the total of MDEs under a stereo microscope (Fig. 3d, see Note 20). 3.3 MDE Germination and Acclimation

1. Pick up MDEs at the cotyledonary stage and place them in solid B5 medium, previously poured into 9 cm Ø high dishes, for germination. Inoculate a maximum of 10 MDEs per dish to allow them to grow without interferences. Germinate them at 25
C and 32 μmol/m2/s light intensity, 12 h photoperiod.

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Fig. 3 Different stages during B. napus microspore culture. (a, b) Dividing, embryogenic microspore as seen by phase contrast (a) and DAPI staining (b). (c) Torpedo-like MDE. (d) Culture dish with MDEs mostly at the torpedo–cotyledonary stages. Bars: (a, b) 10 μm; (c) 100 μm

2. Transfer germinated seedlings to sterile glass small jars with 30 ml of B5 medium for further growth (see Note 21). 3. Properly developed seedlings can be transferred to plastic pots with wet composite soil. 4. Acclimate plantlets in a growth chamber at 25
C and 16 h photoperiod (see Note 22). 3.4 Analysis of Ploidy Level

Flow cytometry is used to check DNA content of microsporederived plantlets and, by comparison with a known diploid donor plant, infer their ploidy level. Thus, diploid donor plants should be analyzed in parallel as controls for 2C DNA content. 1. Cut out young leafs from plants (see Note 23) and place them in dishes inside a box with ice. 2. Add 0.5 ml of lysis buffer on the top of the leaf.

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3. Cut a leaf piece of approximately 1 cm2, and chop it into fine chunks using a razor blade, to release as much intact nuclei as possible to the lysis buffer. 4. Filter the homogenate through to a 30 μm filter and collect the filtered nuclei into vials compatible with the loading port of the flow cytometer to be used (see Note 24). Add 1.5 ml of DAPI staining buffer. 5. Incubate vials for 2 min on ice before analyzing the samples. 6. Start the analysis with the flow cytometer according to manufacturer specifications. Perform a minimum of 10,000 counts per sample.

4

Notes 1. One of the most important parameters to guarantee successful microspore embryogenesis is the proper identification of the most suitable developmental stages of microspores, which in the case of B. napus revolves around the vacuolated microspore and young bicellular pollen stages [4]. For this, it is typical to isolate microspores/pollen from different previously measured bud sizes, and then stain them with DAPI and observe their developmental stage under a fluorescence microscope in order to identify the bud length range where the percentages of vacuolated microspores and young bicellular pollen grains are highest. 2. In our case, we use an Eppendorf® Centrifuge 5804R, refrigerated, with A-4-44 rotor (17.5 cm radius) with adapters for 15/50 ml conical tubes. 3. The number of tea sieves, beakers, and Erlenmeyer flask depends on the number of bud length intervals to be used in parallel (see Note 13). 4. Plating of microspores at the right density is a crucial step for a good embryogenic response. Different counting methods can be used to calculate microspore plating densities, depending on the equipment available in each laboratory. Among them, however, hemocytometers and in particular the Neubauer Improved chamber appears to be the simplest and most suitable approach [9]. 5. For observation of samples stained with DAPI and FDA, slides and coverslips can be reused for a limited number of times, provided that they are carefully cleaned and stored to avoid scratches and dust deposition. All the waste containing DAPI, including microscope slides, should be stored and disposed of following the corresponding safety rules to avoid contamination of the working area.

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6. Once buds are excised from donor plants, it is essential to keep them on ice to arrest microspore development and slow down the processes of cell death. If the workplace is cool enough, keeping the buds on ice and using cold (4
C) disinfecting solutions should be sufficient to prevent loss of embryogenic response. If not, additional measures should be taken. For example, we found that keeping both the solutions and the buds on ice during the whole process of microspore isolation, filtration and plating allows for a successful microspore culture, with good embryogenic response, even in warm workspaces, where air cooling is insufficient or absent. 7. In general, the choice of the dish size depends on the goal of the experiment to be performed. If the goal is to obtain mature MDEs or to collect samples at different days of culture, we usually opt for 6 cm Ø plates. If the goal is different, for example to test the effect on embryogenic response of different compounds, we use 3 cm Ø plates with less culture volume, to optimize the number of chemicals tested in a single experiment. However, it must be noted that dish size has been reported to have an effect on the embryogenic response in other species [10]. 8. For microspore isolation and washing it is possible to use exactly the same NLN-13-based medium used for microspore culture. This would be the easiest and most straightforward alternative. However, preparation of NLN-13 medium is expensive and time-consuming. As an alternative to this, we use the isolation medium. This medium has almost the same osmotic conditions (mainly derived from sucrose) and the same pH value as NLN-13. However, since we only need it for isolation and washing (two relatively rapid steps), there is no need for filtering. It can be autoclaved, which allows for time and money savings, while the efficiency of microspore embryogenesis is not compromised. Once prepared, it must be stored at 4
C for a maximum of 1 month. 9. NLN basal medium (salts and vitamins) can be prepared from stocks (Table 1) or bought as commercial preparation. 10. Whereas DAPI working solution can be prepared in advance from stock solution, FDA working solution must be prepared immediately before use. This is why it will be described later in the Methods section in the context of viability determination. 11. It is also possible to keep plants outdoors instead of indoors. The main advantage is that plants grow more, faster, and produce a larger number of buds to work with, which is highly recommended when performing large-scale cultures. The limitation is that their optimal growth and performance greatly depend on climatic conditions, especially temperature changes.

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This limits the optimal outdoor growth to the months when temperatures do not exceed 20
C during light hours and decrease at night. In addition, outdoor growth exposes them to more frequent pest attacks. To avoid these limitations, it is advisable to keep them in growth chambers under controlled conditions. 12. The number of inflorescences to use depends on the scale of the culture to be carried out. It is better to cut entire inflorescences than individual buds, since this way buds will last longer alive and healthy. 13. As the correlation between bud length and microspore/pollen developmental stage is usually influenced by environmental and donor plant conditions, three consecutive bud length intervals are normally processed in parallel to ensure that at least one of them contains the highest percentage of microspores/pollen at the right developmental stage. For DH4079 and DH12075 lines, the intervals typically used are 3–3.2/ 3.2–3.4/3.4–3.6, and 2.4–2.6/2.6–2.8/2.8–3.0 mm, respectively. 14. Take 40 μl of the initial 1 ml suspension. Pipet 10 μl in each of the two counting cells of the Neubauer chamber (Fig. 2a), leaving the remaining 20 μl in the tube to be used in the next step. Pipet each 10 μl drop on the edge of the chamber coverslip and the suspension will be sucked into the chamber by capillarity. With a microscope, count the microspores on each of the five squares of a single diagonal (Fig. 2b). Repeat this step with the second 10 μl drop, deposited in the other counting cell. Counting is considered coherent when both counts do not deviate more than 15%. If they do, repeat counts with two new drops. With two coherent counts, the average is calculated and then used to estimate the final volume to which resuspend microspores for a final density of 40,000 microspores/ml, according to the following formula: Final volume ¼

Average count  5 4

15. For DAPI and FDA staining, use the 20 μl volume remaining from the 40 μl taken from the initial microspore suspension (see Note 14). For DAPI staining, mix 10 μl of DAPI working solution with 10 μl of the initial microspore suspension. Incubate 15 min in darkness, mount on a slide, add a coverslip, and observe with the fluorescence microscope. It is possible to keep the preparation at 4
C in darkness to be observed later on. For FDA staining, prepare a 1/100 working solution of FDA stock solution in dH2O immediately prior to use. Add 5 μl of the working solution to the remaining 10 μl of the initial

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microspore suspension. Incubate 10 min at room temperature in darkness, mount on a slide, add a coverslip, and observe with fluorescence microscope. 16. To better ensure darkness, it is advisable to wrap dishes in aluminum foil or to place dishes in opaque cardboard or plastic boxes. 17. The duration of the heat stress treatment will affect the process in two ways: efficiency of embryogenesis induction and MDE quality. Prolonged treatments produce more MDEs, but they are more prone to present morphological abnormalities. Conversely, shorter treatments produce less MDEs, but of better quality. It also depends on its combined use with other abiotic stresses or chemicals that enhance embryogenesis induction. This parameter is genotype-dependent and should therefore be established for each new genotype. In such a case, our advice would be to perform an initial test applying 32
C treatments of 24, 48, and 72 h, and then opt for the duration that provides the best compromise between high response and good MDE quality. 18. Depending on the goal of the experiment, it may be necessary to refresh culture medium during 25
C incubation. If the goal is to study early stages of embryogenesis, up to development of globular MDEs, the medium initially poured may be enough. If the goal is to study advanced stages of MDE development or to produce DH plants, it may be necessary to add fresh NLN13 medium 7 days after culture initiation to compensate the loss by consumption and evaporation. This will allow for the development of high-quality MDEs. 19. The procedure of DAPI staining to check for microspore divisions is the same as for stage determination (see Note 15), but using 100 μl of cell suspension taken under sterile conditions (in the flow hood). To avoid culture contamination, it is convenient to have a 3 cm Ø extra dish of each treatment to use for this purpose. 20. Efficiency is usually expressed as the number of MDEs per petri dish, per ml of culture media (dividing by the original volume of each dish counted) or per bud (using the relation of buds per ml, which must be calculated in each culture). The choice depends on the purpose of the study and the intended use of the obtained data. 21. Transfer of germinated seedlings from 9 cm Ø dishes to sterile glass jars provides the space needed for a proper development of roots and shoots. 22. Upon transfer of plantlets to pots with soil, humidity conditions change dramatically, which may severely affect plant growth. To prevent this, plantlets should be first covered with

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intact plastic cups, pinching new holes on the cup every 2 days. This way, holes would progressively increase transpiration and would acclimatize plants to environmental humidity. After several days, remove the cup. 23. The use of young tissues for ploidy analysis relates to the fact that young, proliferating tissues have more cells in the G2 stage of the cell cycle. Thus, the G2 peak will be more clearly seen in the histogram. 24. In our case, 3.5 ml plastic vials compatible with a Partec CyFlow Ploidy Analyzer are used.

Acknowledgments This work was supported by grant AGL2017-88135-R to J.M.S.S. from MINECO jointly funded by FEDER. References 1. Seguı´-Simarro JM, Nuez F (2008) Pathways to doubled haploidy: chromosome doubling during androgenesis. Cytogenet Genome Res 120 (3–4):358–369. https://doi.org/10.1159/ 000121085 2. Seguı´-Simarro JM (2010) Androgenesis revisited. Bot Rev 76(3):377–404. https://doi. org/10.1007/s12229-010-9056-6 3. Shariatpanahi ME, Bal U, Heberle-Bors E, Touraev A (2006) Stresses applied for the re-programming of plant microspores towards in vitro embryogenesis. Physiol Plant 127 (4):519–534 4. Seguı´-Simarro JM, Nuez F (2008) How microspores transform into haploid embryos: changes associated with embryogenesis induction and microspore-derived embryogenesis. Physiol Plant 134:1–12. https://doi.org/10. 1111/j.1399-3054.2008.01113.x 5. Forster BP, Heberle-Bors E, Kasha KJ, Touraev A (2007) The resurgence of haploids in higher plants. Trends Plant Sci 12(8):368–375. https://doi.org/10.1016/j.tplants.2007.06. 007 6. Dwivedi SL, Britt AB, Tripathi L, Sharma S, Upadhyaya HD, Ortiz R (2015) Haploids: constraints and opportunities in plant breeding. Biotechnol Adv 33(6 Pt 1):812–829. https://doi.org/10.1016/j.biotechadv.2015. 07.001

7. Custers J (2003) Microspore culture in rapeseed (Brassica napus L.). In: Maluszynski M, Kasha KJ, Forster BP, Szarejko I (eds) Doubled haploid production in crop plants. Kluwer Academic Publishers, Dordrecht, pp 185–193 8. Huang B, Bird S, Kemble R, Simmonds D, Keller W, Miki B (1990) Effects of culture density, conditioned medium and feeder cultures on microspore embryogenesis in Brassica napus L. cv. Topas. Plant Cell Rep 8 (10):594–597 9. Camacho-Ferna´ndez C, Herva´s D, RivasSendra A, Marı´n MP, Seguı´-Simarro JM (2018) Comparison of six different methods to calculate cell densities. Plant Methods 14 (1):30. https://doi.org/10.1186/s13007018-0297-4 10. Kim M, Park E-J, An D, Lee Y (2013) Highquality embryo production and plant regeneration using a two-step culture system in isolated microspore cultures of hot pepper (Capsicum annuum L.). Plant Cell Tissue Organ Cult 112 (2):191–201. https://doi.org/10.1007/ s11240-012-0222-x 11. Nitsch JP (1972) Haploid plants from pollen. Z Pflanzenzu¨cht 67:3–18 12. Gamborg OL, Miller RA, Ojima K (1968) Nutrient requirements of suspension cultures of soybean root cells. Exp Cell Res 50 (1):151–158. https://doi.org/10.1016/ 0014-4827(68)90403-5

Chapter 20 Anther Culture in Eggplant (Solanum melongena L.) Antonio Calabuig-Serna, Rosa Porcel, Patricia Corral-Martı´nez, and Jose M. Seguı´-Simarro Abstract For a long time, conventional breeding methods have been used to obtain pure, 100% homozygous lines for hybrid seed production in crops of agronomic interest. However, by doubled haploid technology, it is possible to produce 100% homozygous plants derived from precursors of male gametophytes (androgenesis), to accelerate the production of pure lines, which implies important time and cost savings. In this chapter, a protocol for anther culture in eggplant is described, from donor plant growth conditions to regeneration and acclimation of doubled haploid plants, as well as a description of how to analyze ploidy levels of regenerated plants. Key words Androgenesis, Doubled haploid, Haploid, Embryogenesis, Tissue culture

1

Introduction Based on FAOSTAT data from 2016 [1], the global production of eggplant is around 50 million tons annually, with a net value of more than US$10 billion a year, which makes it the fifth most economically important solanaceous crop after potato (Solanum tuberosum), tomato (Solanum lycopersicum), pepper (Capsicum annum), and tobacco (Nicotiana tabacum). In most of these crops, maximum yields are produced by hybrid plants created by the crossing of pure lines with the desired traits. These homozygous lines can be produced by conventional breeding techniques, increasing homozygosity in successive rounds of inbreeding and selection through 6–10 generations. This approach is straightforward and can be successfully applied to virtually all crops. However, it is both time consuming (several years of work) and costly. Alternatively, plant biotechnology allows for the acceleration of the process of homozygous line production by creating doubled haploid (DH) plants derived from (haploid) male gametophyte precursors. This experimental approach, commonly known as androgenesis, has several ways to be induced. The

Martin Bayer (ed.), Plant Embryogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2122, https://doi.org/10.1007/978-1-0716-0342-0_20, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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most successful and widely used is microspore embryogenesis, which consists in the deviation of microspores/young pollen grains from their original gametophytic fate towards embryogenesis. Technically, this can be achieved through in vitro anther culture, whereby DH plants are produced in vitro, and then acclimatized and grown to maturity to finally obtain the desired, fully homozygous pure line. This reduces pure line creation to only one in vitro generation. As a consequence of this, the time and costs of the whole process may be dramatically reduced. The first haploids from anther culture in eggplant were obtained almost simultaneously by the Chinese Research Group of Haploid Breeding [2] and by Isouard et al. [3]. Since then, methods for obtaining DHs through anther culture have been modified, adapted and improved by different research groups [4– 8]. Currently, DHs are typically produced through anther culture for most eggplant varieties of agronomic interest. It is a wellestablished, quick, and easy method for obtaining pure lines. However, it has several limitations including the uncontrolled contribution of anther wall cells to culture conditions, the general low efficiency of the process, and the possibility of occurrence of somatic regenerants derived from anther walls, which makes mandatory the genetic analysis of every single regenerant obtained in order to discard those coming from anther wall tissues and therefore non-DH [9, 10]. In this chapter, a step-by-step protocol for anther culture in eggplant is presented. Every step from donor plant growth conditions to the regeneration and acclimatization of callus-derived DH plants is detailed, together with a description of the ploidy analysis needed to check for the haploid origin of regenerated plants.

2

Materials

2.1

Plant Material

2.2

Equipment

We used as donors, greenhouse-grown plants of the DH36 line—a high androgenic response line generated by us and derived from cv. Bandera [11]. However, the protocols described hereby can be used for other eggplant genotypes (see Note 1). 1. Laminar flow hood. 2. Inverted light microscope. 3. Calipers. 4. Incubator at 35  C and 25  C. 5. Autoclave. 6. Flow cytometer (Partec CyFlow Ploidy Analyzer in our case, or similar device).

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7. Plant growth chamber at 25  C. 8. General laboratory equipment: precision balance, pH meter, stirrer, refrigerator, and so on. 2.3

Materials

1. Box for melting ice. 2. Sterile forceps and scalpel. 3. Sterile glass bottles for liquid media. 4. Sterile baby food jars with plastic lids. 5. Plastic plant pots 90  100 mm (width  height). 6. Sterile Whatman paper. 7. Sterile 50 ml plastic tubes. 8. Sterile culture dishes 90  25 mm (Ø  height). 9. Parafilm. 10. Microscope slides and coverslips. 11. Aluminum foil. 12. Composite soil. 13. Transparent plastic cups. 14. Razor blades. 15. Filters with 30 μm pore size. 16. Plastic tubes 3.5 ml, 55  12 mm (Ø  height).

2.4 Solutions for Anther Culture

1. 70% ethanol (v/v). 2. 4 g/l sodium hypochlorite with 0.05% Tween (v/v). 3. Sterile dH2O (three glass jars) autoclaved at 121  C for 20 min and cooled before use. 4. 4,6-Diamidino-2-phenylindole (DAPI) stock solution: Dissolve 1 mg DAPI in 1–2 drops (~10–20 μl) of DMSO. Add milliQ water up to 1 ml. 5. Nuclear Isolation Buffer (NIB): Prepare a solution of 10 mM spermine-tetrahydrochloride, 10 mM NaCl and 200 mM hexylene glycol (2,5-hexanediol) in Tris–HCl buffer 0.01 M, pH 7. It can be stored at 4  C for up to 1 year. 6. DAPI working solution (1.25 μg/ml) [12]: Briefly, mix well the following components in a small beaker: 5 ml NIB, 12.5 μl DAPI stock solution, 50 μl Triton X-100, and 2 ml glycerol. Fill up to 10 ml with glycerol (around 3 ml more). Cover beaker with aluminum foil and shake overnight. Next day, aliquots of the DAPI working solution can be stored at 4  C for several months. 7. Fluorescein diacetate (FDA) stock solution: 0.2% FDA in acetone (see Note 2).

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Table 1 Macroelements, microelements, and vitamins used in the three basal media utilized for eggplant anther culture (mg/l) C

R

V3

KNO3

2150

2150

1900

NH4NO3

1238

1238

1650

MgSO4·7H2O

412

412

370

MgSO4

–

–

–

CaCl2·2H2O

313

313

440

CaCl2

–

–

–

KH2PO4

142

142

170

Ca(NO3)2·4H2O

50

50

–

Ca(NO3)2

–

–

–

NaH2PO4·H2O

38

38

–

(NH4)2SO4

34

34

–

KCl

7

7

–

MnSO4·H2O

22.130

20.130

0.076

ZnSO4·7H2O

3.625

3.225

1

H3BO3

3.150

1.550

1

KI

0.695

0.330

0.01

Na2MoO4·2H2O

0.188

0.138

–

CuSO4·5H2O

0.016

0.011

0.03

CoCl2·6H2O

0.016

0.011

–

AlCl3·6H2O

–

–

0.05

NiCl2·6H2O

–

–

0.03

Myoinositol

100.00

100.00

100.00

Pyridoxine–HCl

5.500

5.500

5.500

Nicotinic acid

0.700

0.700

0.700

Thiamine–HCl

0.600

0.600

0.600

Calcium pantothenate

0.500

0.500

0.500

Biotin

0.005

0.005

0.005

Glycine

0.100

0.100

0.200

Macroelements

Microelements

Vitamins and amino acids

(continued)

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Table 1 (continued) C

R

V3

Folic acid

–

–

–

L-Glutamine

–

–

–

Glutathione

–

–

–

L-Serine

–

–

–

Na2 EDTA

18.65

18.65

37.30

FeNaEDTA

–

–

–

FeSO4·7H2O

13.90

13.90

27.28

Sucrose

120,000

30,000

30,000

Bacto agar

8000

8000

8000

Chelated iron

8. Lysis buffer (LB01) for flow cytometry: 5 mM Tris (hydroxymethyl) aminomethane, 2 mM Na2EDTA, 0.5 mM spermidine 80 mM KCl, 20 mM NaCl, 15 mM β-mercaptoethanol, and 0.1% (v/v) Triton X-100. Adjust pH to 7.5. 9. Commercial DAPI-based staining buffer Partec CyStain UV Precise P for flow cytometry (Partec GmbH, Mu¨nster, Germany). 2.5

Culture Media

1. Induction medium: C medium (Table 1) supplemented with 5 mg/l kinetin, 5 mg/l 2,4-dichlorophenoxyacetic acid (2,4-D), and 0.2 mg/l B12 Vitamin. Prepare the medium first, then autoclave it (121  C, 20 min), and then add hormones and B12 vitamin from filter-sterilized stocks. Adjust pH to 5.9. 2. Regeneration medium: R medium (Table 1), supplemented with 0.1 mg/l kinetin. Prepare the medium first, then autoclave it, and then add kinetin from filter-sterilized stock. Adjust pH to 5.9 and pour in 90  25 mm sterile culture dishes and sterile baby food jars (200 ml). 3. Rooting medium: V3 medium (Table 1). Adjust pH to 5.9. Autoclave medium at 121  C for 20 min and pour it in 90  25 mm sterile culture dishes and sterile baby food jars (200 ml).

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Methods

3.1 Donor Plant Growth Conditions

1. Sow eggplant seeds and grow seedlings in growth chambers at 20  C, light intensity of 200 μmol/m2/s with 16 h photoperiod and 60–65% relative humidity. 2. As soon as they are 10–15 cm in eight, they can be moved to greenhouse, at 20–25  C under natural light. 3. Upon flowering, they can be used as donor plants for microspore and anther culture. In general, the use of healthy and vigorous plants, with no open flowers or fruits, increases the odds of obtaining a good androgenic response (see Note 3).

3.2 In Vitro Culture of Anthers

1. Select the buds for anther culture (Fig. 1a). Although the precise bud length that contains anthers with microspores/ pollen at the suitable stages should be determined for each genotype [7], in general, optimal buds are totally closed, with petals fully covered by sepals. Excise buds from donor plants and bring them to the lab in plastic tubes on ice. 2. Take the buds to the laminar flow hood and surface disinfect by immersing them in 70% ethanol for 30 s, removing the solution and adding 4 g/l sodium hypochlorite with 0.05% Tween for 5 min (Fig. 1b). Finally, buds are rinsed three times (4 min each) with sterile dH2O (see Note 4). 3. Place buds over a sterile Whatman filter paper to extract the anthers. Excise the basal part of the bud with the scalpel. Then, perform a longitudinal cut to remove both calyx and corolla (Fig. 1c, d). It is very important to avoid damaging the anthers during this operation. 4. With the caliper, measure one excised anther from each bud to determine the precise stage of microspores/pollen (see Note 5). If the anther is at the right stage, discard it (since it has been in contact with the caliper) and use the other five for culture. 5. Remove anther filaments (see Note 6) and place anthers in 90 mm culture dishes (see Note 7) containing solidified induction medium. Inoculate around 20 anthers per dish (Fig. 1e). 6. Seal culture dishes with Parafilm and keep them in darkness at 35  C for 8 days (see Note 8). 7. At day 8, transfer culture dishes to light (12/12 photoperiod) and 25  C. Keep plates in these conditions for 4 days. 8. At day 12, transfer anthers to new culture dishes containing regeneration medium. Keep them in these conditions, with 12/12 light photoperiod and 25  C, until embryos are seen to pop out of anthers (Fig. 1f).

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Fig. 1 Eggplant anther culture. (a) Buds of different size. The arrow points to the bud size that contains microspores/pollen at the most suitable stage to induce embryogenesis. (b) Buds being surface disinfected in a 50 ml tube. (c) Intact bud where the transversal and longitudinal cuts to be made for dissection are marked by the 1 and 2 dashed lines, respectively. (d) Dissected bud showing anther disposition. (e) Anthers inoculated in a culture dish, ready for culture. (f) Cultured anthers with embryos (arrows) popping out of anthers. In the case of the leftmost anther, the embryo is germinating and producing leaves still within the anther. (g) Regenerated plantlet during its process of acclimation from in vitro to ex vitro conditions

9. Transfer developed embryos to solidified germination medium in culture dishes or baby food jars, depending on the developmental stage of the embryo (see Note 9). Keep them growing

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until germination into plantlets. If plantlets need more space, transfer them to larger containers with the same medium composition. 10. As soon as they develop 4–5 leaves and roots, transfer them to pots with soil, cover them with a plastic cup (see Note 10), and acclimatize in growth chambers (Fig. 1g). 11. Three months after the initiation of in vitro culture, transfer responsive anthers to new plates with fresh medium, in order to promote growth of new embryos. Use them as long as they produce embryos. Discard anthers with no response. 3.3 Analysis of Ploidy Level

Microspore-derived plants are analyzed by flow cytometry to check their DNA contents and, by comparison with a known diploid individual, infer their ploidy level. Thus, in parallel to microsporederived plants, analyze also diploid donor plants as control for the 2C DNA content to compare with. 1. Cut out young leafs (see Note 11) from plants and place them in dishes inside a box with ice. Add 0.5 ml of lysis buffer on the top of the leaf. 2. Cut a leaf piece of approximately 1 cm2, and chop it into fine chunks using a razor blade, to release as much intact nuclei as possible to the lysis buffer. 3. Filter the homogenate through a 30 μm filter and collect the filtered nuclei into vials compatible with the loading port of the flow cytometer to be used (see Note 12). Add 1.5 ml of DAPI staining buffer. 4. Incubate vials for 2 min on ice before analyzing the samples. 5. Start the analysis with the flow cytometer according to manufacturer specifications. Perform a minimum of 10,000 counts per sample.

4

Notes 1. We have also used this protocol with other eggplant cultivars and genotypes such as DH44 [11], Bandera (an F1 hybrid from Seminis Vegetable Seeds Iberica, S.A., Spain), Ecavi (an F1 hybrid from Rijk Zwaan Ibe´rica S.A., Spain), and other different eggplant proprietary materials from several private companies. This protocol can be used with other genotypes of eggplant provided that the most suitable bud size has been previously established. The microspore and anther developmental stage is a key factor to succeed in the process of embryogenesis induction.

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2. Whereas DAPI working solution can be prepared in advance from stock solution, FDA working solution must be prepared immediately before use. This is why it will be described later, in Subheading 3, while describing viability determination. 3. Unfortunately, the optimal environmental conditions that allow to maximize the androgenic response of donor plants are not fully understood. However, as in other crops, a good health and absence of biotic or abiotic stress make anthers and microspores more prone to respond to embryogenesis. Eggplant crops use to grow well under warm conditions and long photoperiods with intense sunlight, as in, for example, springsummer in Mediterranean countries. However, excessively high temperatures may affect microspore/pollen viability and reduce androgenic response. Thus, it is advisable to avoid anther or microspore culture during the hot summer months, at least when using greenhouse-grown plants in warm–hot climates. We have also observed that pest attacks reduce the response of anther and microspore cultures. On the other hand, strong or prolonged phytosanitary treatments to prevent or fight against pests may also have a detrimental effect, by reducing the androgenic response, the number of available flowers, or both. In addition, the presence of open flowers and maturing fruits decreases the occurrence of new flowers. This is why it is important to remove open flowers and small fruits from donor plants on a regular basis. 4. Glass jars containing sterilizing solutions and sterile dH2O for washings must be kept at 4  C in order to avoid damage of plant tissue. Buds must be kept on ice until culture begins. During the whole anther or microspore isolation process, solutions and dH2O must be cold. Another important issue to consider is to keep room temperature around 25  C or below, in order to preserve microspores from an extra heat stress that could reduce their androgenic response. If room temperature cannot be kept at 25  C or below, a solution is to work, always inside the laminar flow hood, on a tray filled with melting ice and covered with an aluminum plate to dissipate heat, keeping all vessels containing liquids and tissues in contact with the aluminum plate in order to assure that plant material is kept always cool prior to the application of the heat shock treatment. 5. The developmental stage of the microspores is one of the most important factors affecting androgenic response. The responding stages to induce embryogenesis are vacuolated microspores and young pollen grains. However, the relationship between the length of the anther and the developmental stage of microspores depends on the genotype, so it must be previously determined for each case. Avoid including the anther filament when measuring the anther length.

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6. Usually, filament tissues are very prone to proliferate, producing calli when cultured in in vitro conditions. Then, it is important to remove anther filament as much as possible to avoid the development of calli of somatic origin. 7. Anthers should have the concave part (internal) in contact with culture medium when placed on culture dishes. 8. To maintain darkness in incubators, culture dishes can be wrapped with aluminum foil or placed inside opaque boxes. 9. Embryos may start developing around day 30 of culture, although this time may vary depending on the genotype and culture conditions. It is important to supervise anther cultures every week in order to detect possible contaminations that could affect developing embryos. Well-formed embryos should be transferred to V3 medium as soon as they are found. 10. Once in pots with soil, plantlets are covered with a plastic cup in order to promote progressive acclimatization by pinching small holes in the plastic cup every week. 11. Young, proliferating tissues have more cells in the G2 stage of the cell cycle. Thus, the G2 peak will be more clearly seen in the histogram. 12. In our case, 3.5 ml plastic vials compatible with a Partec CyFlow Ploidy Analyzer.

Acknowledgments This work was supported by grant AGL2017-88135-R to J.M.S.S. from MINECO jointly funded by FEDER. References 1. FAOSTAT (2018). http://faostat.fao.org. Accessed July 2018 2. Research Group of Haploid Breeding (1978) Induction of haploid plants of Solanum melongena. In: Proceedings of the symposium on plant tissue culture. Peking, Science Press, pp 227–232 3. Isouard G, Raquin C, Demarly Y (1979) Obtention de plantes haploides et diploides par culture in vitro d’anthe`res d’aubergine (Solanum melongena L.). C.R. Acad. Sci. Paris Se´rie D 288:987–989 4. Dumas de Vaulx R, Chambonnet D (1982) Culture in vitro d’anthe`res d’aubergine (Solanum melongena L.): stimulation de la production de plantes au moyen de traitements a` 35  C associe´s a` de faibles teneurs en

substances de croissance. Agronomie 2 (10):983–988 5. Chambonnet D (1988) Production of haploid eggplant plants. In: Bulletin interne de la Station d’Ame´lioration des Plantes Maraiche`res d’Avignon, Montfavet, France, pp 1–10 6. Salas P, Prohens J, Seguı´-Simarro JM (2011) Evaluation of androgenic competence through anther culture in common eggplant and related species. Euphytica 182(2):261–274. https:// doi.org/10.1007/s10681-011-0490-2 7. Salas P, Rivas-Sendra A, Prohens J, Segu´ı-Simarro JM (2012) Influence of the stage for anther excision and heterostyly in embryogenesis induction from eggplant anther cultures. Euphytica 184(2):235–250. https:// doi.org/10.1007/s10681-011-0569-9

Anther Culture in Eggplant 8. Rotino GL (2016) Anther culture in eggplant (Solanum melongena L.). In: Germana MA, Lambardi M (eds) In vitro embryogenesis in higher plants, vol. 1359. Methods in molecular biology. Springer, New York, pp 453–466. https://doi.org/10.1007/978-1-4939-30616_25 9. Seguı´-Simarro JM, Corral-Martı´nez P, ParraVega V, Gonza´lez-Garcı´a B (2011) Androgenesis in recalcitrant solanaceous crops. Plant Cell Rep 30(5):765–778. https://doi.org/10. 1007/s00299-010-0984-8 10. Seguı´-Simarro JM (2016) Androgenesis in solanaceae. In: Germana` MA, Lambardi M (eds) In vitro embryogenesis, vol. 1359. Methods in molecular biology. Springer Science +

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Business Media, New York, pp 209–244. https://doi.org/10.1007/978-1-4939-30616_9 11. Rivas-Sendra A, Campos-Vega M, CalabuigSerna A, Seguı´-Simarro JM (2017) Development and characterization of an eggplant (Solanum melongena) doubled haploid population and a doubled haploid line with high androgenic response. Euphytica 213(4):89. https:// doi.org/10.1007/s10681-017-1879-3 12. Custers J (2003) Microspore culture in rapeseed (Brassica napus L.). In: Maluszynski M, Kasha KJ, Forster BP, Szarejko I (eds) Doubled haploid production in crop plants. Kluwer Academic Publishers, Dordrecht, pp 185–193

Correction to: Small RNA In Situ Hybridizations on Sections of Arabidopsis Embryos Katalin Pa´ldi, Magdalena Mosiolek, and Michael D. Nodine

Correction to: Chapter 7 in: Martin Bayer (ed.), Plant Embryogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2122, https://doi.org/10.1007/978-1-0716-0342-0_7

Chapter 7 “Small RNA In Situ Hybridizations on Sections of Arabidopsis Embryos” was previously published non-open access. It has now been changed to open access under a CC BY 4.0 license and the copyright holder has been updated to “The Author(s).” This book has also been updated with this change.

The updated online version of this chapter can be found at: https://doi.org/10.1007/978-1-0716-0342-0_7 Martin Bayer (ed.), Plant Embryogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2122, https://doi.org/10.1007/978-1-0716-0342-0_21, © Springer Science+Business Media, LLC, part of Springer Nature 2020

C1

INDEX A Acridine orange staining ............................. 226, 229, 233 Acriflavine .....................................................196, 198–200 Actin.................................................................... 38, 39, 50 Agrobacterium tumefaciens......................... 245, 248, 252 Androgenesis ........................................................ 269, 283 Anthers ............................................................28, 29, 104, 145, 210, 217, 260–262, 270, 283–292 Anthesis ......................................................................... 104 Arabidopsis thaliana ............. 49, 64, 127, 142, 151, 241

B Biotin ................................. 142, 143, 145, 148, 260, 286 Blocking reagent ................................................ 89, 90, 94 Boechera ......................................128, 130, 133, 135, 136 Brassica napus....................................................... 269–282 BSA .......................................................... 89, 95, 229, 234 Bulk-rupture method..........................113, 116, 117, 124

C cDNA construction....................................................... 114 Cell Atlas.......................................................72, 76–80, 82 Cellulase................................................................ 102, 103 Cell viability .......................................................... 105, 109 Cell volume .................................... 63, 72–75, 77, 81, 82 Chloral hydrate............................................ 6, 65, 68, 254 Clearing ..................................................6, 88, 91, 92, 96, 97, 193, 194, 196–200, 202, 213, 229, 254 Colorimetric reaction......................................... 88, 95, 98 Columella ........................................................................ 76 Cone collection ...................................225, 227, 230, 234 Confocal microscopy ........................................45, 50, 51, 57, 69, 102, 105, 246, 248 Coverslip ........................................................6, 51, 55–61, 68, 69, 78, 79, 94, 95, 97, 172, 195, 200, 202, 229, 232–234, 259–263, 271, 278, 280, 281, 285 Cytoskeleton..............................................................38, 50

D Data processing ............................................................. 168 Defective kernel (dek)......................................4, 9, 26, 31 Denhardt’s solution ..................................................89, 93 DEPC-treated water ................................... 88, 89, 92, 95 Dextran sulfate ................................................... 89, 93, 97

4,6-Diamidino-2-phenylindole (DAPI) ....................... 69, 143, 144, 147, 192, 236, 271, 272, 276–281, 285, 289, 291 DIG Nucleic Acid Detection Kit .......................... 90, 223 DNA fragmentation ................... 223, 225–227, 233, 234 Doubled haploid (DH)..............269, 270, 281, 283, 284

E Ear ............................9–11, 25, 26, 28–31, 193, 195–199 Egg cells......................................102, 258, 260, 261, 265 Eggplant ............................................................... 283–292 Emasculation ............................................... 104, 109, 148 Embedding ................................................... 6, 37, 88, 90, 91, 96, 97, 101, 129–132, 135, 141, 152, 162, 192, 206, 209, 213, 218, 220, 221, 226, 246, 248, 249, 257 emb mutants ...................................................................... 4 Embryo development .................................104, 105, 142, 148, 184, 191, 205–207, 211, 217, 224, 241–254 general counterstaining solution (EGC) ................. 51 general mounting solution (EGM).......................... 51 isolation ...................................................50, 104–107, 114–117, 170, 184 microtubule counterstaining solution (EMTC) ............................................................... 51 microtubule mounting solution (EMTM) .............................................................. 51 morphology ....................................... 5, 244, 247, 252 proper ....................................................109, 205–207, 211, 212, 216, 218–220 sac.............................................................44, 105–107, 109, 115, 169, 191–202, 257 Embryogenesis ...........................................................5, 37, 49–61, 101, 102, 127, 141, 151, 168, 192, 205–221, 223–236, 241–244, 257, 258, 269, 270, 274, 278, 279, 281, 284, 289, 291 Embryogenic cell line .......................................... 245, 248 EMS treatment .................................................... 7, 15, 16, 18, 22, 25, 26, 29, 31 Endosperm ....................................................3–11, 26, 29, 31, 43, 50, 101, 110, 113, 127, 128, 136, 141, 152, 201, 212, 220, 257 Enhancer trap ............................................................5, 7, 9 Eosin Y......................................................... 89, 91, 96, 97

Martin Bayer (ed.), Plant Embryogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2122, https://doi.org/10.1007/978-1-0716-0342-0, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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296 Index

AND

PROTOCOLS

Erythrosine .................................................................... 172 1–Ethyl–3–(3–dimethylaminopropyl)carbodiimide (EDC) ...............................................87, 89, 92–94 Ethyl methanesulfonate (EMS).............. 7, 15–22, 25–31

F Farmer’s fixation solution.................................... 128, 130 Feeder cells ...................................................260–262, 266 Fertilization .......................................................7, 26, 167, 192, 193, 205, 207, 224, 257–259 Feulgen staining ................................................... 191–202 Fiji ....................................................................... 71, 72, 76 Fixative solution ............................. 65, 68, 208, 212, 213 Flow cytometer ................................................... 271, 272, 277, 278, 284, 287, 289 Flower buds .........................................104, 148, 274, 275 Fluorescein diacetate (FDA)............................... 102, 104, 105, 109, 110, 271, 272, 276, 278–280, 285, 291 Fluorescence-activated nuclear sorting (FANS)..............................................128, 151–164 Fluorescence lifetime .......................................... 172, 174, 175, 177, 179, 180, 182, 186 Fluorescent proteins.................6, 7, 50, 51, 65, 142, 172 Formaldehyde-acetic acid-alcohol fixative ..................... 89 Formamide .................................................. 90, 93, 94, 97 FRET-FLIM ......................................................... 168, 171

G Gaussian blur .............................................................72, 76 Genetic screens..................................................... 3–11, 26 Germinal mutations ........................................................ 26 Germline sector ................................................................. 9 Globular stage .............................................. 51, 135, 157, 206–208, 211, 212, 214–216, 218–220 Glutaraldehyde ..................................................... 208, 246 Glycine ................................... 89, 92, 153, 157, 245, 286 GUS reporter gene ..................................... 243, 246, 250

H Hand dissection...................................113, 115, 116, 124 Heart stage ............................................................ 5, 7, 55, 56, 59, 152, 168, 171 HistoClear ..................................90, 92, 96, 97, 246, 249 Histological analysis ............................................. 246–249 Hoyer’s solution................................................... 6, 65, 68 Hybridization salts ....................................................89, 93

I Imaging........................................5, 6, 40, 44, 50, 57, 60, 63, 64, 67, 69, 143, 168–170, 173, 184, 191–202 Immersion medium ..................................................57, 61 Inaccessible tissues ...................................... 152, 155, 160 Induction medium ............................................... 287, 288

Inner cells ............................................................. 151, 157 In situ hybridizations ...............................................87–98, 207–209, 211–215, 219, 220 In vitro fertilization (IVF) .................................. 102, 258, 259, 262, 264, 265 Isolation medium ........................................ 272, 275, 279

K Karyogamy..................................................................... 258

L Laser capture microdissection (LCM) ............... 205–209, 211–216, 218–221 Lateral root................................................ 64, 65, 68, 151 Library generation .......................................114, 117–121 Live-cell imaging .......................................................37–46

M Macerozyme .................................................................. 103 Maize ............................................................. 3–11, 25–31, 102, 127, 191–202, 207, 258, 259 Maleic acid ....................................................................... 89 Mannitol ............................ 102, 103, 110, 259–262, 265 Maturation medium................... 243, 245–247, 249–252 Maturing embryos (MEs)........................... 102, 103, 251 Methylimidazole........................................................89, 92 Methyl salicylate .........................192, 196, 199, 200, 202 Microdissection ...................................102, 127–138, 205 Micro-RNAs .................................................................... 16 Microscopy .......................................6, 50, 51, 64, 68–69, 130, 131, 200, 205, 236, 246, 248, 254, 258 Microspore ........................ 269–282, 284, 288, 289, 291 Microspore culture.......................................269–282, 291 Microtome .......................................................89, 91, 129, 131, 192, 209, 214, 215, 218, 219, 226, 232 Microtubules (MTs)..................... 38, 44, 50, 51, 60, 258 Microtubule stabilizing buffer ....................................... 51 Mineral oil .......................................................25, 27, 102, 105, 110, 259, 261, 262 Modified pseudo-Schiff propidium iodide (mPS-PI) ........................................................65, 66 MorphoGraphX.........................................................63–82 2-(N-Morpholino) ethanesulfonic acid hydrate (MES) .............................................. 102, 103, 251 Morphological analysis ..................................................... 5 Mounting medium...................................................54–56, 60, 61, 68, 95, 229, 230 Mutagenesis.............................. 4, 7–9, 11, 15–22, 25–31

N NBT/BCIP stock solution .......................................90, 95 NGS library preparation ............................. 117, 119, 121 Nicotiana tabacum ..................................... 102, 270, 283

PLANT EMBRYOGENESIS: METHODS NLN-13 medium ........................................ 272, 279, 281 Non-concordant kernels ...........................................29–32 Norway spruce ..................................................... 241–254 N5T medium...................................................... 38, 41, 43 Nuclear Isolation Buffer (NIB) ..........152, 155, 272, 285 Nuclei purification buffer (NPB) ............... 144, 146, 147 Nucleus .....................................37–39, 44, 177, 223, 258 N6Z medium ....................................................... 260, 261

O Ovaries ...................... 9, 32, 45, 191–201, 260, 262, 265 Ovule cultivation medium .............................................. 38

AND

PROTOCOLS Index 297

Rhodamine ........................................................... 172, 174 Rice ......................................................102, 127, 257–266 RNA extraction .......................................... 114, 117–121, 133, 135, 137, 153, 157, 158 RNAlater............................ 114–117, 124, 152, 155, 162 RNA sequencing (RNA-seq).............................. 108, 133, 135, 149, 152, 157, 160, 207, 221 RNaseZap .................................89, 92, 95, 114–117, 128 Rooting medium .................................................. 262, 287 Roots............................................................. 5, 26, 37, 57, 64–66, 68, 69, 72–79, 142, 151, 152, 160, 183, 224, 242, 243, 281, 290

P

S

Paraplast plus ................................................................... 90 Parent-of-origin effects .................................... 7, 9–11, 31 PDMS micropillar array device.................................38, 40 Periodic acid ................................... 65, 68, 192, 196, 198 Phaseolus vulgaris .......................................................... 217 Phosphate-buffered saline (PBS) ........................... 65, 78, 89, 92, 144–146, 148, 152, 153, 162, 225, 229–231, 233, 234, 236, 246 Photon counting mode .................................................. 57 Picea ............................................................................... 244 Pinus sylvestris....................................................... 224, 244 Pistil ............................................................................... 104 Plant development ................................................. 65, 101 Ploidy analysis......................................271, 272, 282, 284 Pollen ...................................................... 9–11, 17, 25–31, 196, 210, 211, 217, 224, 261, 262, 269, 270, 278, 280, 284, 288, 289, 291 Pollen mutagenesis............................................. 26, 27, 29 Pollination .......................................................27–31, 104, 105, 109, 135, 148, 196, 210, 211, 217, 224 Prematuration medium....................................... 243, 245, 246, 248–250, 252 Probe design..............................................................91, 96 Programmed cell death (PCD) .......................... 223–225, 235, 242, 244, 247 Proliferation medium..........................245, 247, 248, 252 Propidium iodide (PI) ....................................... 64–66, 68 Proteinase K ............................................................ 88, 89, 92–95, 97, 153, 162, 230, 233 Protein interaction ........................................................ 167 Protoderm ...........................................151, 162, 242, 244

Scarlet runner bean .............................................. 205–221 Schiff’s reagent..................................................... 192, 200 Scots pine......................................................223–236, 244 SCRI Renaissance........................................ 51, 65, 68, 69 SeaPlaque agarose ....................................... 246, 248, 250 Sectioning ................................................. 88, 91, 96, 137, 192, 193, 201, 209, 214, 220, 226, 231, 232, 254 Seed collection ..................................................... 104, 105 Seed development .............................4, 7, 9–11, 191–202 Seed isolation system .................................................... 143 Seed mutant................................................................... 4, 9 Segmentation ............................................................63–82 SHORT-ROOT (SHR) and SCARECROW (SCR) (SCR-SHR) ................................... 168, 171 Siliques........................................................ 5, 8, 9, 20, 21, 32, 41–43, 45, 50, 53–57, 59, 69, 87, 91, 96, 97, 115, 116, 128, 130, 135, 136, 145, 148, 155, 162, 169, 170, 184 Slide preparation .................................................. 129–132 Small RNAs ...............................................................87–98 Sodium hypochlorite .................................. 271, 274, 285 Solanum ......................................................................... 128 Solanum melongena.............................................. 283–292 Somatic embryogenesis........................................ 242–244 Sperm cells.........................................................25, 26, 29, 31, 102, 258, 260–262, 265 SSC.............................................................................89, 94 Stereomicroscope ...................................... 53, 55, 59, 169 Stigma ....................................................... 53, 55, 60, 109, 145, 210, 211, 217, 260, 265 Streptavidin.................................................................... 142 Subcellular structures................................................50, 57 Suspensor............................................109, 201, 205–207, 211, 212, 216–221, 224, 242, 243, 253

R Rapeseed ............................................................... 270, 275 Regeneration media ...................................................... 260 Renaissance staining.................................... 65, 66, 68, 69 Reporter genes ........................................ 5, 243, 246, 250 Reverse transcription (RT) ................................... 68, 117, 118, 153, 158–160, 163

T Tagmentation ............................................. 119, 120, 154, 155, 160, 161, 164 Tassels ........................................................................28–31

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298 Index

AND

PROTOCOLS

Tissue culture ................................................................ 292 Tissue embedding ................................. 89, 129–132, 209 Tissue enrichment test ........................114, 115, 122–124 Tissue fixation ...........................................................88, 91 Tissue-specificity............................................................ 137 TNM5 buffer.............................................................89, 95 TNP buffer ......................................................... 89, 95, 98 Tobacco ...................................... 101–110, 258, 270, 283 Toluidine blue staining ........................................ 229, 232 Tomato ................................................128, 130, 135, 283 Torpedo stage...................... 8, 38, 51, 60, 152, 157, 162 Transcriptomes ............................................. 63, 101, 102, 105, 108, 113–125, 127, 128, 137, 141, 144, 145, 147, 148, 151, 152, 205, 216, 218 Transgenic cell lines ............................................ 243, 247, 249, 251, 252, 254 Tris-buffered saline (TBS) .............................................. 89

tRNA..........................................................................89, 93 TUNEL assay ...................................................... 224, 225, 227, 229, 233, 234, 236 Tungsten needles ................................................. 114–116 Two-photon excitation microscopy (2PEM) ..........................................................38, 40

V Voxel size ..................................................... 71, 72, 76, 78

Z Zea mays......................................................................... 191 Zygotes ................................................................ 7, 37–39, 43–46, 50, 101, 102, 106, 109, 257–266 Zygotic genome activation (ZGA)............................... 101