Biolistic DNA Delivery: Methods and Protocols [1 ed. 2020] 1071603558, 9781071603550

Table of contents :
Foreword
Preface
Contents
Contributors
Part I: Tribute to Professor Diter von Wettstein
Chapter 1: Diter von Wettstein, Professor of Genetics and Master of Translating Science into Applications
1 Family History
2 A Brief Biography
3 A Glimpse into Diter´s Personality and Research
3.1 Fundamental Research
3.2 Interphase Research
3.2.1 Breeding for Root and Crown Rot Resistance
3.2.2 Production of Nutritionally Enhanced ``Celiac-Safe´´ Wheat Genotypes
3.2.3 Production of Wheat Genotypes with Improved Bioavailability of Dietary Fibers
3.3 Applied Research
3.3.1 Breeding for Imidazolinone Tolerance
3.3.2 Breeding for Proanthocyanidin-Free Malting Barley
3.3.3 Breeding for Increased Bioavailability of β-Glucans
4 Closing Remark
References
Chapter 2: Diter von Wettstein and The Meiotic Program of Pairing and Recombination
References
Part II: Background and Overview
Chapter 3: A Short History and Perspectives on Plant Genetic Transformation
1 Introduction
2 Agrobacterium-Mediated Plant Genetic Transformation
3 Biolistics or Particle Bombardment-Mediated Genetic Transformation of the Nuclear Genome
4 Biolistics-Mediated Chloroplast Transformation
5 Bioactive Beads-Mediated Gene Transfer
6 Pollen and Pollen Tube Pathway-Mediated Plant Transformation
7 Silicon Carbide Whisker- and Nanoparticle-Mediated Plant Transformation
8 Electrophoretic Transfection and Electroporation-Mediated Plant Transformation
9 Microinjection-Mediated Plant Transformation
10 Progress in Recalcitrant Plant Transformation
11 Gene Integration Through Homologous Recombination
12 CRISPR/Cas9-Mediated Genome Editing and Beyond
13 Conclusions
References
Chapter 4: Choice of the Promoter for Tissue and Developmental Stage-Specific Gene Expression
1 Introduction
2 Materials
2.1 Fruit-Specific Promoters
2.2 Flower-Specific Promoters
2.3 Anther/Pollen-Specific Promoters
2.4 Seed-Specific Promoters
2.5 Root-Specific Promoters
2.6 Developmentally Regulated Promoters
3 Notes
References
Chapter 5: Choice of Explant for Plant Genetic Transformation
1 Introduction
2 Plant Genetic Transformation
2.1 General Strategies for Transgene Delivery
2.2 Biolistics or Particle Bombardment Method
3 Different Types of Explants Used in Plant Genetic Transformation
4 Factors Influencing the Choice of Explants
4.1 Explant Age
4.2 Explant Size
4.3 Explant Excise Position
4.4 Explant Orientation
4.5 Explant Genotype
5 Explant Pretreatment
6 Explant Preparation
7 Application of Particle Bombardment Method in Plant Genetic Transformation
8 Conclusion
References
Chapter 6: Biolistic Approach for Transient Gene Expression Studies in Plants
1 Introduction
2 Advantages and Limitations of Transient Expression Mediated by Biolistics
2.1 Range of Species and Tissue Targets
2.2 Target Organelles
2.3 Plant Sample Preparation
2.4 Vector Preparation
2.5 Invasiveness
2.6 Early Events Following Microparticle Bombardment
2.7 Time-Frame of Biolistic Experiments
3 Examples of Biolistic-Mediated Transient Expression Studies
3.1 Transcriptional Regulation and Promoter Activity
3.2 Protein Subcellular Localization
3.3 Cell-to-Cell Protein Movement
3.4 Virus Inoculation
3.5 RNA Silencing Induction
3.6 Targeted Genome Editing
4 Discussion and Conclusion
References
Chapter 7: Nanobiolistics: An Emerging Genetic Transformation Approach
1 Introduction
2 Discussion
2.1 Synthesis and Characterization of Nanoparticles for Nanobiolistics
2.1.1 Synthesis of Mesoporous Silica Nanoparticles
2.1.2 Synthesis of Gold Nanoparticles
2.1.3 Nanoparticle Characterization
2.2 Testing Nanobiolistics in Animals
2.3 Emerging Studies of Nanobiolistics in Plants
3 Conclusions
References
Part III: Protocols
Chapter 8: Biolistic Transformation of Japonica Rice Varieties
1 Introduction
2 Materials
2.1 Plasmid Constructs
2.2 Plant Material (See Note 2)
2.3 Stock Solutions (See Note 3)
2.4 Culture Media (See Note 5)
2.5 Equipment and Related Supplies
3 Methods
3.1 Microprojectile Preparation (See Note 7)
3.1.1 Plasmid Confirmation
3.1.2 Gold Microprojectile Preparation
3.1.3 DNA/Gold Coating and Loading to Macrocarriers
3.2 Embryo Explant Preparation
3.2.1 Seed Sterilization
3.2.2 Explant Preparation
3.2.3 Osmotic Treatment, Bombardment and Resting
3.3 Selection of Transformed Calli
3.4 Regeneration and Rooting
3.5 Growing Plants in Growth Chambers
3.6 Transgene Inheritance Analysis
4 Notes
References
Chapter 9: Biolistic DNA Delivery in Maize Immature Embryos
1 Introduction
2 Materials
2.1 DNA Constructs (See Note 1)
2.2 Plant Material
2.3 Stock Solutions
2.4 Culture Media
2.5 Equipment and Related Supplies
3 Methods
3.1 Microprojectile Preparation (See Note 8)
3.1.1 Preparing of Gold Microprojectiles
3.1.2 Coating DNA on the Gold Particles
3.1.3 Loading the DNA/Gold onto the Macrocarrier
3.1.4 Preparing the Biolistic Gun Device for Bombardment
3.2 Maize Embryo Explant Preparation
3.2.1 Embryo Dissection
3.2.2 Pre-bombardment Osmotic Treatment
3.3 Bombardment
3.3.1 Launch the Biolistic Device
3.3.2 Microprojectile Bombardment of DNA (See Note 25)
3.3.3 Post-bombardment Biolistic Gun Care
3.3.4 Post-bombardment Osmotic Treatment and Transient Assays
3.4 Selection and Regeneration
3.4.1 Selection of Stable Transgenic Events
3.4.2 Callus Maturation
4 Notes
References
Chapter 10: Biolistic DNA Delivery and Its Applications in Sorghum bicolor
1 Introduction
2 Sorghum Tissue Culture
2.1 Explants
2.2 Media
2.3 Environment
3 Biolistic Delivery Parameters
3.1 Nanoparticles
3.2 Selection
4 Applications
4.1 Expression of Transgenes
4.2 RNAi
4.3 Genome Editing and CRISPR/Cas9
5 Future of Particle Bombardment in Sorghum
References
Chapter 11: Biolistics-Mediated Gene Delivery in Sugarcane
1 Introduction
2 Materials
2.1 Target Tissue
2.2 Stock Solutions and Media
2.3 DNA for Bombardment into Sugarcane Cells
2.4 Basic Equipment and Reagents for Bombardment
3 Methods
3.1 Preparation of Target Tissue
3.2 Preparation of Gold Particles
3.3 Microcarrier Preparation for Bombardment
3.4 Microprojectile Bombardment
3.5 Post Bombardment
3.6 Molecular Analysis of Regenerants
4 Notes
References
Chapter 12: Genetic Transformation of Common Wheat (Triticum aestivum L.) Using Biolistics
1 Introduction
2 Materials
2.1 Media
2.1.1 Stock Solutions
2.1.2 Plant Culture Media Stock Solutions
2.1.3 Final Culture Media
2.2 Isolation of Target Explants
2.2.1 Growth of Donor Plants
2.2.2 Sterilization of Immature Caryopses
2.3 Components for Particle Bombardment
3 Methods
3.1 Isolation of Target Explants
3.1.1 Collection and Sterilization of Immature Caryopses
3.1.2 Isolation of Immature Scutella
3.2 Preparation of Gold Microcarriers for Particle Bombardment
3.2.1 Preparation of Gold Microcarriers Stock
3.2.2 Coating of Gold Microcarriers with DNA
3.3 Transformation Via Particle Bombardment
3.3.1 Preparation of Particle Gun and Components
3.3.2 Assembling the Gun
3.3.3 Firing the Gun
3.3.4 Disassembling the Gun
3.3.5 Conclusion of Bombardment
3.4 Regeneration and Selection of Transgenic Plantlets Following Bombardment
3.4.1 Induction of Embryogenic Callus
3.4.2 Introduction of Selection
3.4.3 Increasing Selection Pressure
3.4.4 Regeneration of Plantlets
3.4.5 Rooting of Plantlets
3.4.6 Potting Putatively Transformed Plantlets to Soil
3.4.7 Analysis of Plants and Further Growth
4 Notes
References
Chapter 13: Biolistic DNA Delivery in Turfgrass Embryonic Callus Initiated from Mature Seeds
1 Introduction
2 Materials
2.1 Plant Material
2.2 Plasmids
2.3 Culture Media and Solutions
2.3.1 Chemical Stock
2.3.2 Initiation of Embryogenic Callus from Mature Seeds
2.3.3 Plant Selection and Regeneration Media
3 Methods
3.1 Initiation and Proliferation of Embryonic Callus from Mature Seeds
3.2 Callus Preparation Prior to Bombardment
3.3 Biolistic Transformation of Creeping Bentgrass Embryogenic Calli with pSBUbibar-35SGUS
3.4 Selection and Regeneration of Transformed Colonies
3.5 Regenerated Plants Maintenance
3.6 Molecular Analysis of Transgene Integration and Expression in the Transformed Plants
3.6.1 PCR Analysis
3.6.2 Southern Blot Analyses
3.6.3 Northern Blot Analyses
4 Notes
References
Chapter 14: Use of Microspore-Derived Calli as Explants for Biolistic Transformation of Common Wheat
1 Introduction
2 Materials
2.1 Plants
2.2 Stock Solutions
2.3 Specific Laboratory Equipment and Supplies
3 Methods
3.1 Growth Conditions and Selection of Spikes
3.2 Pretreatment
3.3 Isolation of Microspores
3.4 Preparation of Plant Material for Microprojectile Bombardment
3.5 Preparation of Gold Particles Stock Solution for Biolistic Transformation of Microspore-Derived Calli
3.6 Preparation of Gold Particles for Bombardment
3.7 Coating the Macrocarrier
3.8 Microprojectile Bombardment
3.9 Regeneration
3.10 Colchicine Treatment
3.11 Determination of Ploidy by Chromosome Analysis
4 Development of Wheat Genotypes with Reduced Immunogenicity
4.1 Construction of dTALE Repressor (Donor) and CRISPR/Cas9 Nuclease Constructs
4.2 Identification of Candidate Transformants with Adduct Lesions in the Wheat Dre2 Gene Homoeologs
5 Notes
References
Chapter 15: Plant Transformation Techniques: Agrobacterium- and Microparticle-Mediated Gene Transfer in Cereal Plants
1 Introduction
1.1 State-of-the-Art Transformation Techniques in Cereals
2 Agrobacterium-Mediated Transformation
2.1 Materials
2.2 Plant Material and Growth Conditions
2.3 Preparation of A. tumefaciens Electrocompetent Cells
2.4 Surface Sterilization of Immature Barley Seed
2.5 Isolation of Immature Embryos and Inoculation with A. tumefaciens
2.6 Generation of Plants from Calli
2.7 Determination of GFP in Transformed Tissue
3 Biolistic Transformation
3.1 Materials
3.2 Collection and Sterilization of Wheat Caryopses
3.3 Isolation of Immature Wheat Embryos
3.4 Coating of Gold Particles with DNA (for Six Shots)
3.5 Delivery of DNA-Coated Gold Particles
3.6 Histochemical Assays for GUS Detection
4 Notes
References
Chapter 16: Proteolistics: A Protein Delivery Method
1 Introduction
2 Materials
2.1 Plant Materials
2.2 Stock Solutions and Media
2.3 Proteins (See Note 4)
2.4 Equipment and Related Supplies
2.5 Other Supplies
3 Methods
3.1 Gold Particle Preparation
3.2 Protein-Gold Particle Suspension Preparation
3.3 Drying the Suspension on the Macrocarrier
3.3.1 Air-Drying
3.3.2 Freeze-Drying
3.3.3 Storage of Macrocarrier-Set Until Bombardment and Post-Drying Protein Check
3.4 Plant Target Tissue Preparation
3.4.1 Preparation of Onion Epidermis Tissue
3.4.2 Preparation of Maize Immature Embryos
3.5 Bombardment Using PDS-1000/He Biolistic Device (See Note 18)
3.6 Monitoring Intracellular Delivery
4 Notes
References
Chapter 17: Biolistic Delivery of Programmable Nuclease (CRISPR/Cas9) in Bread Wheat
1 Introduction
2 Materials
2.1 Materials for Designing and Validating CRISPR/Cas9 RNPs
2.1.1 sgRNA Cloning
2.1.2 In Vitro Transcription of sgRNA
2.1.3 Cas9 RNP Synthesis and Validation
2.2 Material for Biolistic Delivery of Cas9 RNPs and Mutant Screening
2.2.1 Plant Material
2.2.2 Sterilization Material
2.2.3 Particle Bombardment
2.2.4 Regeneration of Embryos
2.2.5 Mutant Screening
3 Methods
3.1 Designing and Validation of CRISPR/Cas9 RNPs
3.1.1 Designing and Synthesis of Single-Guide RNA (sgRNA) for Cas9
3.1.2 Designing ssDNA oligos (Only for Homology Directed Repair)
3.1.3 Cloning of sgRNA into Scaffold Vector
3.1.4 In Vitro Transcription of sgRNA
3.1.5 Production of Cas9 Protein
3.1.6 Validation of Cas9 RNPs Activity
In Vitro Assay
In Vivo Assay (Using Protoplast)
Assessment of Cas9 RNP Activity
3.1.7 Production of Cas9 RNPs
3.2 Biolistic Delivery of CRISPR/Cas9 RNPs and Mutant Screening
3.2.1 Wheat Growth Conditions
3.2.2 Harvesting Spikes and Sterilization
3.2.3 Isolation of Immature Embryo
3.2.4 Preparation of Gold Particle Suspension
3.2.5 Coating of Cas9 RNPs on Gold Nanoparticles
3.2.6 Biolistic Delivery of Cas9 RNPs
3.2.7 Verification of Cas9 RNP Delivery
3.2.8 Tissue Culture Regeneration of Bombarded Embryos
3.2.9 Mutant Screening
4 Notes
References
Index

Citation preview

Methods in Molecular Biology 2124

Sachin Rustgi · Hong Luo Editors

Biolistic DNA Delivery in Plants 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.

Biolistic DNA Delivery in Plants Methods and Protocols

Edited by

Sachin Rustgi Department of Plant and Environmental Sciences, School of Health Research, Pee Dee Research and Education Center, Clemson University, Florence, SC, USA

Hong Luo Department of Genetics and Biochemistry, Clemson University, Clemson, SC, USA

Editors Sachin Rustgi Department of Plant and Environmental Sciences School of Health Research Pee Dee Research and Education Center Clemson University Florence, SC, USA

Hong Luo Department of Genetics and Biochemistry Clemson University Clemson, SC, USA

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-0355-0 ISBN 978-1-0716-0356-7 (eBook) https://doi.org/10.1007/978-1-0716-0356-7 © Springer Science+Business Media, LLC, part of Springer Nature 2020 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: 1 New York Plaza, New York, NY 10004, U.S.A.

Foreword Diter von Wettstein was among the last of the twentieth century’s renaissance men of science and the scion of a distinguished family of scientists, including his grandparents, parents, and siblings. Following in their grand tradition, he was a naturalist with knowledge and expertise in a wide variety of subjects. At a time when many of his contemporaries were becoming highly specialized, he used his versatility and vast knowledge by making significant contributions to yeast genetics, plant breeding, epigenetics, and in pioneering studies on barley mutants, organelle (chloroplasts) biology including ultrastructural studies, lipid and chlorophyll biosynthesis, mapping and cloning of genes, and biotechnology. Although much of his work was devoted to seeking answers to important biological questions, he believed strongly in the social responsibilities of scientists and in application of science to the good of humanity and the world we live in. Long before it became fashionable to talk about “New Plant Breeding Technologies” (the so-called NPBTs), he practiced them all. For example, he contributed to the development of wheat strains with significantly reduced amounts of gluten to help people suffering from celiac disease, transgenic strains of barley that were resistant to stem rust and others that could be used for chicken feed instead of corn, and his work on yeast and barley that led to improvements in beer production. He was thus deeply disappointed and disturbed that the fruits of biotechnology were being denied to farmers and consumers alike for ideological and political rather than scientific reasons not only in Europe but more importantly in the developing economies of Africa and Asia which stand to benefit most from biotech crops. He was a strong advocate of public education and involvement in science. I did not know Diter well but had the great pleasure of meeting him many times. On two occasions, he was my host in Copenhagen, first in 1972 when I was at the University of Copenhagen, and again in 1980 at the Carlsberg Laboratory, which he transformed from a little-known corporate group to an internationally renowned institution. This made his corporate bosses happy and ensured continued generous funding. After my seminar in 1972, which was held in an ornate high-ceilinged hall in an old building in the Botanical Garden, he took me home where I was delighted to meet his charming wife Penny von Wettstein-Knowles, a distinguished plant biochemist. Their warm and gracious hospitality made it a relaxed and memorable evening of great food, wine, and wonderful conversation. He looked on proudly as Penny talked to me about her latest research on plant surface waxes. Unlike the tight-lipped rigid formality of his counterparts in Europe, he had a friendly, informal, and open manner which was partly a result of his having lived and worked for a time in the United States. Diter was a great raconteur who enjoyed sharing his rich body of personal and professional experiences. I witnessed this first-hand at the first International Congress of Plant Molecular Biology held in Savannah, GA, in 1985, when I went to greet him and Penny who were sitting at a table in the bar area during a reception. He was in a jovial mood surrounded by an eager group enthralled by with his unique and often witty stories. This volume is a fitting and fond tribute to Diter von Wettstein, the man and the scientist. University of Florida, Gainesville, FL, USA

v

Indra Vasil

Preface In the post-genomics era, the reliable gene delivery methods for expression, functional characterization, and subcellular localization of proteins are of paramount importance. Both transient expression and stable genetic transformation methods have their unique importance in advancing the basic understanding of different plant processes and in the development of agronomically desirable cultivars. Today, a large proportion of crops (staple and horticultural) grown worldwide are genetically engineered either for agronomical traits such as resistance to major insect pests, drought, and herbicides or industrially important traits such as better end-use quality or extended shelf life. Microprojectile bombardment via a biolistic particle delivery system or gene gun is one such method that allows physical delivery of the foreign biomolecules into the viable fungal, oomycete, plant, or mammalian cells. The method relies on precipitation of the nucleic acid, protein, or nucleoprotein complex on the biologically inert microscopic gold or tungsten particles (i.e., microprojectiles) or more recently nanoparticles of variable origins and sizes and their delivery by a ballistic device into a viable cell. The method of biolistic DNA delivery was discovered in the late 1980s, soon after the discovery of the Agrobacterium-mediated gene delivery method. The Agrobacterium-based genetic transformation system is primarily genotype dependent and relies heavily on the interactions among genes in the Agrobacterium genome, its tumor-inducing (Ti) or hairy root-inducing (Ri) plasmid and the host cell. Transcriptomic and proteomic studies revealed that the Agrobacterium and host cell interactions result in differential regulation of several stress-induced and pathogenesis-related genes. This host–parasite interaction is mainly responsible for the genotype dependence of Agrobacterium. The researchers soon recognized this limitation and started to seek for a versatile genetic transformation system. Discovery of the biolistic gene delivery method of genetic transformation came as a breakthrough, and protocols to transform different cell types and tissues as well as plant species were sought. Discovery of the biolistic and Agrobacterium-mediated gene delivery methods have marked the beginning of agricultural biotechnology. The science of genetic transformation has moved long ways since the discovery of the biolistic gene delivery method, and several alternative transformation procedures have been identified. A few noteworthy examples include the use of polyethylene glycol, microinjection, sonication, high-frequency ultrasound, low-power laser, electroporation, liposome, silicon carbide whisker, cell-penetrating peptides, virus vectors, direct spray, and more recently the nanoparticles. Despite the availability of a variety of genetic transformation methods, the biolistic gene delivery method has always remained one of the most reliable and widely used methods and is expected to retain its popularity in the foreseeable future. In the last couple of years, the biolistic gene delivery method has been put to a variety of uses including delivery of virus particles, pathogen effectors, gene silencing and gene expression constructs, programmable meganucleases, nucleoprotein complex, and vaccine. The progress made in this research area was summarized in a number of excellent review articles, book chapters, and an earlier volume of the Methods in Molecular Biology series, entitled Biolistic DNA Delivery. The previous volume was more general in its treatment of the subject, whereas this volume is largely focused on the research performed in plants.

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Delivery of this volume became necessary to provide plant researcher with a consolidated laboratory manual for use. This volume compiles reliable protocols for the use of the biolistic DNA delivery method in different plant species and also covers a few general topics related to the subject. A comprehensive collection of protocols is intended to provide a practical guide for the novice as well as the advanced user in the field of plant genetic transformation. For the sake of simplicity, this volume is divided into three parts, where Part I covers two chapters (Chapters 1 and 2). These chapters are dedicated to the life and research of Professor Diter von Wettstein, one of the landmark names in the field of agricultural biotechnology and one of the original editors of proposed volume. Part II includes the non-protocol chapter that covers many relevant topics of interest and provides a broad overview of the field as well as through light on other exciting modifications of the system (Chapters 3–7). Whereas Part III covers reliable plant transformation procedures in different plant species (Chapters 9–17). To sum up, this volume brings together knowledge and the experience of experts in the field of genetic transformation, who contributed to the invention of methods covered in this volume or are using them regularly. The chapters give step-by-step instructions to implement biolistic transformation into scientific practice reproducibly. Each chapter is supported by a helpful “Notes” section, which offers valuable advice for the particular transformation procedure described. Thus, this methods book represents a useful resource for students, postdoctoral fellows, and principal investigators who work on plant genetic transformation. Florence, SC, USA Clemson, SC, USA

Sachin Rustgi Hong Luo

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

PART I

TRIBUTE TO PROFESSOR DITER VON WETTSTEIN

1 Diter von Wettstein, Professor of Genetics and Master of Translating Science into Applications. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sachin Rustgi and Birgitte Skadhauge 2 Diter von Wettstein and The Meiotic Program of Pairing and Recombination . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Denise Zickler

PART II

v vii xi

3

19

BACKGROUND AND OVERVIEW

3 A Short History and Perspectives on Plant Genetic Transformation . . . . . . . . . . . 39 Thakku R. Ramkumar, Sangram K. Lenka, Sagar S. Arya, and Kailash C. Bansal 4 Choice of the Promoter for Tissue and Developmental Stage-Specific Gene Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 69 Olga G. Smirnova and Alex V. Kochetov 5 Choice of Explant for Plant Genetic Transformation . . . . . . . . . . . . . . . . . . . . . . . . 107 Nibedita Chakraborty, Priyanka Chakraborty, Moutushi Sen, and Rajib Bandopadhyay 6 Biolistic Approach for Transient Gene Expression Studies in Plants . . . . . . . . . . . 125 Benoıˆt Lacroix and Vitaly Citovsky 7 Nanobiolistics: An Emerging Genetic Transformation Approach . . . . . . . . . . . . . . 141 Francis J. Cunningham, Gozde S. Demirer, Natalie S. Goh, Huan Zhang, and Markita P. Landry

PART III

PROTOCOLS

8 Biolistic Transformation of Japonica Rice Varieties . . . . . . . . . . . . . . . . . . . . . . . . . . 163 Raviraj Banakar and Kan Wang 9 Biolistic DNA Delivery in Maize Immature Embryos. . . . . . . . . . . . . . . . . . . . . . . . 177 Kan Wang, Huilan Zhu, and Morgan McCaw 10 Biolistic DNA Delivery and Its Applications in Sorghum bicolor . . . . . . . . . . . . . . . 197 Guoquan Liu, Karen Massel, Basam Tabet, and Ian D. Godwin

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Contents

Biolistics-Mediated Gene Delivery in Sugarcane . . . . . . . . . . . . . . . . . . . . . . . . . . . . Priya A. Joyce and Yue Sun Genetic Transformation of Common Wheat (Triticum aestivum L.) Using Biolistics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Caroline A. Sparks and Angela Doherty Biolistic DNA Delivery in Turfgrass Embryonic Callus Initiated from Mature Seeds . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Man Zhou, Junming Zhao, Dayong Li, Shuangrong Yuan, Ning Yuan, Zhigang Li, Haiyan Jia, Fangyuan Gao, Bekir San, Qian Hu, and Hong Luo Use of Microspore-Derived Calli as Explants for Biolistic Transformation of Common Wheat . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sachin Rustgi, Samneet Kashyap, Nii Ankrah, and Diter von Wettstein Plant Transformation Techniques: Agrobacteriumand Microparticle-Mediated Gene Transfer in Cereal Plants . . . . . . . . . . . . . . . . . . Jafargholi Imani and Karl-Heinz Kogel Proteolistics: A Protein Delivery Method . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Susana Martin-Ortigosa and Kan Wang Biolistic Delivery of Programmable Nuclease (CRISPR/Cas9) in Bread Wheat . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Abhishek Bhandawat, Vinita Sharma, Vikas Rishi, and Joy K. Roy

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

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Contributors NII ANKRAH • Department of Crop and Soil Sciences, Washington State University, Pullman, WA, USA SAGAR S. ARYA • TERI-Deakin NanoBiotechnology Centre, The Energy and Resources Institute, New Delhi, India RAVIRAJ BANAKAR • Department of Agronomy and Crop Bioengineering Center, Iowa State University, Ames, IA, USA RAJIB BANDOPADHYAY • Department of Botany, UGC-Center of Advanced Study, The University of Burdwan, Golapbag, West Bengal, India KAILASH C. BANSAL • TERI-Deakin NanoBiotechnology Centre, The Energy and Resources Institute, New Delhi, India ABHISHEK BHANDAWAT • National Agri-Food Biotechnology Institute (NABI), Mohali, Punjab, India NIBEDITA CHAKRABORTY • Department of Botany, UGC-Center of Advanced Study, The University of Burdwan, Golapbag, West Bengal, India; Department of Biotechnology, National Institute of Technology, Durgapur, West Bengal, India PRIYANKA CHAKRABORTY • Department of Botany, UGC-Center of Advanced Study, The University of Burdwan, Golapbag, West Bengal, India VITALY CITOVSKY • Department of Biochemistry and Cell Biology, State University of New York, Stony Brook, NY, USA FRANCIS J. CUNNINGHAM • Department of Chemical and Biomolecular Engineering, University of California, Berkeley, CA, USA GOZDE S. DEMIRER • Department of Chemical and Biomolecular Engineering, University of California, Berkeley, CA, USA ANGELA DOHERTY • Plant Sciences Department, Rothamsted Research, Hertfordshire, UK FANGYUAN GAO • Department of Genetics and Biochemistry, Clemson University, Clemson, SC, USA IAN D. GODWIN • Centre for Crop Science, Queensland Alliance for Agriculture and Food Innovation, The University of Queensland, Brisbane, Australia NATALIE S. GOH • Department of Chemical and Biomolecular Engineering, University of California, Berkeley, CA, USA QIAN HU • Department of Genetics and Biochemistry, Clemson University, Clemson, SC, USA JAFARGHOLI IMANI • Institute of Phytopathology, Research Centre for BioSystems, Land Use and Nutrition, Justus Liebig University Giessen, Giessen, Germany HAIYAN JIA • Department of Genetics and Biochemistry, Clemson University, Clemson, SC, USA PRIYA A. JOYCE • Sugar Research Australia, Brisbane, QLD, Australia SAMNEET KASHYAP • Department of Plant and Environmental Sciences, School of Health Research, Clemson University Pee Dee Research and Education Centre, Florence, SC, USA ALEX V. KOCHETOV • Institute of Cytology and Genetics, Siberian Branch, Russian Academy of Sciences, Novosibirsk, Russia KARL-HEINZ KOGEL • Institute of Phytopathology, Research Centre for BioSystems, Land Use and Nutrition, Justus Liebig University Giessen, Giessen, Germany

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xii

Contributors

BENOIˆT LACROIX • Department of Biochemistry and Cell Biology, State University of New York, Stony Brook, NY, USA MARKITA P. LANDRY • Department of Chemical and Biomolecular Engineering, University of California, Berkeley, CA, USA; California Institute for Quantitative Biosciences, QB3, University of California, Berkeley, CA, USA; Chan-Zuckerberg Biohub, San Francisco, CA, USA SANGRAM K. LENKA • TERI-Deakin NanoBiotechnology Centre, The Energy and Resources Institute, New Delhi, India DAYONG LI • Department of Genetics and Biochemistry, Clemson University, Clemson, SC, USA ZHIGANG LI • Department of Genetics and Biochemistry, Clemson University, Clemson, SC, USA GUOQUAN LIU • Centre for Crop Science, Queensland Alliance for Agriculture and Food Innovation, The University of Queensland, Brisbane, Australia HONG LUO • Department of Genetics and Biochemistry, Clemson University, Clemson, SC, USA SUSANA MARTIN-ORTIGOSA • Department of Agronomy and Crop Bioengineering Center, Iowa State University, Ames, IA, USA; KWS SAAT SE, Einbeck, Germany KAREN MASSEL • Centre for Crop Science, Queensland Alliance for Agriculture and Food Innovation, The University of Queensland, Brisbane, Australia MORGAN MCCAW • Department of Agronomy and Crop Bioengineering Center, Iowa State University, Ames, IA, USA THAKKU R. RAMKUMAR • Agronomy Department, IFAS, University of Florida, Gainesville, FL, USA VIKAS RISHI • National Agri-Food Biotechnology Institute (NABI), Mohali, Punjab, India JOY K. ROY • National Agri-Food Biotechnology Institute (NABI), Mohali, Punjab, India SACHIN RUSTGI • Department of Plant and Environmental Sciences, School of Health Research, Clemson University Pee Dee Research and Education Centre, Florence, SC, USA; Department of Crop and Soil Sciences, Washington State University, Pullman, WA, USA BEKIR SAN • Department of Genetics and Biochemistry, Clemson University, Clemson, SC, USA MOUTUSHI SEN • Department of Botany, UGC-Center of Advanced Study, The University of Burdwan, Golapbag, West Bengal, India; Department of Biotechnology, National Institute of Technology, Durgapur, West Bengal, India; Department of Botany, Durgapur Government College, Durgapur, West Bengal, India VINITA SHARMA • National Agri-Food Biotechnology Institute (NABI), Mohali, Punjab, India BIRGITTE SKADHAUGE • Carlsberg Research Laboratory, Copenhagen, Denmark OLGA G. SMIRNOVA • Institute of Cytology and Genetics, Siberian Branch, Russian Academy of Sciences, Novosibirsk, Russia CAROLINE A. SPARKS • Plant Sciences Department, Rothamsted Research, Hertfordshire, UK YUE SUN • Sugar Research Australia, Brisbane, QLD, Australia BASAM TABET • Centre for Crop Science, Queensland Alliance for Agriculture and Food Innovation, The University of Queensland, Brisbane, Australia DITER VON WETTSTEIN • Department of Crop and Soil Sciences, Washington State University, Pullman, WA, USA KAN WANG • Department of Agronomy and Crop Bioengineering Center, Iowa State University, Ames, IA, USA

Contributors

xiii

NING YUAN • Department of Genetics and Biochemistry, Clemson University, Clemson, SC, USA SHUANGRONG YUAN • Department of Genetics and Biochemistry, Clemson University, Clemson, SC, USA HUAN ZHANG • Department of Chemical and Biomolecular Engineering, University of California, Berkeley, CA, USA JUNMING ZHAO • Department of Genetics and Biochemistry, Clemson University, Clemson, SC, USA MAN ZHOU • College of Natural, Applied and Health Sciences, Wenzhou Kean University, Wenzhou, Zhejiang, China HUILAN ZHU • Department of Agronomy and Crop Bioengineering Center, Iowa State University, Ames, IA, USA DENISE ZICKLER • Institute for Integrative Biology of the Cell (I2BC), Centre National de la Recherche Scientifique (CNRS), CEA, Universite´ Paris-Sud, Universite´ Paris-Saclay, Gifsur-Yvette, France

Part I Tribute to Professor Diter von Wettstein

Chapter 1 Diter von Wettstein, Professor of Genetics and Master of Translating Science into Applications Sachin Rustgi and Birgitte Skadhauge Abstract The present and subsequent chapters in this volume are dedicated to the life and research of Professor Diter von Wettstein who contributed immensely to the development of science and education. His contributions spanned various fields of science such as genetics, physiology, ultrastructural analysis, molecular biology, genomics, and biotechnology including genome editing. He performed and promoted pioneering research in the fields of epigenetics, directed evolution of enzymes, synthetic biology (promoter and gene optimizations), and genomics (genome sequencing of baker’s yeast). Glimpses of his time at the Carlsberg Laboratory and Washington State University, with examples from the research performed at these institutions, are included in this chapter. His life is an inspiration to the next generation of biologists. Despite difficult situations, his persistent efforts and keen desire to learn enabled him to overcome obstacles. He always tried to attain the best, excelling in translating fundamental knowledge into applications. Key words Genetic transformation, Epigenetics, Gene editing, Wheat, Barley, Biotechnology

1

Family History Diter von Wettstein (Dietrich Holger Wettstein, Ritter von Westersheim) was born on September 20, 1929 into a family of nobles, whose roots trace back to Johann Rudolf Wettstein, a Swiss diplomat, who played a pivotal role in the Swiss independence, and later became the Lord Mayor of Basel. His place in history can be recognized from the appearance of his picture on the Swiss stamps. In Diter’s lineage, his grandfather, Richard Wettstein, Ritter von Westersheim, who was Professor of Botany in Prague and Director of the botanical garden there, attained extraordinary prominence due to his seminal research in plant evolution and systematics. The glory he enjoyed can be witnessed from his picture appearing on the Austrian 50 Schilling note. Richard married Adele (Diter’s grandmother), who was also a botanist by training and daughter of another renowned botanist Anton Joseph Kerner (Anton Kerner, Ritter von Marilaun) who was Professor of Systematic Botany at the

Sachin Rustgi and Hong Luo (eds.), Biolistic DNA Delivery in Plants: Methods and Protocols, Methods in Molecular Biology, vol. 2124, https://doi.org/10.1007/978-1-0716-0356-7_1, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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Sachin Rustgi and Birgitte Skadhauge

Fig. 1 Diter von Wettstein’s lineage. Some of these pictures were obtained from Diter von Wettstein and Penny von Wettstein-Knowles at different occasions

University of Vienna, and also curator of the botanical garden there. Diter’s father Fritz von Wettstein (Friedrich Wettstein, Ritter von Westersheim) was the director of the Kaiser Wilhelm Institute (now Max Planck Institute) for Biology in Berlin-Dahlem and a revered geneticist of his time. During his reign the institute gained prominence in plant breeding and botanical research. Diter’s uncle Otto von Wettstein (son of Richard von Wettstein and elder brother of Fritz von Wettstein) was curator of the herpetological collection at the Natural History Museum in Vienna and a renowned zoologist of his time. Carrying on the family tradition Diter’s elder sister, Udda Lundqvist also developed an interest in biological sciences and later became a scientist in the Nordic Genetic Resource Center (NordGen), where she is still involved in maintaining a massive collection of >10,000 barley mutants. Diter’s younger brother, Gerhard von Wettstein, like his older siblings, was a meritorious student of biology. He studied biochemistry under the Nobel laureate Adolf Butenandt at the Max Planck Institute for Biochemistry. Diter’s long-term consort Penny von Wettstein-Knowles is Professor of Biology at Copenhagen University and a revered

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biochemist and geneticist. She has made seminal contributions to our current understanding of the genetics of plant wax traits. Diter is survived by her, their two daughters Heidi and Kimbery, and two grandchildren Naomi and Joshua.

2

A Brief Biography His scientific education started at an early age, as botany dominated his home environment. Though the conditions were favorable for education the time was not in Diter’s favor, as he had lost his grandfather at a tender age of two and his father at the age of 15. Moreover, the adverse political condition of the country and breaking out of the World War II, all affected his primary education. Despite these difficulties and the hearing impairment Diter had since birth, he never let any of it interfere with his learning. He obtained two Ph.D. degrees, in fact, one in Biochemistry from the University of Tu¨bingen (Germany) and another in Genetics from Stockholm University (Sweden), at an early age of 23 and mastered four languages: English, German, Danish, and Swedish. Subsequently, in 1957, at an age of 27 he obtained a D.Sc. degree in Genetics from Stockholm University. In the same year, he joined Stockholm University as Assistant Professor of Genetics and served there until 1962. Diter obtained a Rockefeller fellowship in 1958, and traveled to the USA for a year to carry out research at the California Institute of Technology in Pasadena, the Cold Spring Harbor Laboratory in New York, and the Carnegie Institute of Washington at Stanford University. Diter moved in 1962 to Denmark as Professor of Genetics and Head of the Institute of Genetics at the University of Copenhagen. A decade later (1972) he was invited to join the Carlsberg Laboratory as Head of the Department of Physiology, a position he held until retirement in 1996. Thereupon he joined Washington State University as the R.A. Nilan Distinguished Professor where he served until 2016. While working at WSU he also served as adjunct professor in the Justus-Liebig University, Germany (2007–2010), and the Northeast Normal and Wuhan Universities in China. The above summary reveals that he was an active scientist in various capacities for 66 years in five different countries. His research career spanning over seven decades can be arbitrarily divided into three major research phases (Fig. 2). Some of the major achievements from each of these phases are elaborated in the following paragraphs.

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Fig. 2 Different phases of Diter’s academic life. Some of these pictures were obtained from Diter von Wettstein and Penny von Wettstein-Knowles at different occasions

3

A Glimpse into Diter’s Personality and Research In 1970 the Danish breweries Carlsberg and Tuborg, both of which had their own research departments, merged. At the same time with the 100 year celebration approaching for the foundation of the Carlsberg Laboratory in 1976, the management wanted to augment the Carlsberg Laboratory. This led eventually to an enormous expansion of the original building and the search for a new head for its Department of Physiology. The short version of a complicated process was that in 1972 Diter von Wettstein (DvW), professor of genetics at the University of Copenhagen, was headhunted to oversee theoretical and practical work that would be undertaken in a brand-new, ultra-modern Carlsberg Laboratory in the heart of Copenhagen, which would also house the Carlsberg Research Centre consisting of the combined applied research departments of the Carlsberg and Tuborg breweries. Only in 2015 were the Carlsberg Research Centre and the Carlsberg Laboratory joined, giving rise to the Carlsberg Research Laboratory. DvW was designated head of the Laboratory’s Department of Physiology in 1972, whose other Department, namely of Chemistry was headed by Professor Martin Ottesen. In the interim until 1976 while the Laboratory was being modernized, DvW immediately started to learn as much as possible about the brewing process. This led to his realization, for example, that the already substantial number of anthocyanin-free barley mutants identified conceivably offered a new pathway toward producing clear rather than cloudy beer (see Subheading 3.3.2). When moving to the Carlsberg Laboratory, DvW brought with him a team of scientists and technicians from the university and

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with the considerable funds allotted was able to hire additional coworkers. Energized by new assignments, fine surroundings and a highly motivated staff, DvW took on scientific responsibility for the physiological processes of two of the organisms, barley and yeast, relevant to beer production. The wide-ranging remit gave him ample scope for basic research initiatives in plant physiology, as well as those focusing on benefits to brewing, that might be derived from barley and malt. At the same time, he initiated research efforts in microbiology, with the emphasis on yeast and fermentation. DvW’s work with plant physiology and microbiology was complemented by extensive, practical breeding efforts to search for and develop improved organisms for the production of beer. It was obvious from DvW’s first day at the Carlsberg Laboratory that he was the right person to deliver what the leadership of the brewery had been looking for. Not only was he ambitious, full of initiative, meticulous and extremely hard working; he was also part of an influential scientific network. It was therefore no surprise that in the 1980s DvW advanced to become the leading scientist at Carlsberg Laboratory, assuming the title of Chairman of the Research Council. In that role, DvW’s task was to master the alignment, that is, link up the wishes of the Carlsberg Research Centre, Laboratory, brewery and advisers—a task that he considered enjoyable and interesting. On top of that, DvW was one of the driving forces behind projects that relied on international cooperation, such as those relating to barley breeding and yeast genomics. The Physiology Department was a mecca for post docs from all over the world. In a laboratory perspective, DvW was an enthusiastic leader with analytical talents and an innovative mind set. Experiments were carefully tracked, always with helpful, rational comments. And when it came to proofreading manuscripts and reports, new authors would soon get used to DvW’s tiny handwritten comments at the top of printouts. DvW also paid special attention to the design and presentation of journal illustrations and slideshows of results. Generating the perfect photo, image or graph was not as easy as it is with today’s technology. DvW always wanted to provide managers, peers and the public with benchmark presentations of data and ideas. In the best possible pedagogical way, he aspired to bring research to life and disseminate it. Another facet of the latter is demonstrated with respect to his strong views on the benefits that could be obtained from genetic engineering. DvW realized that to overcome the generally negative view of such experiments a handson introduction was required already in the high schools. Thus, he set up practical courses for high school teachers using yeast at the Laboratory on Saturdays giving them the background to then carry out such experiments with their own students, as well as providing the necessary financial support therefore. More than 30 years later this approach is still being exploited.

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Fig. 3 Order of Dannebrog awarded to Diter von Wettstein for his contribution to sciences. This picture was graciously provided by Penny von Wettstein-Knowles

Most of DvW’s employees—and not only those from his Carlsberg years—remember him as exciting, inspiring and, not least, fun. Born, it would seem, with a genetically dominant party gene, he enjoyed socializing with all kinds of colleagues, whether at parties, formal dinners, on excursions or in other situations where he delighted in providing entertainment with his humorous presentations. Not only did DvW establish social networks, procure proper housing and jobs for the significant others of the many collaborators who visited the Department of Physiology over the years, but he also involved local and state authorities in the work of the Laboratory, echoing Carlsberg founder J.C. Jacobsen’s mentality of engaging with the surrounding society. Thanks to his special drive and engagement, DvW was admired by many. During his time at the Carlsberg Laboratory and before, DvW was credited with numerous valuable discoveries in basic and applied science, always aimed at translating results into practice and thus helping to promote a general interest in innovation and research. In view of his numerous contributions to the Danish community, DvW was honored with the Order of Dannebrog. The Order is awarded to members of the armed forces or to meritorious civilians for an outstanding contribution to the sciences, arts or business (Fig. 3). In fact, this high civilian honor presented to DvW had helped the Brewery leaders in handpicking Diter to lead the Department of Physiology at the Carlsberg Laboratory.

Yongchun Wu, Jayaveeramuthu Nirmala Claudia Osorio Jennifer Van Fleet, Ventria Bioscience, Fort Collins Jennifer Van Fleet, Ventria Bioscience, Fort Collins Rainer Stahl, MALTAgen Forschung GmbH, Andernach, Germany Rainer Stahl, MALTAgen Forschung GmbH, Andernach, Germany Rainer Stahl, MALTAgen Forschung GmbH, Andernach, Germany

Ventria Bioscience, Fort Collins Ventria Bioscience, Fort Collins Ventria Bioscience, Fort Collins

Barley Trichoderma harzianum Barley Homo sapiens sapiens Barley Homo sapiens sapiens Barley Homo sapiens sapiens Barley Homo sapiens sapiens Barley Homo sapiens sapiens Barley Homo sapiens sapiens Barley Human influenza virus Barley African swine fever virus Barley Homo sapiens sapiens Barley Homo sapiens sapiens Barley Barley Barley Barley Barley

Endochitinase

Human type I collagen

Human lactoferrin

Human lysozyme

Human antithrombin III,

α-Antitrypsin,

Serum albumin

HIV-1 antibody

E2 envelope glycoprotein of Swine fever virus

Somatropin

Defensins

Adjuvant C1

BPBF

bZIP

Ventria Bioscience, Fort Collins

Ventria Bioscience, Fort Collins

Henny Horvath-O’Geen, Gamini Kannangara

Barley Synthetic (Bacillus species)

Coworkers

Heat stable (1,3-1,4)-β-glucanase

Source

Crop

Transgene

Table 1 List of transgenic cereals produced by Diter von Wettstein and colleagues at the Washington State University, Pullman Reference

[27]

[27]

2003–2005

2003–2005

2003–2005

2003–2005

2003–2005

2003–2005

2003–2005

(continued)

2000–2002 [27]

2000–2002 [27]

2000–2002 [27]

2002

2002

2003–2004 [26]

1998–2010 [11, 25]

1996–2003 [20–24]

Year

Necrology of Prof. Diter von Wettstein: Part I 9

Nii Ankrah, Sachin Rustgi Nii Ankrah, Sachin Rustgi

Rhoda Brew-Appiah, Sachin Rustgi Claudia Osorio, Sachin Rustgi

Nuan Wen, Sachin Rustgi

Weiguo Liu, Elizabeth Kohl, Sachin Rustgi

Mingming Yang, Nii Ankrah, Sachin Rustgi 2013–2016 [14]

Wheat Escherichia coli Wheat Trichoderma harzianum Wheat Synthetic Wheat Rice Wheat Flavobacterium meningosepticum, barley Wheat Pyrococcus furiosus, barley Wheat Synthetic Wheat Bacillus subtilis Wheat Pseudomonas fluorescens

Endochitinase (ThEn42)

DEMETER specific hairpin RNA (hpRNA)

DEMETER specific artificial microRNA (amiRNA)

Prolyl endopeptidase and Endoprotease B2

Prolyl endopeptidase and Endoprotease B2

DEMETER specific TALE repressor and DRE2 specific CRISPR/Cas9

β-Xylanase

Pyrrolnitrin operon (prnABCD)

2010–2016 [3, 4]

2010–2016 [3, 4]

2009–2016 [3]

2009–2012 [2, 3]

2003–2006 [12]

Samneet Kashyap, Nii Ankrah, Sachin Rustgi 2013–2019 [14, 15]

Shanshan Wen, Sachin Rustgi

2009–2013 [12]

2009–2013 [12]

2006–2010 [12]

2001–2003 [28]

β-Glucuronidase gene (uidA)

Nii Ankrah, Sachin Rustgi

Henny Horvath-O’Geen, Robert Brueggeman

Wheat Jelly fish

Reference

GFP

Year

Barley Barley

Coworkers

Rpg1

Source

Crop

Transgene

Table 1 (continued)

10 Sachin Rustgi and Birgitte Skadhauge

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A few selected projects directed by DvW are outlined in brief below. Most were carried out with various collaborators (see Table 1), highlighting DvW as an early and true science-to-application translator. 3.1 Fundamental Research

A major research project focused on the elucidation of the molecular basis of storage protein synthesis and deposition in the vacuoles of the developing barley endosperm was undertaken [1]. The obtained results gave both basic and applied insights. Under this research project gene and promoter sequences as well as molecular structures of the complex loci encoding B-, C-, and γ-hordeins were obtained. In addition, the mechanism of transcriptional regulation of the B-hordein genes was deciphered. It was one of the first demonstrations of the epigenetic regulation of gene expression or imprinting in cereal endosperm or in plants [1]. Later this phenomenon was rediscovered in Arabidopsis and studied in great detail, and the genes involved in epigenetic regulation of endosperm-specific gene expression were identified. DvW and group made use of this knowledge to develop reduced-immunogenicity wheat and barley genotypes for celiac patients, setting a great example of translational research [2–4]. The basic research on grain development specifically on the synthesis, subcellular transport, accumulation and catabolism of biomolecules such as the seed storage proteins and the cell wall polysaccharides have allowed identification of the endosperm specific promoters of the hordein genes and the low- and high-PI α-amylase genes. The team has tested these promoters by attaching them to the reporter genes for their spaciotemporal activities, expression levels, and influence of internal/external cues [5]. This basic knowledge has allowed characterization of a number of endosperm specific promoters, which can be used to express and accumulate large quantities of desired proteins in the cereal endosperm. While working on grain storage proteins, DvW and colleagues also discovered that storage protein synthesis during grain development is regulated at the transcriptional level and is dependent on the supply of nitrogen. In this connection, the group identified a GCN4 motif and a bifunctional endosperm element in the promoter of C-hordein genes that sense the nitrogen level and regulate transcription of the hordein genes. This research finding has far reaching implications in developing storage protein genes with constitutive expression—a desirable goal—to reduce the necessity for large amounts of fertilizer for sustainable high yield of proteins across environment years; and to develop promoters for biopharming [1]. Another basic problem which DvW and group worked on was engineering of proteins for thermostability, pH optimum, and dual functionality (multienzyme activity). It was one of the first demonstrations of protein engineering, where proteins were tailored to

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meet the industry requirements [1]. This research led to the development of (1,3;1,4) β-glucanase variants with enhanced thermostability, a wider pH optimum, or a multienzyme activity with properties of both a β-glucanase and a cellulase [1]. Following similar principles, the thermostable variants of a Flavobacterium meningosepticum prolyl endopeptidase (PE-FmPep) and a barley glutamine specific-endoprotease (EP-HvB2) were subsequently also developed by the group [3]. The advances made in the field of protein engineering by DvW and group in late 1980s [6], have later evolved into the full-blown field of the directed evolution of enzymes, which is finding great implications in the food, feed, brewing, biofuel, and the health industries (cf. Ref. 7). DvW always had a longing to understand the mechanism of chloroplast biogenesis and the biosynthesis of the photosynthetic membrane, as well as the process of chromosome pairing and crossing over. He performed seminal research in both of these research areas and lately summarized his findings in these areas in an outstanding review [8]. Moreover, his contributions to the fields of chloroplast biogenesis and chromosome pairing were recently revisited by Hoober [9] and Zickler [10], respectively. Therefore, discussion on these topics has been omitted here. Close to his retirement from the Carlsberg Laboratory, DvW added another dimension to his research and joined a team of international researchers to participate in the sequencing of the yeast genome. In this effort, his group has contributed the nucleotide sequences of yeast chromosomes III, X, and XI [8]. This research helped pave the way for the subsequent sequencing of other eukaryotes, including human and more recently bread wheat. 3.2 Interphase Research 3.2.1 Breeding for Root and Crown Rot Resistance

About 86% of the US wheat and 50% of the world’s wheat is produced under water scarcity (http://drought.mssl.ucl.ac.uk). Therefore, to conserve moisture, direct seeding operations were adopted in 35.5% of the US crop land planted to wheat, barley, corn, cotton, and soybeans. The management practices that leave crop residues on the surface also create an ideal environment for the increase of soil borne pathogens such as Rhizoctonia solani and Fusarium culmorum, the causal agents of Rhizoctonia root rot and Fusarium crown rot, respectively. Yield losses due to such pathogens make them the main limiting factors to the adoption of direct seeding in dry land cropping systems around the globe. To deal with this problem DvW and coworkers adapted two strategies: (1) Ectopic expression of the Trichoderma endochitinase gene in wheat and barley genomes. The chitinase enzyme catalyzes degradation of chitin molecules, a major component of the cell wall in true fungi including Rhizoctonia and Fusarium. Using the existing knowledge, DvW et al. expressed the Trichoderma harzianum ThEn42 gene in wheat and barley genomes, and studied these plants for tolerance to root and crown rot pathogenesis in the

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greenhouse under artificial inoculation. The wheat transformants were developed in the background of two commercial spring bread wheat cultivars Louise and Express using a microspore electroporation-based genetic transformation method. Whereas, barley transformants were developed by cocultivation of scutellar calli with Agrobacterium in the background of a regeneration competent cv Golden Promise and the transgene was later transferred to a commercial spring barley cv Baronesse by a conventional cross breeding approach [11–14]. (2) Biomimetic engineering of wheat to produce a natural agricultural fungicide in roots. Pyrrolnitrin is one of the natural agricultural fungicides produced by fluorescent Pseudomonas found in suppressive soils. This antibiotic is synthesized from the amino acid tryptophan by the concerted action of four enzymes encoded by an operon. To assure pyrrolnitrin production in wheat roots DvW and group engineered an expression vector by optimizing pyrrolnitrin operon genes for expression in wheat and provided it with a shared rice root specific promoter. The production of pyrrolnitrin was confirmed by triple-quadrupole (QQQ) mass spectrometer and laboratory bioassay with root and crown rot pathogens [14]. 3.2.2 Production of Nutritionally Enhanced “Celiac-Safe” Wheat Genotypes

Nutritional quality and food safety were largely ignored at the time of the “Green Revolution,” but this perception is rapidly changing, and in later years more attention has been vested in the improvement of nutritional quality and food safety of the harvest. About 7.5% of the US population and 1% of the world population suffers from the “gluten syndrome.” The only effective therapy known so far for these disorders is lifelong adherence to an abstinent diet, which is difficult to practice if not impossible. Therefore, to develop dietary therapies for the individuals suffering from celiac disease or allergy to wheat and its derivatives, DvW and coworkers followed three different strategies: (1) epigenetic elimination of immunogenic gluten proteins by tissue-specific (via RNA interference) or systemic (via induced mutagenesis) silencing of the wheat DEMETER gene, a master-regulator of gluten biosynthesis in grains [3]; (2) post-transcriptional silencing of immunogenic gluten proteins via endosperm-specific expression of a chimeric hairpin construct, which was specifically designed to target transcripts of the prolamin gene family [3]; and (3) post-translational detoxification of the wheat gluten proteins by endosperm-specific expression of two “glutenases” [4]. The outcome of these research activities was published in a series of research articles with a significant finding being the identification of wheat genotypes exhibiting up to a 76% reduction in the amounts of immunogenic prolamins and wheat lines expressing glutenases in the endosperm [2–4, 14]. To further increase the extent of gluten elimination, the group has lately targeted wheat DEMETER and DRE2 genes (responsible for activation of the DEMETER enzyme) using the CRISPR/Cas9

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approach [14, 15]. The reduced-immunogenicity wheat genotypes developed under this research project are expected to impact the lives of millions of individuals around the globe by improving the quality of their lives and by reducing healthcare cost. 3.2.3 Production of Wheat Genotypes with Improved Bioavailability of Dietary Fibers

Arabinoxylans or pentosans are the predominant component of the wheat endosperm cell wall that constitute ~6–7% of the dry matter in the grains. Unfortunately, mammals cannot digest these biological fibers, which significantly reduce their bioavailability to the consumer, and also cause discomfort on the consumption of these fibrous foods. As there is a great emphasis on the consumption of whole-grain products DvW and team transformed wheat to express a codon-optimized version of 1,4-β-xylanase from Bacillus subtilis [12]. These transformed wheat lines are expected to exhibit high nutritional value in the feeding trials performed on rodents and primates.

3.3

The acreage for barley is constantly declining which can be partially attributed to the large-scale application of imidazolinone herbicides (IMIs) and the adaptation of IMI-tolerant crops. In view of reestablishing barley in crop rotation schemes DvW and coworkers have started a screen for IMI-tolerant mutant(s) in a sodium azidetreated M2/M3 population of barley cultivar Bob [16]. This screen has resulted in identification of a mutant line carrying a single-point mutation leading to a serine to asparagine amino acid substitution in the herbicide-binding site of the barley acetohydroxy acid synthase (AHAS) enzyme [16]. Since the IMI-tolerant mutation was identified in the background of an outdated feed barley cultivar Bob it was transferred to six barley cultivars two each belonging to three market classes (food, feed, and malting) of barley using DNA marker-assisted selection [3, 13, 17]. The IMI-tolerant lines developed as above showed up to 90% recovery of the carrier chromosome, and can be released as improved varieties after necessary testing.

Applied Research

3.3.1 Breeding for Imidazolinone Tolerance

3.3.2 Breeding for Proanthocyanidin-Free Malting Barley

Proanthocyanidins (condensed tannins) exist naturally in the barley seed coats and are carried from the malt into the wort and from the wort into the beer, where they cause precipitation of proteins and haze formation especially upon refrigeration, which reduces the shelf life of beer. To produce brilliant clear beer today, proofing or stabilizing treatment with chemicals that chelate proanthocyanidins such as polyvinylpyrrolidone, or proteases are performed. As noted above one of DvWs first projects initiated at the Carlsberg Laboratory was to exploit and enlarge the collection of barley mutants that are free of anthocyanins and proanthocyanidins which today number circa 560 [1]. Several of these mutants were high yielding and gave rise to malting varieties for cultivation in Europe such as Caminant [ant28–484 (Grit)] and Clearity

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[ant27–488 (Zenit)]. After moving to Washington, DvW and crew bred two more proanthocyanidin-free barley varieties, Radiant [ant29–667 (Harrington)] with a yield matching that of Baronesse [18], and Fritz [ant-499 (Apex)] with a yield at least 1000 lb./a more than the large acreage varieties (i.e., Bob, Baronesse, and Champion) [13, 19]. It should be mentioned that haze stability of beer made from proanthocyanidin-free malt of Fritz is much better than that achieved by the presently employed chemical treatments. Fritz is, additionally, a personal favorite for many customers of the Skagit Valley Malting Company at Burlington, WA. These results illustrate DvW’s tenacity to continue with projects he believed in, in this case over 40 years, despite setbacks and impediments encountered along the way. 3.3.3 Breeding for Increased Bioavailability of β-Glucans

80% of the endosperm cell walls of barley consist of (1,3;1,4)-β-Dglucans, which cause unique problems in the use of barley for brewing and/or as poultry feed. During the malting process, for α-amylases to reach the starch grains accumulated in the endosperm cells it is essential for the β-glucan walls to depolymerize. Albeit the aleurone layer of endosperm in barley grains secretes β-glucanase the native enzyme does not survive the high temperature during the kilning process. Incomplete depolymerization of the β-glucan walls during germination in the malting process therefore causes an unacceptable high viscosity of wort, which is a significant problem in the full-scale filtration or centrifugation steps. For this reason, most barley varieties cannot be used for malting. On the other hand, the low nutritional value of barley for poultry is due to the absence of an intestinal enzyme for efficient depolymerization of (1,3;1,4)-β-D-glucans. The absence of appropriate digestive enzymes in poultry animals leads to high viscosity of β-D-glucans in their intestines, causing limited nutrient uptake, decreased growth rate, and unhygienic sticky droppings, which adhere to chickens and floors of the production cages. Consequently, the eight billion broiler chickens produced annually in the United States are primarily raised on corn-soybean diets [1]. In view of these problems, DvW and team set out to breed transgenic barley plants, which express a protein-engineered thermostable (1,3;1,4)-β-glucanase enzyme during germination/malting in the aleurone cells to allow secretion of enzyme into the endosperm. As expected, the resultant barley transformants expressing thermostable (1,3;1,4)-β-glucanase brewed into excellent beer without a need for filtration [5]. The team additionally, bred the transgene from Golden Promise into the widely cultivated spring feed barley cultivar Baronesse, and tested it for superior feed characteristics [1, 5].

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Closing Remark The US society has always been very open to new technologies, albeit the idea of adapting genetically modified food crops has never received general acceptance, which bothered DvW, who has always been an advocate of genetic engineering technologies to breed crops for enhanced productivity and safety. In fact, when the new legislature about the labeling of all GM ingredients on foods was proposed, Diter contradicted the idea. His purpose was not to conceal or provide incomplete information to the consumers, but to avoid spread of misconceptions or induction of panic among the public. In this connection, he encouraged the WSU administration to allow delivery of a new hands-on training course on the production of transgenic crops, with the expectation to increase the awareness of the general public about this technology and to improve the career prospects of the participating students. DvW also organized a National Academy of Sciences (NAS) symposium on the “Functional Genomics of Model Organisms to Crop Plants for Global Health” with a specific session on “Engineering Crops for the Future” in association with Roger Beachy and Robert Goldberg in 2006. Diter always viewed genetic engineering as a major solution to the complex problems encountered by mankind today and in the near future. The unsubstantiated negative attitude toward the transgenic technology and uninformed decisions made by the governments or the general public due to lack of knowledge frustrated DvW at times. In fact, in an extensive review of his work, DvW stated that “At present, breweries and malting companies will not use transgenic barley that is popularly, and nonsensically, called GM or GMO barley (standing for genetically modified or genetically modified organism barley). They are waiting for consumers to be enlightened that transgenic barley is not different and equally well tested as safe food as barley bred by mutation, selection, and hybridization for the past 10,000 years” [1]. The negative attitude of the general public toward transgenics, however, never prompted DvW to stop working in this research area (for examples, see Table 1), as he always hoped that “the time will come when transformed cultivars will be as accepted and considered as ‘traditional’ as crop plants containing induced mutations” [8].

Acknowledgments The authors would like to thank Penny von Wettstein-Knowles for her support, corrections, and providing pictures used in necrology and the financial support from the State of South Carolina grant S009, the Life Sciences Discovery Fund Grant 3143956-01, and the Clemson Faculty Succeeds Grant 1502211.

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References 1. von Wettstein D (2007) From analysis of mutants to genetic engineering. Annu Rev Plant Biol 58:1–19 2. Wen S, Wen N, Pang J, Langen G, BrewAppiah RAT, Mejias JH, Osorio C, Yang MM, Gemini R, Moehs CP, Zemetra RS, Kogel KH, Liu B, Wang X, von Wettstein D, Rustgi S (2012) Structural genes of wheat and barley 5-methylcytosine DNA glycosylases and their potential applications for human health. Proc Natl Acad Sci U S A 109:20543–20548 3. Rustgi S, Wen N, Osorio C, Brew-Appiah RAT, Wen S, Gemini R, Mejias JH, Ankrah N, Moehs CP, von Wettstein D (2014) Natural dietary therapies for the ‘gluten syndrome’. The Royal Danish Academy of Sciences and Letters, Copenhagen 4. Osorio C, Wen N, Mejias JH, Liu B, Reinbothe S, von Wettstein D, Rustgi S (2019) Development of wheat genotypes expressing a glutamine-specific endoprotease from barley and a prolyl endopeptidase from Flavobacterium meningosepticum or Pyrococcus furiosus as a potential remedy to celiac disease. Funct Integr Genomics 19:123–136 5. Horvath H, Huang J, Wong OT, von Wettstein D (2002) Experiences with genetic transformation of barley and characteristics of transgenic plants. In: Slafer GA, Molina-Cano JL, Savin R, Araus JL, Romagosa I (eds) . Food Products Press, Barley science resent advances from molecular biology to agronomy of yield and quality, pp 143–176 6. Borriss R, Olsen O, Thomsen KK, von Wettstein D (1989) Hybrid bacillus endo-(1–3,1–4)-β-glucanases: construction of recombinant genes and molecular properties of the gene products. Carlsb Res Commun 54:41–54 7. The Royal Swedish Academy of Sciences (2018) Scientific background on the Nobel prize in chemistry 2018 – directed evolution of enzymes and binding proteins. https://old. nobelprize.org/che-sci.pdf?_ga¼2. 228216788.850160180.15385606991556534510.1538560699 8. von Wettstein D (2006) Fascination with chloroplasts and chromosome pairing. Progress Bot 67:3–28 9. Hoober JK (2017) Diter von Wettstein (Dietrich Holger Wettstein Ritter von Westersheim): September 20, 1929–April 13, 2017. Photosynth Res 134:107–110 10. Zickler D (2020) Diter von Wettstein and the meiotic program of pairing and recombination. In: Rustgi S, Luo H (eds) Biolistic DNA delivery in plants. Springer, New York, NY (chapter 2, this volume)

11. Kogel KH, Voll LM, Sch€afer P, Jansen C, Wu Y, Langen G, Imani J, Hofmann J, Schmiedl A, Sonnewald S, von Wettstein D, Cook RJ, Sonnewald U (2010) Transcriptome and metabolome profiling of field-grown transgenic barley lack induced differences but show cultivarspecific variances. Proc Natl Acad Sci U S A 107:6198–6203 12. Brew-Appiah RAT, Ankrah N, Liu W, Konzak CF, von Wettstein D, Rustgi S (2013) Generation of doubled haploid transgenic wheat lines by microspore transformation. PLoS One 8: e80155 13. Rustgi S, von Wettstein D (2015) Breeding barley ornamented with the novel agronomical attributes. Med Aromat Plants 4:2 14. Rustgi S, Kashyap S, Gandhi N, von Wettstein D, Ankrah N, Gemini R, Reisenauer P (2018) Novel wheat genotypes designed to meet the future needs for safe and surplus food. Ann Wheat Newslet 64:56–61 15. Rustgi S, Kashyap S, Ankrah N, von Wettstein D (2020) Use of microspore-derived calli as explants for biolistic transformation of common wheat. In: Rustgi S, Luo H (eds) Biolistic DNA delivery in plants. Springer, New York, NY (chapter 14, this volume) 16. Lee H, Rustgi S, Kumar N, Burke I, Yenish JP, Gill KS, von Wettstein D, Ullrich SE (2011) Single nucleotide mutation in the barley acetohydroxy acid synthase (AHAS) gene confers resistance to imidazolinone herbicides. Proc Natl Acad Sci U S A 108:8909–8913 17. Rustgi S, Matanguihan J, Mejias JH, Gemini R, Brew-Appiah RAT, Wen N, Osorio C, Ankrah N, Murphy KM, von Wettstein D (2014) Assessment of genetic diversity among barley cultivars and breeding lines adapted to the US Pacific Northwest, and its implications in breeding barley for imidazolinoneresistance. PLoS One 9:e100998 18. von Wettstein D, Cochran JS, Ullrich SE, Kannangara CG, Jitkov VA, Burns JW, Reisenauer PE, Chen X, Jones BL (2004) Registration of ‘Radiant’ barley. Crop Sci 44:1859–1860 19. Rustgi S, Brouwer B, von Wettstein D, Reisenauer PE, Lyon S, Ankrah N, Jones S, Guy SO, Chen X (2020) Registration of ‘Fritz’, a two-row, spring barley. J Plant Registr. (under review) 20. Jensen LG, Olsen O, Kops O, Wolf N, Thomsen KK, von Wettstein D (1996) Transgenic barley expressing a protein-engineered, thermostable (1,3-1,4)-beta-glucanase during germination. Proc Natl Acad Sci U S A 93:3487–3491

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21. Jensen LG, Politz O, Olsen O, Thomsen KK, von Wettstein D (1998) Inheritance of a codon-optimized transgene expressing heat stable (1,3-1,4)-β-glucanase in scutellum and aleurone of germinating barley. Hereditas 129:215–225 22. Horvath H, Huang J, Wong O, Kohl E, Okita T, Kannangara CG, von Wettstein D (2000) The production of recombinant proteins in transgenic barley grains. Proc Natl Acad Sci U S A 97:1914–1919 23. von Wettstein D, Mikhaylenko G, Froseth JA, Kannangara CG (2000) Improved barley broiler feed with transgenic malt containing heat-stable (1,3-1,4)-beta-glucanase. Proc Natl Acad Sci U S A 97:13512–13517 24. von Wettstein D, Warner J, Kannangara CG (2003) Supplements of transgenic malt or grain containing (1,3-1,4)-β-glucanase to barley based broiler diets lift their nutritive value to that of corn. Br J Poult Sci 44:438–449

25. Wu Y, von Wettstein D, Kannangara CG, Nirmala J, Cook RJ (2006) Growth inhibition of the cereal root pathogens Rhizoctonia solani AG8, R. oryzae and Gaeumannomyces graminis var. tritici by a recombinant endochitinase from Trichoderma harzianum. Biocontrol Sci Tech 16:631–646 26. Osorio C (2004) Development of transgenic barley expressing human type I collagen. M. Sc., thesis, Department of Crop and Soil Sciences, Washington State University, Pullman WA 27. Stahl R, Horvath H, Van Fleet J, Voetz M, von Wettstein D, Wolf N (2002) T-DNA integration into the barley genome from single and double cassette vectors. Proc Natl Acad Sci U S A 99:2146–2151 28. Horvath H, Rostoks N, Brueggeman R, Steffenson B, von Wettstein D, Kleinhofs A (2003) Genetically engineered stem rust resistance in barley using the Rpg1 gene. Proc Natl Acad Sci U S A 100:364–369

Chapter 2 Diter von Wettstein and The Meiotic Program of Pairing and Recombination Denise Zickler Abstract Recombination and pairing are prominent features of meiosis where they play an important role in increasing genetic diversity. In most organisms recombination also plays mechanical roles in mediating pairing of homologous chromosomes during prophase and in ensuring regular segregation of homologous pairs at the first meiotic division. The laboratory directed by D. von Wettstein identified six key steps in the meiotic process: (1) Recombination mediated processes occur in physical and functional linkage with the synaptonemal complex (SC), a highly conserved, meiosis-specific structure that links homologous axes along their lengths. (2) The pairing process involves formation and resolution of chromosomal entanglements/interlockings. (3) The SC normally forms specifically between homologous chromosomes, but in unusual situations can form between nonhomologous chromosomes or regions resulting in two-phase SC formation. (4) In hexaploid common wheat, extensive multivalents form with multiple, pairing partner shifts, indicating homology recognition and SC formation among homoeologs as well as homologs. (5) Linkage between recombination and the SC is revealed by crossover-correlated nodules localized in the SC central region. (6) Modified SCs sometimes play a direct role in homolog segregation, providing the required connection between homologs in absence of crossovers/chiasmata. Key words Meiosis, Synaptonemal complex, Interlockings, Recombination nodules

By his exceptional insight into complex biological systems, his rigorous experimentation, and repeatedly combining genetic approaches with molecular biology and microscopy, Diter von Wettstein made seminal achievements in several different research areas. He produced high-profile papers both with respect to quantity and quality throughout his career and contributed to science propagation by organizing several meetings and workshops. Moreover, his work on plant biology played a key role in translating results from fundamental into applied research, and is therefore universally known and highly appreciated. Details concerning this part of his work are elaborated in the preceding chapter of this book. Although Diter is perhaps best known for his work on plant physiology, he made also enduring contributions to our Sachin Rustgi and Hong Luo (eds.), Biolistic DNA Delivery in Plants: Methods and Protocols, Methods in Molecular Biology, vol. 2124, https://doi.org/10.1007/978-1-0716-0356-7_2, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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understanding of chromosome pairing and recombination during meiosis. Meiosis is a highly conserved and complex process with a central role in most eukaryote life cycles. In contrast to mitotic divisions, meiotic divisions reduce the diploid chromosome complement by half as required for sexual reproduction. This is achieved by one round of DNA replication followed by two rounds of divisions without intervening replication. Paternal and maternal homologous chromosomes segregate to opposite poles during the first division whereas their sister chromatids (issued from premeiotic replication) segregate during the second division. Successful execution of the two meiotic divisions requires that during the first division, homologous chromosomes (homologs) are connected to one another in order to segregate to opposite poles of the spindle. Connections are achieved by reciprocal homologous recombination (crossovers, seen cytologically as chiasmata) between one sister chromatid of each homolog plus cohesion between sister chromatids all along their lengths. Connected homologs thus form a unit called a “bivalent” in which maternal and paternal centromeres/kinetochores can be attached to microtubules from opposite poles at division I. Before segregation, cohesion links between homologs are degraded along arms but not at centromeres to permit release of chiasmata without loss of sisterchromatid association. Release of sister-chromatid centromeric cohesion allows then their separation during the second division. Many of the most unique aspects of meiosis are devoted, therefore, to providing at least one chiasma per bivalent in order to achieve their correct segregation during the first division. Consequently, meiotic recombination at the DNA level plays a double role: an indispensable mechanical role for chromosome segregation and promotion of genetic diversity (reviews in Refs. 1–3). When, together with Mogens Westergaard, Diter von Wettstein developed a research group devoted to “meiotic pairing and recombination,” after his nomination as head of the Carlsberg Department of Physiology in 1972 (Fig. 1A), the existence of the meiotic process at the chromosomal level was mainly elucidated by light microscopy of fixed and stained chromosome preparations. Except for certain favorable species, it was difficult to study the complex process of homolog recognition and pairing, which is a central unique feature of meiosis. Furthermore, recombination could only be estimated by counts of chiasmata (which is not always easy because of hyper condensation of the chromosomes at the stage when chiasmata are visible) and/or, in favorable organisms, genetic analyses of the resulting gametes. A fantastic tool to study the early meiotic stages was provided by the discovery of the evolutionarily well-conserved meiotic pairing-structure called the Synaptonemal Complex by MJ Moses and DW Fawcett [4, 5].

Fig. 1 (A) Picture of D. von Wettstein at the electron microscope (from Ref. 89). (B) Electron micrograph of the Synaptonemal Complex (SC) of Neotiella. The tripartite SC is formed by two striped lateral elements (LE) and a central element located at equal distance from the two LEs. Chromatin of the parental homologous chromosomes is visible on both sides of the SC (from Ref. 7). (A, D) Two examples of interlockings. (C) Electron micrograph of a spread nucleus of Bombyx with one bivalent trapped in the other bivalent that is synapsing (arrow) (from Ref. 25). (D) Drawing of two entangled bivalents (large arrow) that are partially synapsed. One of them already shows late recombination nodules (small arrows). Bars at the end of the chromosomes indicate their attachment to the nuclear envelope; section numbers are shown (from Ref. 35). (E) Drawing of SC formation in a triploid Bombyx female: the SC joins two of the three homologous chromosomes while the third one remains as a single chromosome. Note that their telomeres are attached to the nuclear envelope (black line) in close proximity. Numbers indicate the section number (from Ref. 31). (F) Electron micrograph of a spread wheat nucleus at zygotene. This part of the nucleus shows a mixture of already synapsed regions (SC, arrows) and still completely separated chromosomes. The image also illustrates the difficulty to trace chromosomes one by one (from Ref. 27). (G) Electron micrograph of the Sordaria humana SC with a late Recombination Nodule (RN, arrow) located in the CE. It is placed under the Neotiella SC to show the conservation in size and structure of the two SCs, with the exception that Sordaria LEs are not striped and their chromatin is less dense (from Ref. 59)

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The synaptonemal complex (SC), which forms between homologous chromosomes during pairing, is a proteinaceous tripartite structure, which can be easily silver stained or contrasted by conventional electron microscopy (EM) methods. Preceding the appearance of the SC, each chromosome is associated with an axial element (AE). Components of the SC central element then assemble progressively between two homologous AEs and, by extension allow the complete synapsis of homologs (Fig. 1B). The SC is disassembled at the end of pachytene, except at the chiasma sites. This progressive formation of both axial and central elements allows one to follow not only complete SCs at pachytene, but also individual chromosomes during all preceding steps of homolog recognition and pairing, even if the surrounding chromatin remains too diffuse to allow a clear recognition of the chromosomes. Two additional types of dense structures/nodules are present in the central region of the SC. Those present at zygotene are generally small and irregular in shape and their role still remains to be defined. The nodules present at pachytene are larger and correspond in number, and distribution along the homologs, to the sites of crossovers (reviews in Refs. 1, 6; below). Studying the SC components, therefore, allows not only the characterization of chromosome behavior during pairing, but also, through analysis of nodules, quantification of the number and localization of reciprocal exchanges occurring in each nucleus (below). Mogens and Diter saw the SC as the tool with which one could quickly answer fundamental questions about chromosome pairing [7–11]. They rapidly identified important areas of research that were either understudied or had not been examined at all. At the time, no SC proteins were known, prohibiting the use of antibodies or fluorescent–tags to the corresponding proteins, as currently used now (reviews in Refs. 3, 12). Furthermore, the universal 100 nm width of the SC does not allow its study by conventional light microscopy. Therefore, Diter, Søren Rasmussen and Preben Holm developed the only experimental system that could answer the critical questions of the time: serial sectioning of entire nuclei followed by three-dimensional reconstruction of SCs or unsynapsed AEs from electron micrographs (discussion in Refs. 13, 14). The serial-sectioning technique, although time-consuming, currently remains the best to give the exact and relative positions of all chromosomes in a nucleus, and to reveal changes in their disposition and morphology during meiotic prophase (e.g., reviews in Refs. 1, 2, 15–17). The group also used surface spreading techniques for EM (developed by Montrose Moses [18]) in which stained nuclear contents are displayed on a surface. Instead of the almost 6 months necessary to reconstruct a single zygotene nucleus of hexaploid wheat (with 42 chromosomes, often associated as multivalents), this technique enabled all the chromosomes of a nucleus to be observed at once in a two-dimensional view

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[19–24]. Rupture of the nuclear envelope by spreading, however, disrupts the three-dimensional arrangements of the chromosomes. The nodules present in the central region of the SC are also no longer easily identifiable in the spread silver-stained SCs. Despite these drawbacks the technique permitted the analysis of many more nuclei than could be analyzed ultrastructurally from serial sections to study specific problems such as chromosome entanglements/ interlockings during pairing (e.g., [25, 26]), SC patterns in polyploid plants like hexaploid wheat (e.g., [19–22]), haploid meiosis [24], and hybrids [27]. Even by today’s standards, the achievements done in Diter’s laboratory are astonishing. From 1972 to 1988, over 1700 meiotic nuclei from the following organisms were serially sectioned, observed by EM, printed on paper, their AEs and SCs reconstructed (from mostly over 100 micrographs for each nucleus), the behavior of each chromosome analyzed, and over 40 papers published (see the reference list). – Common fruit fly Drosophila melanogaster [28, 29]. – Silk worm Bombyx mori female [30–33] and male [25, 26, 34, 35]. – Human Homo sapiens spermatocytes [16, 36–44] and oocytes [45, 46]. – Mouse Mus musculus spermatocytes [47]. – Easter lily Lilium longiflorum [48, 49]. – Barley Hordeum vulgare [50]. – Corn Zea mays [51]. – Common wheat Triticum aestivum [19–24, 27, 52–54]. – Discomycete Neotiella rutilans [7, 8]. – Ascomycete Neurospora crassa [55]. – Ascomycete, budding yeast Saccharomyces cerevisiae [56–58]. – Ascomycete Sordaria humana [59]. – Basidiomycete Coprinus cinereus [60, 61]. – Basidiomycete, commune [62].

splitgill

mushroom

Schizophyllum

This was made possible thanks to the enthusiastic and supportive help of Diter (who never put his name on the papers, despite continuous critical reading and suggestions) plus the stimulating atmosphere and facilitating environment of the laboratory. Also, Søren, Preben and Jane Sage, the fantastically competent engineer of the laboratory, devised an ingenious method for collecting serial sections on a grid, along with a remarkably simple, very efficient little tool to stain several grids together. Computerization initiated by Søren allowed easier quantitative analysis of the series of

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micrographs, Ann-Sofi Steinholz printed electron micrographs, Nina Rasmussen made most of the drawings, Inge Sommer typed all manuscripts and Bent Hansen maintained and repaired the very busy electron microscope. All together provided a warm social climate, where most scientific or technical difficulties were solved in long animated discussions often sustained on Friday by the free Carlsberg beer. Several post-doctorates and researchers, such as Chris Gillies, Denise Zickler, Maya Bojko, Ben Lu, Glyn Jenkins, Bente Wischmann, Joakim Glamann, Pheya Carmi, Yigal Koltin, Palle Hobolth, and Xingzhi Wang (I apologize for possible missing names), spent some time in the laboratory to work on their own project or in collaboration with Søren and Preben on various projects. Most have gone on to build successful careers in research and teaching in academia or industry. The insights obtained from the “Carlsberg’s” work brought the technique and the results to the attention of researchers interested in meiosis. Diter therefore organized two EMBO workshops on “Chromosome pairing and crossing over” in 1978 and 1979 at the Carlsberg Laboratory. During 1 week, applicants not only learned how to fix, section, reconstruct, and analyze meiotic nuclei but also participated every day in a conference with invited “stars” of the field followed by discussions. This was the golden age of serial sectioning and new discoveries concerning the chromosome behavior during meiotic pairing and recombination. Succinctly listing all of the contributions of Diter’s laboratory to the many aspects of the meiotic process is difficult, but six main areas can be distinguished; all were important discoveries and, in most cases, were confirmed by genetic analyses of mutants, molecular approaches, and immunolocalization of recombination and SC proteins. First, analysis of 14 phylogenetically different organisms (above) revealed a remarkable similarity in the course of chromosome pairing. (1) The SC definitively mediates the intimate connection of homologs. (2) Before SC formation, chromosome ends are attached to the inner surface of the nuclear envelope and these attachment sites are initially distributed over most of the nuclear periphery. (3) Ends next become redistributed via chromosome movements into a limited region of the nuclear envelope, defining the “bouquet stage” described in the early 1900s [63] and now shown to be a universal feature of meiosis (reviews in Refs. 1, 3, 15, 64). (4) The bouquet stage is followed by redistribution of chromosome ends evenly around the nuclear periphery. (5) Although SC initiation is mostly observed at chromosome ends, the juxtaposition of homologs is not a simple zippering from one chromosome end, but in most organisms (especially with long chromosomes) short stretches of SCs are formed at several independent places along the homologs (e.g., [45, 49]). (6) At the end of pachytene, the SC is degraded, leaving only remnants at diplotene (reviews in Refs. 1, 15, 16, 37, 44).

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Second, Diter’s group showed for the first time that formation of entanglements/interlocks of unrelated chromosomes is a regular pairing step in all organisms (Figs. 1C, D). First described by Gelei [65] in the flatworm Dendrocoelum lacteum, such interlocks were considered as rare events visible only at diplotene/metaphase I, thus after pairing. However, the pre-SC random distribution of homologous chromosomes in the nucleus combined with chromosome movement during homology recognition and the fact that SC initiates in more than one site along homologs provide a high potential risk of chromosomal entanglements. Indeed, three-dimensional reconstructions from serial sections revealed clearly that two types of interlockings occur during pairing. Either one chromosome (unsynapsed or one homolog of a partially synapsed bivalent, Fig. 1C) or a pair of homologs (partially or fully synapsed) is trapped in a loop between two synapsed stretches of another bivalent (reviews in Refs. 1–3, 15, 16, 26). Interlocks are rare in organisms with small chromosomes (e.g., Coprinus, [60]) and more frequent in organisms with longer chromosomes (up to 20 per nucleus in wheat, [17, 19]). While routinely observable during the first steps of pairing, interlocks are almost never detected when the SC is completely formed at pachytene, thus explaining their paucity in light microscopic studies. A mechanism for interlock resolution must therefore exist. In the fungus Sordaria macrospora, recombination protein Mlh1 (the eukaryotic homologue of the bacterial mismatch repair protein MutL), which is implicated in finalization of crossover recombinational interactions, is also required for interlock resolution. Mlh1 likely eliminates constraining recombination connections formed by the recombination process on the SCs located on both sides of the entanglement site [66] (Fig. 1D). Because frequent AE and SC interruptions were observed in association with interlocks, Diter, Preben, and Søren suggested that interlocks are resolved by breakage and rejoining of chromosomes or bivalents through Type II Topoisomerase that can pass one DNA duplex through another [1, 26]. This particularly fine example of Diter’s prescience was recently proven correct in Arabidopsis by analyses of hypomorph topII mutants: interlockings can indeed be resolved by TOPII activity, but only in addition with chromosome movement [67]. Third, they showed that pairing patterns sometimes change over time resulting in two-phase pairing/SC formation. In all studied organisms (above), SC formation occurs preferentially between homologous partners. This is best illustrated by pairing of reciprocal translocations (where chromosome segments are exchanged between two nonhomologous chromosomes): the observed SC configurations correspond to those predicted by underlying chromosomal homology (as illustrated by SC formation in the translocation quadrivalents of Coprinus [60]).

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These constraints can change overtime. (1) Straight bivalents sometimes replace the former quadrivalents in Coprinus in absence of crossovers on both sides of the pairing-partner exchange ([60]; review in Ref. 1). (2) In presence of inversions or duplications of chromosome segments on one homolog, the corresponding bivalents usually exhibit first only homologous SC formation (with the inverted or duplicated region forming a loop); this configuration is followed by nonhomologous SC formation after synaptic adjustment at late pachytene. Also, in a human XYY male, the two Y chromosomes were found paired by the SC beyond the PAR homology segment normally synapsed by the SC between X and Y chromosomes [43]. All these observations imply early SC destabilization in the vicinity of the rearrangement discontinuity followed by reformation of the SC that no longer reflects the underlying nonhomology (review in Ref. 14; other examples in Refs. 68, 69 and for a human reciprocal translocation in [39]). (3) Extensive nonhomologous SC formation is also observed in haploid meioses (e.g., [50]), indicating a “driving force” that seeks for maximization of SC formation when the homologous partner is missing. (4) An interesting case of changes in SC patterns was discovered during the study of triploid females of silk worm (Bombyx mori), which provides an example of competition between three homologous chromosomes (S. Rasmussen [31], in collaboration with B.L. Astaurov, Moscow). Before SC formation, the three homologous chromosome ends are attached to the nuclear envelope in close proximity (Fig. 1E). The SC initially forms in the telomere regions, pair wise between all three possible homologous partners, including double SCs at any place along the chromosomes. However, at the end of pachytene, nuclei contain only bivalents and univalents (Fig. 1E), which often, in turn, undergo foldback SC formation or associate with one or more nonhomologous chromosomes with SC morphologies indistinguishable from those of homologous SCs. This result clearly indicates that this second round of SC formation is no longer dependent on homology. Similarly, in autotetraploid Bombyx females, the quadrivalents observed during the early synapsis steps are mostly replaced later by bivalents. This is a perspicuous indication of a two-phase SC formation that, moreover, results in this case in viable gametes [33]. Such correction of pairing and SC formation again involves SC destabilization plus reformation between different chromosomes. This mechanism is only observed in females, which lack crossover and chiasma formation. In Bombyx males, by contrast, which follow the standard program of crossover formation [35] and thus recombination-dependent pairing and SC formation, SC correction is less efficient or nonexistent and trivalents or quadrivalents persist until their segregation at metaphase I [34]. In triploid Coprinus with a standard crossover program, trivalents are often synapsed by two SC central regions combining the three

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homologous axes along their entire length [61]. SC formation, therefore, can involve all three homologs at the same sites. Such configurations are maintained unaltered throughout pachytene [61]. In Bombyx male and in Coprinus, occurrence of crossovers has been proposed to prevent transformation of multivalents into bivalents, thus leading to infertility (reviews in Refs. 1, 15, 16). In conclusion: correction and successive nonhomologous SC formation depending upon crossover formation is likely a regular feature of the meiotic prophase. These findings also indicate that at early steps of SC formation homology is predominant, but that at later stages this restrictive condition is alleviated, allowing SC formation and progress in nonhomologous segments, therefore maximizing a continuous SC. SC formation in diploid, triploid or tetraploid Bombyx females remains a mystery. In most organisms including Bombyx males, which form crossovers, DNA–DNA recombinationrepair interactions after programmed double-strand breaks, mediate the recognition and pairing of homologous chromosomes plus SC initiation (review in Ref. 3). In absence of DNA double-strand breaks in Bombyx females, other mechanisms likely take place (e.g., homologous recognition at the telomere regions like in the worm Caenorhabditis elegans, [70]). These phenomena remain a fascinating subject for the future. Fourth, ultrastructural analysis of common wheat Triticum aestivum provided the first complete picture of how this allohexaploid pairs and synapses. Many naturally evolved and/or cultivated plants contain multiple genome complements, which may be genetically identical (homologous) or divergent from one another to various degrees (homoeologous). They arise either by multiplication of a basic set of chromosomes (autopolyploidy) or result from combinations of related but not completely homologous genomes (allopolyploidy). This raises an intriguing issue because pairing can occur between homologous and homoeologous chromosomes when restriction of pairing to homologous chromosomes is a prerequisite for reproductive fertility and thus stability in these polyploids. The common wheat is an allohexaploid (2n ¼ 6x ¼ 42 chromosomes) combining three diploid genomes (AA BB DD) of T. urartu, T. speltoides, and T. tauschii. Breeding and genetic analyses have shown that each pair of chromosomes is partly homoeologous to a chromosome pair in each of the two other genomes (Zhang et al, among whom Diter von Wettstein, [71]). In spite of this close relationship among the three genomes, crossovers and thus chiasmata form only between homologs and never between homoeologs, the outcome being the formation of 21 bivalents at metaphase I (review in Refs. 17, 19, 71). To test how those bivalents are formed, Palle Hobolth [52], Glyn Jenkins [53], and Preben Holm [17, 19] undertook an ultrastructural analysis by serial sectioning (Hobolth and Jenkins) or spreading (Holm; Fig. 1F) of early prophase nuclei and discovered

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several interesting and intriguing behaviors. First, the chromosomes are not prealigned when meiosis starts, excluding a premeiotic pairing mechanism to explain the formation of regular metaphase I bivalents. Second, at early prophase, extensive multivalents form with multiple pairing partner shifts, indicating homology recognition and SC formation among homoeologs as well as homologs. Third, SC initiation at both chromosome ends and interstitially generates multiple entanglements /interlockings (above), which are partially repaired at pachytene. Finally, by the end of synapsis, however, SC formation is restricted to homologs, thus revealing again (as in the Coprinus and Bombyx polyploids, above) the presence of a two-phase SC formation. The presence of only bivalents indicates that the transformation of multivalents into bivalents happens before crossover occurrence or that crossover formation is delayed until the completion of the second phase of SC correction and/or indicates the presence of a mechanism that suppresses crossovers between paired segments of homoeologs. The regular bivalent formation seen in common wheat is not observed when lines having multiple duplicated genome complements are generated de novo by plant breeding (review in Refs. 71, 72). This indicates that special stringency determinants have developed in the genomes of the natural allopolyploids as in common wheat, enhancing their viability. Among the genetic determinants identified is the Pairing homoeologous 1 (Ph1) locus discovered in 1958 based on the observation that wheat plants lacking chromosome 5B exhibited homoeologous bivalent formation (review in Ref. 73). The locus is present only in the B genome on the long arm of chromosome 5. When present as two copies, although multivalents are observed during SC formation (above), only bivalents are formed at metaphase I, and crossovers/chiasmata are restricted to homologs (e.g., [74], reviews in Refs. 2, 17, 19–22). Further studies of the role of the Ph1 locus by Diter’s group comprise analyses of wheats (1) nullisomic for chromosome 5B [21], (2) monosomic for 5B [22], (3) those in which chromosome 5B was replaced by one, two, or three copies of an isochromosome for the long arm of chromosome 5B [21], (4) those carrying an isochromosome for the long arm of 5B, and (5) wheat–rye hybrids nullisomic or monosomic for 5B (review in Ref. 23) as well as (6) haploid wheat [24]. When the Ph1 locus (or the arm of chromosome 5B) is absent, the number of multivalents increase as well as the number of pairing-partner exchanges and chiasmata are no longer solely restricted to homologs; moreover, synapsis is never complete and multivalents remain unresolved until metaphase I [21]. When six copies of the locus or of 5B are present, by contrast, primarily only univalents are observed at metaphase I, indicating a strong reduction in the number of crossovers even among the homologs. However, SC form, although incompletely and the high frequency of interlockings indicates that SC form between

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homologs and homoeologs (review in Refs. 15, 17; Sidhu et al, among whom Diter von Wettstein [75]). In the hexaploid, trihaploid and wheat–rye hybrids, the increase in the number of pairingpartner exchanges in absence of chromosome 5B is observed very early and cannot be attributed to differences in the two-phase SC correction process. Absence of 5B likely results, therefore, in relaxation of the homolog recognition and/or synapsis stringency. The conclusion of these studies was that both increases and decreases in the copy number of chromosome 5B, or of Ph1 containing regions, augment homoeologous and even nonhomologous chromosome SC formation, while the diploid number of the Ph1 locus ensures correction of multivalents through dissolution and reassembly of the SC and finally exclusive bivalent formation (reviews in Refs. 15, 17, 75). However, the presence of the Ph1 locus in nascent allohexaploid wheat (identical in genome constitution to common wheat) is not sufficient to ensure strict homologous meiotic pairing. This results in a wide range of chromosomal aneuploidy in these newly synthesized multiple wheat lines. Their persistence, even after consecutive selection for euploidy over multiple generations, suggests that additional mechanisms are required to stabilize the postpolyploidization generations [71]. Several types of hypotheses have been advanced to account for the pairing and recombination behavior of allohexaploid wheat and especially for the role of Ph1. Holm and Wang [23] suggested that the Ph1 locus controls the stringency of both pairing and crossover formation/suppression between homologs. This control could involve the DNA mismatch repair process [2]. Indeed, in budding yeast, homoeology of 1% reduces crossovers several fold, and this effect is reversed by elimination of mismatch repair functions (e.g., [76]). Recent cloning and sequencing of the Ph1 locus define this locus as containing heterochromatin with a ZIP4-B2 paralog inserted into a cluster of Cdk2-like genes (cell division cycle 2) interspersed with methyl transferase genes. The Zip4 homologues are also located on chromosomes 3A, 3B and 3D [77]. In budding yeast, Zip4 is required for both SC and crossover formation, while in plants, the protein is especially required for crossover formation (review in Ref. 3). The presence of a ZIP4 paralog in the Ph1 locus is thus interesting, but more studies will be necessary to know the exact role, if any, of this inserted ZIP4-like gene. Several other candidate genes (e.g., C-Ph1 identical in chromosomes 5A and 5D, but absent in 5B) have been described (Bhullar et al. among whom Diter von Wettstein [78]). ASY1 (Asynapsis 1), a component of the SC lateral element shows abnormal localizations in absence of Ph1 [79]. ASY1 is essential for SC and crossover formation in Arabidopsis and rye: when absent or mutated, plants show homologous and nonhomologous SC formation [80]. Its abnormal localization in the ph1 mutant could destabilize the SC and consequently prevent the second-phase SC formation observed in

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wild-type wheat. The overall conclusion is that the molecular mode of action of the Ph1 locus remains elusive in part because of the complexity of the locus. Perturbation of any biochemical process required to implement SC formation will also reduce the efficiency of discrimination between homologous and homoeologous pairing/SC formation without being informative about the Ph1 function per se. Fifth, the hypothesis that the nodules associated with the SC central region are involved in meiotic recombination was extensively supported by Diter’s laboratory. Electron-dense, more or less ovoid structures were/are observed associated with forming and formed SC in all studied organisms (above, Fig. 1D, G). A correlation between these structures and crossovers was first recognized by Carpenter [81] in oocytes of D. melanogaster. Further investigations revealed two types of nodule-like structures along SCs. One class seen at pachytene correlates with crossover recombination events in all studied organisms and is thus called late nodules (LNs) or recombination nodules (RNs; Fig. 1G). The second class (called early nodules), essentially present in forming SCs, is smaller in size and more numerous than the later RNs and thus not correlated with the crossover/chiasma number of the organism. In accord with an implication of late RNs in the crossover process, female Bombyx lacking chiasma formation also lack RNs, while chiasmate males exhibit RNs along their homologs [82]. The correlation between crossovers and RNs was subsequently confirmed by immunolocalization of several recombination proteins (including Mlh1, above). The central components of the SC are required for the maintenance and turnover of the recombination proteins participating in maturation of the double-strand breaks into crossovers (review in Ref. 3). Systematic analysis of the number and distribution of RNs through meiotic prophase in human spermatocytes [36], human oocytes [46], Bombyx spermatocytes [35] and Coprinus [60] showed that early nodules are not only more numerous than late nodules but that they are also evenly distributed along homologs. This is in marked contrast to late RNs, which are never randomly distributed (below). In triploid Coprinus where SC formation between all three homologs is frequent, nodules associate with SC central regions sometimes at the same position [60]. Diter’s group confirmed that late RNs reflect the operation of a crossover regulation system. (1) Whatever the size of the chromosome, there is always at least one late RN per homolog pair. (2) Both types of nodules are found predominantly in euchromatin and are rare or absent from centromere regions and heterochromatin. (3) The remaining fragments of SCs seen at diplotene mostly contain one RN, and their global number corresponds to the number of chiasmata seen by light microscopy. (4) In Bombyx, Coprinus and human, RNs are further converted into chromatin-chiasma

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structures that remain visible up to prometaphase I. (5) Although late RNs tend to occur at different positions along chromosomes in different nuclei, when two or more RNs are seen along one homolog they are spaced well apart as are crossovers. This results from the process of interference, discovered during genetic analyses of Drosophila recombination by Sturtevant [83] and Muller [84]. Namely, a crossover occurring at one position along a chromosome reduces the probability that another crossover will occur nearby (reviews in Refs. 1–3). In all studied organisms, furthermore, a tight correlation exists between the SC length of a bivalent and the number of RNs per bivalent (review in Ref. 85). This correlation is also observed when male and female behaviors are compared. Human female SCs are twice as long as male SCs, with an accompanying slight increase in the number of RNs [41–43, 46]. Further analyses of several hundred nuclei by immunocytology of Mlh1 foci, which mark the sites of crossovers and colocalize with late RNs in EM spreads, revealed that the number of Mlh1 foci in females is double the level of foci seen in male (e.g., [86, 87]). Sixth, the lack of crossover can be compensated by retention of the SC for correct homolog segregation. In nearly all organisms, the formation of crossovers/chiasmata ensures a link between homologs until their segregation at anaphase I. This is observed in Bombyx diploid males [35]. Bombyx diploid females, by contrast, lack crossover and chiasmata [32, 88]. Nonetheless, both sexes exhibit normal SCs. The same asymmetry for crossover formation between female and male (shown above for Bombyx) is also observed in other insects, and in the studied cases, both sexes form normal SC at pachytene (reviews in Refs. 2, 30, 32). However, instead of SC depolymerization after crossover completion as in chiasmate males, in the achiasmate females, SCs are maintained until homolog segregation at anaphase I. The remaining SCs are either conserved in their original form or transformed into a dense rod to which homologous chromosomes remain attached [30, 88]. The Bombyx study provided for the first time an explanation for the correct homolog segregation observed by light microscopy in achiasmate organisms or sexes: the SC (modified or not) can provide a substitute connection (thus assuming the function of chiasmata) as required for the maintenance of homologs until their regular disjunction at anaphase I. When I arrived at Carlsberg for my postdoctoral training in October 1973, to study budding-yeast meiosis, I was impressed by Diter’s broad thinking, focus on key questions and adaptability. Being a passionate scientist he had high expectations of himself and people around him. Moreover, Diter’s energy seemed boundless in both research and parties. At meetings, he was able to dance all night and nevertheless participative actively during all presentations the next day. I am, of course, indebted to Diter and Mogens for a fantastic postdoctoral experience and sweet evenings shared

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with their wives Penny von Wettstein-Knowles and Ebba Westergaard. I am also indebted to Søren and Preben for providing me with so much pleasure during our long discussions in the laboratory and during our numerous parties, which both illuminated the long fall and winter evenings. Overall, my postdoctoral experience in Diter’s laboratory comforted my idea that meiosis provided a wonderful playing field of research, which I still enjoy, and that a warm social climate in the laboratory is almost as important as tireless work.

Acknowledgments I thank Penny von Wettstein-Knowles and Sachin Rustgi for comments that improved the manuscript. I am grateful and much indebted to Preben Holm and Søren Rasmussen for their longstanding warm friendship and for their steady encouragement and critical reading of the manuscript. We all three would like to thank the editors for this opportunity to reflect on ways that Diter von Wettstein affected and enriched our lives. References 1. von Wettstein D, Rasmussen SW, Holm PB (1984) The synaptonemal complex in genetic segregation. Annu Rev Genet 18:331–413 2. Zickler D, Kleckner N (1999) Meiotic chromosomes: integrating structure and function. Annu Rev Genet 33:603–754 3. Zickler D, Kleckner N (2015) Recombination, pairing, and synapsis of homologs during meiosis. Cold Spring Harb Perspect Biol 7: a016626 4. Moses MJ (1956) Chromosome structures in crayfish spermatocytes. J Biophys Biochem Cytol 2:215–218 5. Fawcett DW (1956) The fine structure of chromosomes in the meiotic prophase of vertebrate spermatocytes. J Biophys Biochem Cytol 2:403–406 6. Carpenter ATC (1988) Thoughts on recombination nodules, meiotic recombination and chiasmata. In: Kucherlapati R, Smith GR (eds) Genetic recombination. American Society for Microbiology, Washington, DC, pp 549–574 7. Westergaard M, von Wettstein D (1970) Studies on the mechanism of crossing over. IV. The molecular organization of the synaptinemal complex in Neottiella (Cooke) saccardo (Ascomycetes). C R Trav Lab Carlsberg 37:239–268

8. Westergaard M, von Wettstein D (1972) The synaptinemal complex. Annu Rev Genet 6:71–110 9. von Wettstein D (1971) The synaptinemal complex and the four-strand crossing over. Proc Natl Acad Sci U S A 68:851–855 10. von Wettstein D (1977) The assembly of the synaptinemal complex. Phil Trans R Soc Lond B 277:235–243 11. Stern H, Westergaard M, von Wettstein D (1975) Presynaptic events in meiocytes of Lilium longiflorum and their relation to crossing-over: a preselection hypothesis. Proc Natl Acad Sci U S A 72:961–965 12. Gao J, Colaia´covo MP (2018) Zipping and unzipping: protein modifications regulating Synaptonemal Complex dynamics. Trends Genet 34:232–245 13. Holm PB, Rasmussen SW, von Wettstein D (1979) The possible contribution of electron microscopy to the understanding of the mechanism of non-disjunction in man. Mutat Res 61:115–119 14. Holm PB, Rasmussen SW, von Wettstein D (1982) Ultrastructural characterization of the meiotic prophase. A tool in the assessment of radiation damage in man. Mutat Res 95:45–59

Necrology of Prof. Diter von Wettstein: Part II 15. Rasmussen SW, Holm PB (1980) Mechanics of meiosis. Hereditas 93:187–216 16. Rasmussen SW, Holm PB (1984) The synaptonemal complex, recombination nodules and chiasmata in human spermatocytes. Symp Soc Exp Biol 38:271–292 17. Holm PB (1986) Ultrastructural analysis of meiotic recombination and chiasma formation. Tokai J Exp Clin Med 11:415–436 18. Moses MJ (1977) Synaptonemal complex karyotyping in spermatocytes of the Chinese hamster (Cricetulus griseus). I. Morphology of the autosomal complement in spread preparations. Chromosoma 60:99–125 19. Holm PB (1986) Chromosome pairing and chiasma formation in allohexaploid wheat, Triticum aestivum, analyzed by spreading of meiotic nuclei. Carlsberg Res Commun 51:239–294 20. Holm PB (1988) Chromosome pairing and synaptonemal complex formation in hexaploid wheat, monosomic for chromosome 5B. Carlsberg Res Commun 53:57–89 21. Holm PB (1988) Chromosome pairing and synaptonemal complex formation in hexaploid wheat, nullisomic for chromosome 5B. Carlsberg Res Commun 53:91–110 22. Holm PB (1988) Chromosome pairing and synaptonemal complex formation in hexaploid wheat, monoisosomic and diisosomic for the long arm of chromosome 5B. Carlsberg Res Commun 53:111–133 23. Holm PB, Wang X (1988) The effect of chromosome 5B on synapsis and chiasma formation in wheat, Triticum aestivum cv. Chinese Spring. Carlsberg Res Commun 53:191–208 24. Wang X (1988) Chromosome pairing analysis in haploid wheat by spreading of meiotic nuclei. Carlsberg Res Commun 53:135–166 25. Rasmussen SW (1986) Initiation of synapsis and interlocking of chromosomes during zygotene in Bombyx spermatocytes. Carlsberg Res Commun 51:401–432 26. Rasmussen SW (1986) Chromosome interlocking during synapsis – a transient disorder. Tokai J Exp Clin Med 11:437–451 27. Wang X, Holm PB (1988) Chromosome pairing and synaptonemal complex formation in wheat-rye hybrids. Carlsberg Res Commun 53:167–190 28. Rasmussen SW (1973) Ultrastructural studies of spermatogenesis in Drosophila melanogaster. Z Zellforsch 140:125–144 29. Rasmussen SW (1975) Synaptonemal polycomplexes in Drosophila melanogaster. Chromosoma 49:321–331

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30. Rasmussen SW (1977) The transformation of the synaptonemal complex into the “elimination chromatin” in Bombyx mori oocytes. Chromosoma 60:205–221 31. Rasmussen SW (1977) Chromosome pairing in triploid females of Bombyx mori analyzed by three dimensional reconstructions of synaptonemal complexes. Carlsberg Res Commun 42:163–197 32. Rasmussen SW (1977) Meiosis in Bombyx mori females. Philos Trans R Soc Lond B Biol Sci 277:343–350 33. Rasmussen SW, Holm PB (1979) Chromosome pairing in autotetraploid Bombyx mori females. Mechanism for exclusive bivalent formation. Carlsberg Res Commun 44:101–125 34. Rasmussen SW (1987) Chromosome pairing in autotetraploid Bombyx males. Inihibition of multivalent correction by crossing over. Carlsberg Res Commun 52:211–242 35. Holm PB, Rasmussen SW (1980) Chromosome pairing, recombination nodules and chiasma formation in diploid Bombyx males. Carlsberg Res Commun 45:483–548 36. Rasmussen SW, Holm PB (1978) Human meiosis II: chromosome pairing and recombination nodules in human spermatocytes. Carlsberg Res Commun 42:275–327 37. Rasmussen SW, Holm PB (1978) Human meiosis IV: the elimination of synaptonemal complex fragments from metaphase I bivalents of human spermatocytes. Carlsberg Res Commun 43:423–438 38. Holm PB, Rasmussen SW (1977) Human meiosis I. The human pachytene pachytene karyotype analyzed by three dimensional reconstruction of the synaptonemal complex. Carlsberg Res Commun 42:283–323 39. Holm PB, Rasmussen SW (1978) Human meiosis III. Electron microscopical analysis of chromosome pairing in an individual with a balanced translocation 46,XY,t(5p;22p+). Carlsberg Res Commun 43:329–350 40. Holm PB, Rasmussen SW (1983) Human meiosis V. Substages of pachytene in human spermatogenesis. Carlsberg Res Commun 48:351–383 41. Holm PB, Rasmussen SW (1983) Human meiosis Vl. Crossing over in human spermatocytes. Carlsberg Res Commun 48:385–413 42. Holm PB, Rasmussen SW (1983) Human meiosis VII. Chiasma formation in human spermatocytes. Carlsberg Res Commun 48:415–456 43. Berthelsen JG, Holm PB, Rasmussen SW (1980) Three ultrastructure markers on pachytene bivalents of human spermatocytes. Carlsberg Res Commun 45:25–28

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44. Holm PB, Rasmussen SW (1984) The synaptonemal complex in chromosome pairing and disjunction. In: Bennett MD, Gropp A, Wolf U (eds) Chromosomes today, vol 8. Georg Allen & Unwin, Crows Nest, NSW, pp 104–116 45. Bojko M (1983) Human meiosis. VIII. Chromosome pairing and formation of the synaptonemal complex in oocytes. Carlsberg Res Commun 48:457–483 46. Bojko M (1985) Human meiosis. IX. Crossing over and chiasma formation in oocytes. Carlsberg Res Commun 50:43–72 47. Glamann J (1986) Crossing over in the male mouse as analysed by recombination nodules and bars. Carlsberg Res Commun 51:143–161 48. Holm PB (1977) The premeiotic DNA replication of euchromatin and heterochromatin in Lilium longiflorum (Thunb.). Carlsberg Res Commun 42:249–281 49. Holm PB (1977) Three-dimensional reconstruction of chromosome pairing during the zygotene stage of meiosis in Lilium longiflorum (Thunb.). Carlsberg Res Commun 42:103–151 50. Gillies CB (1974) The nature and extent of synaptonemal complex formation in haploid barley. Chromosoma 48:441–453 51. Mogensen HL (1977) Ultrastructural analysis of female pachynema and the relationship between synaptonemal complex length and crossing-over in Zea mays. Carlsberg Res Commun 42:475–497 52. Hobolth P (1981) Chromosome pairing in allohexaploid wheat var. Chinese Spring. Transformation of multivalents into bivalents, a mechanism for exclusive bivalent formation. Carlsberg Res Commun 46:129–173 53. Jenkins G (1983) Chromosome pairing in Triticum aestivum cv. Chinese Spring. Carlsberg Res Commun 48:255–283 54. Wischmann B (1986) Chromosome pairing and chiasma formation in wheat plants trisomic for the long arm of chromosome 5B. Carlsberg Res Commun 51:1–25 55. Gillies CB (1972) Reconstruction of the Neurospora crassa pachytene karyotype from serial sections of synaptonemal complexes. Chromosoma 36:119–130 56. Zickler D, Olson LW (1975) The synaptonemal complex and the spindle plaque during meiosis in yeast. Chromosoma 50:1–23 57. Petersen JGL, Olson LW, Zickler D (1978) Synchronous sporulation of Saccharomyces cerevisiae at high cell concentrations. Carlsberg Res Commun 43:241–253 58. Zickler D (1981) Ultrastructure of the yeast nucleus. In: Gull K, Oliver SG (eds) The fungal

nucleus, British mycological society symposium 5. Cambridge University Press, Cambridge, pp 63–83 59. Zickler D, Sage J (1981) Synaptonemal complexes with modified lateral elements in Sordaria humana: development of and relationship to the “recombination nodules”. Chromosoma 84:305–318 60. Holm PB, Rasmussen SW, Zickler D, Lu BC, Sage J (1981) Chromosome pairing, recombination nodules and chiasma formation in the basidiomycete Coprinus cinereus. Carlsberg Res Commun 46:305–346 61. Rasmussen SW, Holm PB, Lu BC, Zickler D, Sage J (1981) Synaptonemal complex formation and distribution of recombination nodules in pachytene trivalents of triploid Coprinus cinereus. Carlsberg Res Commun 46:347–360 62. Carmi P, Holm PB, Koltin Y, Rasmussen SW, Sage J, Zickler D (1978) The karyotype of Schizophyllum commune analyzed by three dimensional reconstruction of synaptonemal complexes. Carlsberg Res Commun 43:117–132 63. Eisen G (1900) The spermatogenesis of Batrachoseps. J Morphol 17:1–117 64. Zickler D, Kleckner N (1998) The leptotenezygotene transition of meiosis. Annu Rev Genet 32:619–697 65. Gelei J (1921) Weitere Studien u¨ber die Oogenese des Dendrocoelum lacteum. II. Die L€angskonjungation der Chromosomen. Arch Zellforsch 16:88–169 66. Storlazzi A, Gargano S, Ruprich-Robert G, Falque M, David M, Kleckner N, Zickler D (2010) Recombination proteins mediate meiotic spatial chromosome organization and pairing. Cell 141:94–106 67. Martinez-Garcia M, Schubert V, Osman K, Darbyshire A, Sanchez-Moran E, Franklin FCH (2018) TOPII and chromosome movement help remove interlocks between entangled chromosomes during meiosis. J Cell Biol 217:4070–4079 68. Moses MJ, Poorman PA (1981) Synaptosomal complex analysis of mouse chromosomal rearrangements. II. Synaptic adjustment in a tandem duplication. Chromosoma 81:519–535 69. Moses MJ, Poorman PA, Roderick TH, Davisson MT (1982) Synaptonemal complex analysis of mouse chromosomal rearrangements. IV. Synapsis and synaptic adjustment in two paracentric inversions. Chromosoma 84:457–474 70. Rog O, Dernburg AF (2013) Chromosome pairing and synapsis during Caenorhabditis elegans meiosis. Curr Opin Cell Biol 25:349–356

Necrology of Prof. Diter von Wettstein: Part II 71. Zhang H, Bian Y, Gou X, Zhu B, Xu C, Qi B, Li N, Rustgi S, Zhou H, Han F, Jiang J, von Wettstein D, Liu B (2013) Persistent wholechromosome aneuploidy is generally associated with nascent allohexaploid wheat. Proc Natl Acad Sci U S A 110:3447–3452 72. Bomblies K, Jones G, Franklin C, Zickler D, Kleckner N (2016) The challenge of evolving stable polyploidy: could an increase in crossover interference distance play a central role? Chromosoma 125:287–300 73. Sears ER (1976) Genetic control of chromosome pairing in wheat. Annu Rev Genet 10:31–51 74. Feldman M (1966) The effect of chromosomes 5B, 5D and 5A on chromosomal pairing in Triticum aestivum. Proc Natl Acad Sci U S A 55:1447–1453 75. Sidhu GK, Rustgi S, Shafqat MN, von Wettstein D, Gill KS (2008) Fine structure mapping of a gene-rich region of wheat carrying Ph1, a suppressor of crossing over between homoeologous chromosomes. Proc Natl Acad Sci U S A 105:5815–5820 76. Borts RH, Haber JE (1987) Meiotic recombination in yeast: alteration by multiple heterozygosities. Science 237:1459–1465 77. Rey MD, Martı´n AC, Smedley M, Hayta S, Harwood W, Shaw P, Moore G (2018) Magnesium increases homoeologous crossover frequency during meiosis in ZIP4 (Ph1 gene) mutant wheat-wild relative hybrids. Front Plant Sci 9:509 78. Bhullar R, Nagarajan R, Bennypaul H, Sidhu GK, Sidhu G, Rustgi S, von Wettstein D, Gill KS (2014) Silencing of a metaphase I-specific gene results in a phenotype similar to that of the Pairing homeologous 1 (Ph1) gene mutations. Proc Natl Acad Sci U S A 111:14187–14192 79. Boden SA, Langridge P, Spangenberg G, Able JA (2009) TaASY1 promotes homologous chromosome interactions and is affected by deletion of Ph1. Plant J 57:487–497

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80. Sanchez-Moran E, Santos JL, Jones GH, Franklin FCH (2007) ASY1 mediates AtDMC1-dependent interhomolog recombination during meiosis in Arabidopsis. Genes Dev 21:2220–2233 81. Carpenter ATC (1975) Electron microscopy of meiosis in Drosophila melanogaster females. II. The recombination nodule-a recombination-associated structure at pachytene? Proc Natl Acad Sci U S A 72:3186–3189 82. Rasmussen SW, Holm PB (1982) The meiotic prophase in Bombyx mori. In: King RC, Akai H (eds) Insect ultrastructure, vol 1. Plenum Publishing Corp, New York, NY, pp 61–85 83. Sturtevant AH (1915) The behavior of the chromosomes as studied through linkage. Z Induct Abstammungs Vererbungsl 13:234–287 84. Muller HJ (1916) The mechanism of crossing over. Am Nat 50:193–434 85. von Wettstein D (1984) The synaptonemal complex and genetic segregation. Symp Soc Exp Biol 38:195–231 86. Gruhn JR, Rubio C, Broman KW, Hunt PA, Hassold T (2013) Cytological studies of human meiosis: sex-specific differences in recombination originate at, or prior to, esaishment of double-strand breaks. PLoS One 8: e85075 87. Wang S, Hassold T, Hunt P, White MA, Zickler D, Kleckner N, Zhang L (2017) Inefficient crossover maturation underlies elevated aneuploidy in human female meiosis. Cell 168:977–989 88. Rasmussen SW (1976) The meiotic prophase in Bombyx mori females analyzed by three dimensional reconstructions of synaptonemal complexes. Chromosoma 54:245–293 89. Holter H, Max Møller K (1976) The Carlsberg Laboratory 1876/1976, Rhodos International Science and Art Publishers, New York, NY 379

Part II Background and Overview

Chapter 3 A Short History and Perspectives on Plant Genetic Transformation Thakku R. Ramkumar, Sangram K. Lenka, Sagar S. Arya, and Kailash C. Bansal Abstract Plant genetic transformation is an important technological advancement in modern science, which has not only facilitated gaining fundamental insights into plant biology but also started a new era in crop improvement and commercial farming. However, for many crop plants, efficient transformation and regeneration still remain a challenge even after more than 30 years of technical developments in this field. Recently, FokI endonuclease-based genome editing applications in plants offered an exciting avenue for augmenting crop productivity but it is mainly dependent on efficient genetic transformation and regeneration, which is a major roadblock for implementing genome editing technology in plants. In this chapter, we have outlined the major historical developments in plant genetic transformation for developing biotech crops. Overall, this field needs innovations in plant tissue culture methods for simplification of operational steps for enhancing the transformation efficiency. Similarly, discovering genes controlling developmental reprogramming and homologous recombination need considerable attention, followed by understanding their role in enhancing genetic transformation efficiency in plants. Further, there is an urgent need for exploring new and low-cost universal delivery systems for DNA/RNA and protein into plants. The advancements in synthetic biology, novel vector systems for precision genome editing and gene integration could potentially bring revolution in crop-genetic potential enhancement for a sustainable future. Therefore, efficient plant transformation system standardization across species holds the key for translating advances in plant molecular biology to crop improvement. Key words Plant genetic engineering, Biolistic-mediated transformation, Agrobacterium-mediated transformation, Organelle transformation, CRISPR/Cas9

1

Introduction The cellular totipotency in the plant kingdom, the ability of Agrobacterium tumefaciens to infect plant cells and to genetically modify it, has been combined to open up a new field in plant biotechnology, as transgenic plant biology. Plant tissue culture and plant biotechnology have their foundations in the cell theory [1, 2] and the bacterial genetic transformation [3], respectively. Haberlandt— the father of tissue culture—established in vitro culture of plant

Sachin Rustgi and Hong Luo (eds.), Biolistic DNA Delivery in Plants: Methods and Protocols, Methods in Molecular Biology, vol. 2124, https://doi.org/10.1007/978-1-0716-0356-7_3, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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cells from multiple species and predicted to maintain and develop mature plant from those cells; however, his attempts did not succeed [4]. Later, plant callus with indefinite growth was achieved by three independent groups simultaneously [5–7]. Ball demonstrated the development of the whole plant in vitro using shoot tips of Tropaeolum majus and Lupinus albus [8]. On the other hand, Muir et al. established a technique for single cell cultures from callus [9, 10]. In parallel, Skoog and Miller established the hormonal control of organ development in plant cell cultures by fine-tuning and using a balanced ratio of auxins and cytokinins [11]. In this series, Steward and group demonstrated the development of somatic embryos that originated from a single cell and coined the term totipotency as the ability of single plant cell to develop into a whole plant [12–14]. Guha and Maheshwari reported that androgenic haploid embryos can be initiated using pollen grains of Datura [15]. Later, isolated protoplasts from Nicotiana glauca and N. langsdorffii were fused and interspecific somatic hybrid tobacco plants were regenerated [16]. In the genetic engineering field, the 1970s saw a new beginning with restriction enzymes. HindIII was the first restriction enzyme to be isolated and characterized from the bacterium Haemophilus influenzae. Smith and Nathan’s group made seminal contributions to this research [17–20]. These discoveries eventually led to the construction of the first recombinant (simian vacuolating virus 40) SV40 DNA by Jackson et al., with the insertion of DNA segments containing E. coli galactose operon and lambda phage genes [21]. However, the first recombinant plasmid in bacterium was introduced by Cohen et al., which is regarded as the first recombinant organism [22]. Meanwhile, two-dimensional electrophoresis of proteins was developed by O’Farrell [23]. Following this breakthrough, DNA sequencing technology was established by Sanger et al. [24]. And in 1983, Kary Mullis developed the method to amplify DNA fragments in vitro called a polymerase chain reaction (PCR) using DNA polymerase [25–27]. The Agrobacterium biology started with the identification of Agrobacterium tumefaciens, originally Bacterium tumefaciens, a gram-negative soilborne bacterium, as a causative agent of tumors in plant species [28]. Later studies by Braun and colleagues proved that the infecting pathogen A. tumefaciens is required only to initiate the plant infection but not for the tumor development, while the nutritional physiology of the infected cells is permanently altered, conceptualizing it as a tumor-inducing principle (TIP) [29, 30]. Later, Zaenen et al. reported that the gall formation is associated with unusually large tumor-inducing (Ti) plasmid [31]. Subsequently, transferred DNA (T-DNA) region was identified in the Ti plasmid, which was further characterized and found to be transferred from the A. tumefaciens Ti plasmid and integrated into the plant genome [32–35]. Zambryski et al. further identified

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that the T-DNA region contains specific border sequences—right border (RB) and left border (LB) and studied them in detail [35]. In the meantime, Drummond et al. reported that the T-DNA region is transcribed in the plant cell [36], while Gelvin et al. further proved that, the transcription rate was relatively lower in the bacterial cell and higher in the host plant cell [37]. Hoekema et al. were the first to use A. tumefaciens for plant genetic transformation [38]. However, development of the transgenic plant using A. tumefaciens was first reported by Herrera-Estrella et al. [39], and in quick succession by Fraley et al. [40]. A. tumefaciens can infect a broad range of dicot plants [41], while in monocots, Hiei et al. reported the successful transformation of rice a decade later [42]. Monocots are considered relatively recalcitrant [43], despite natural infection reported as early as in 1984 in members of Liliaceae and Amaryllidaceae [44]. Apart from plants, the A. tumefaciens, under specific laboratory conditions was also shown to genetically transform algae, fungi, and human cells, thus covering phylogenetically diverse kingdom [45–48]. Apart from A. tumefaciens mediated plant genetic transformation; several other bacterial species have been shown to genetically transform plants. For instance, A. rhizogenes, a closely related Agrobacterium species, that carries similar machinery and mechanism with Ri plasmid in place of the Ti plasmid as in A. tumefaciens, can infect the host plant root and genetically modify them, and induce hairy root phenotype. Surprisingly, bacterial species such as Ensifer adhaerens, Rhizobium etli, and Mesorhizobium loti, which are non-Agrobacterial but closely related species, are also shown to genetically transform plants if provided with T-DNA machinery. Viruses are also genetically manipulated to act as vectors to introduce foreign genetic material. These viral vector systems can be developed and applied to a wide variety of plants within a short span of experimental time; however, the viral transformations turned out to be transient. Cauliflower mosaic virus (CaMV) was the first virus to be studied elaborately and was proposed to act as an expression vector [49, 50]. Howell et al. reported that cloned copies of CaMV can be infectious to plants upon mechanical inoculation [51]. Grimsley et al. combined T-DNA machinery with CaMV viral genome to infect turnips and introduced a method called “agroinfection,” in which a modified binary vector with T-DNA carrying viral genome sequences was launched using A. tumefaciens [52]. Further, Grimsley et al. also shown that maize streak virus (MSV) based vectors can be used to agroinfect maize plants, a monocot plant, which are often considered recalcitrant to A. tumefaciens mediated transformation [53]. Following this, an RNA virus, the beet western yellows virus was successfully used to agroinfect the plants [54]. Viral vectors were also modified to initiate gene silencing, termed as virus-induced gene silencing (VIGS) and employed in reverse genetic approach for plant functional genomics [55, 56].

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Further, foreign DNA elements can be introduced by chemical or physical means. These direct gene transfer methods can be applied to diverse plant species as these methods do not depend on the plant–parasite interaction for gene delivery, as was the case with Agrobacterium. However, such direct gene transfer methods often lead to multicopy integration and scrambling of the foreign DNA element. The most preferred and commonly used nonbiological method for plant transformation is particle bombardment (also termed as biolistics). The particle bombardment was reported and successfully established for several recalcitrant plant species, especially cereals, quickly after the development of A. tumefaciens mediated transformed plants [57, 58]. This method is also universally used to target chloroplasts for genetic transformation.

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Agrobacterium-Mediated Plant Genetic Transformation The plant pathogenic Agrobacterium species A. tumefaciens and A. rhizogenes have evolved and uniquely equipped themselves to infect the host plant cells. Upon infection, these pathogens initiate the unidirectional transfer of desire DNA segment (transferred DNA/T-DNA) from their large plasmid leading to stable integration of the T-DNA into the host nuclear genome [59]. Agrobacterium’s potential for genetic transformation has been exploited by plant biologists to generate transgenic plants by modifying/inserting the foreign genes into the T-DNA region (Fig. 1). A. tumefaciens harbors the Ti plasmid that induces tumors on

Fig. 1 Agrobacterium-mediated transformation; wounded plant cells release certain phytochemicals, especially phenolics (P). These phenolics are sensed by VirA which activates VirG by phosphorylation. VirG further induces the expression of other vir genes. The gene products of the vir region facilitate the transfer of T-DNA with a gene of interest (GOI) to the plant cell. VirD1/D2 protect and export the T-DNA when it passes through the VirB transmembrane traversing apparatus. VirE2 and VirD2 together are responsible for the transport of T-DNA inside the plant cell. The T-DNA-VirE2-VirD2 complex enters into nucleus through the nuclear pore complex (NPC). The gene of interest with T-DNA left (LB) and right border (RB) is inserted into the plant chromosome via illegitimate recombination

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host plants’ crown region, whereas the A. rhizogenes contains the Ri plasmid that initiates hairy root formation. The plasmids Ti (pTi) and Ri (pRi) are the determining factors for the species specificity for A. tumefaciens and A. rhizogenes. Both the Ti and Ri plasmids have in their T-DNA regions, bacterial oncogenes (onc), structurally mimicking plant gene cassettes. These onc genes code for enzymes involved in the production of opines, a class of secondary amino acids conjugated to a sugar moiety and are responsible for hormonal alteration in the host cells resulting in callus formation. Opines are metabolized and used as the carbon and nitrogen source by the pathogen but not by the host cell. Based on the type of opines produced, Agrobacterium strains can be classified as nopaline, octopine, agropine, or succinamopine types [60]. Hooykaas and Beijersbergen reported that nopaline and octopine strains are the most prevalent. The A. tumefaciens C58, a nopaline-type strain, had been genome sequenced in 2001 [61]. The bacterial genome is bipartite and carries a circular and a linear chromosome. Along with, it also harbors two large plasmids, pTiC58 (pTi) and pAtC58 [62–64]. The pTi carries the T-DNA region. Upon A. tumefaciens infection, the T-DNA gets copied, which translocate and stably integrate into the plant nuclear genome through a complex process. The whole process involves the participation of multiple pTi-virulence (vir) region-derived vir proteins [65–67]. In order to biotechnologically exploit A. tumefaciens, the T-DNA has been modified to carry the gene of interest in place of the onc genes, without affecting the DNA delivery mechanism, a process dubbed disarming. This T-DNA molecule gets copied in the bacterial cell, which subsequently is transferred into the plant cell and gets integrated into the host plant genome. T-DNA carries a 25-bp imperfect direct repeats flanking both the ends called as border (RB and LB) sequences. The internal sequences within these border sequences can be tailored to carry the desired transgenes [68, 69]. Hoekema et al. developed the binary vector system, in which the pTi components were separated [38]. The vir genes are set to reside in a residential Ti plasmid, while the T-DNA region along with its border sequences is carried in a small handy plasmid that can replicate autonomously both in E. coli and Agrobacterium. The virulence genes VirA, VirB, VirC, VirD, VirE, and VirG, from the Ti plasmid, play a major role in T-DNA transfer, while Agrobacterium genome with the chromosomal virulence (chv) genes play a critical role in host recognition and attachment to the host cell surface [59]. Additionally, VirD5, VirE3, VirF, VirH, VirJ, VirK, VirL, VirM, VirP, and VirR were found as pTi-residing genes with roles in host-specificity [70]. Agrobacterium gets chemotactically attracted by the chemical exudates from the wounded plant tissue. These exudates, loaded with phenolics and with a slightly acidic pH (5.0–5.8), induce vir

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gene expression in A. tumefaciens. Acetosyringone (AS, 3,5-dimethoxy acetophenone) is the well-characterized compound [65, 71]. The low level expressing virA and virG genes get activated upon virulence induction. The transmembrane VirA, along with genome encoded ChvE detect external stimuli [72]. VirA gets autophosphorylated and transphosphorylates the transducer VirG; thus, VirA-VirG represents a two-component regulatory system [73]. The activated VirG binds to vir box region present in the promoter region of all vir genes [74]. VirH2, a P450-like enzymatic protein, is involved in detoxifying the host-generated toxic phenolics such as ferulic acid [75, 76]. Monosaccharides and derivatives such as galacturonic acid, glucuronic acid, arabinose, galactose, glucose, mannose, and cellobiose are also reported to enhance vir gene induction [74]. The chromosomal encoded ChvG protein, a part of ChvG-ChvI two-component regulatory system was proposed to play a role in external low pH-induced vir expression [77]. Upon vir induction, single-stranded T-DNA molecule (T-strand, the bottom strand of T-DNA region) is synthesized. VirD1 cooperates with the site-specific endonuclease VirD2 to cleave the T-strand between the third and the fourth nucleotide of both border sequences [78]. The virC operon participates in T-DNA processing by binding with the overdrive and not to the RB [79]. VirD2 covalently binds to the 50 end of the T-strand and recruits the single strand binding protein VirE2 to coat the T-strand to form T-complex [80]. The VirD2 and VirE2, both with nuclear localization signals (NLS), remain attached with the T-strand as T-complex, both in the bacterium and the host cell. VirE1 stabilizes VirE2 and prevents self-aggregation [81]. Concurrently, eleven VirB members and VirD4 forms the type IV secretion system (T4SS) with two major components: (a) VirB2, along with VirB5 and VirB7, forms the T-pilus for establishing bacterium–plant cell interaction, and (b) a membrane-associated transporter complex to translocate the T-complex [82, 83]. VirD4 acts as coupling receptor protein, the VirB1 performs a localized lysis in the bacterial peptidoglycan layer, and coordinates T-pilus formation, while VirB3, VirB4, VirB6, VirB8, and VirB11 act as inner membrane translocase, VirB7, VirB9, and VirB10 form the outer-membrane core complex, and VirB2 and VirB5 forms T-pilus [82, 84]. The covalently attached VirD2 acts as a pilot during T-complex translocation from bacterium to plant cell. Agrobacterium proteins VirD5, VirE2, VirE3, VirF, MobA, and Atu6154 are also reported to be translocated alongside VirD2 into the host cell [85–87]. VirE2 also interacts with host plasma membrane lipid molecules to form a transmembrane channel for the passage of T-complex [88]. VirD2 and VirE2 harbor NLS sequences and are reported to interact with several host proteins in the process of T-complex trafficking to the nucleus. VirD2 interacts with KAP

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(karyopherin) and three cyclophilin (ROC1, ROC4, and CypA) proteins. KAP was identified as nuclear import receptor in Arabidopsis, while the cyclophilins were proposed to play a role in proper conformation of VirD2 in host cell until T-DNA integration [89– 91]. VirE2, with NLS sequence, was shown no direct involvement in T-complex nuclear import, however, VirE2 interacts with VIP1 (VirE2-interacting factor), which in turn, interacts with KAP, forming a complex with VirE3 acting as an adaptor during nuclear import [92–95]. The VirF plays a role in the removal of VirE2 and VIP1 from the T-complex, while VirD2 remains intact with the T-strand [67, 96, 97]. The T-DNA integrates into plant chromosome by nonhomologous recombination, with microhomologies in the 30 region of T-strand with the host target loci sequences, often rich in A-T [98]. A nick in top strand of host genome’s integration locus leads the initiation of complementary strand synthesis for T-strand from 30 end to the VirD2 attached 50 end [99]. Both the T-strand and complementary T-strand anneals to form the doublestranded T-DNA. The generated T-DNA gets ligated into the targeted host locus, often with short deletions [98, 100].

3 Biolistics or Particle Bombardment-Mediated Genetic Transformation of the Nuclear Genome Biolistic transformation, also known as particle bombardment, is a physical means of forcing DNA molecule into the plant cells. It is the most preferred method of genetic transformation next only to the A. tumefaciens-mediated plant transformation. Unlike A. tumefaciens-mediated plant transformation, biolistic-mediated plant transformation does not depend on host genotype or receptivity. On the other hand, biolistic-mediated plant transformation often results in multiple and scrambled integrations [101]. The first successful particle bombardment system for plant cells was established by Klein et al., and was quickly followed in multiple models and other recalcitrant crop species such as wheat, rice, onion, and maize [57, 58, 102, 103]. The biolistic approach provides the opportunity to deliver large fragments of DNA such as bacterial artificial chromosomes [104]. Exogenous DNA integration occurs following both illegitimate and homologous recombination [101, 105]. Multiple comparative studies in plants such as maize, barley, and fescue, suggest particle bombardment-mediated transformation results in frequent complex and scrambled integrations in comparison to Agrobacterium-mediated transformation [106–108]. However, studies in sugarcane proved otherwise [109]. Particle bombardment-mediated transformation, however, is the most preferred method in experiments that demand rapid analysis and with transient expression such as promoter analysis,

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protein localization, transcription factor characterization, pathway elucidation, hormonal regulation of genes, and promoter component identification [110–118]. Particle bombardment allows for delivering the exogenous DNA not only to the nuclear genome but also to organellar genomes as well. For instance, chloroplasts in higher plants and chloroplast as well as mitochondria in algae were successfully transformed by the biolistic genetic transformation approach [119–122]. The biolistic machinery involves a gene gun apparatus with helium at high pressure that triggers the exogenous DNA molecule coated onto the gold/tungsten carriers (microprojectile) with high velocity into the recipient target tissue (Fig. 2). The microprojectile passes through the cell, while the coated DNA stays within the cell [101, 123]. Originally tungsten particles were used as the carrier; however, tungsten was replaced with gold particles as tungsten was observed to inhibit cell culture growth and damage DNA by inducing double-strand breakages [124, 125]. Torney et al. attempted to use silica particles with adsorbed DNA; however, these particles delivered DNA only in the presence of additional gold particles, suggesting that silica particles lacked the required momentum (please consult Chapter 8 in this volume for further discussion on this topic) [126]. Gold or tungsten microprojectiles when complexed with viral particles, E. coli, or yeast cells resulted in transient gene expression in the bombarded tobacco cells [127]. Particle bombardment is also used to deliver whole viral particle and viral

Fig. 2 Biolistic transformation; tungsten or gold microprojectiles are coated with a plasmid harboring the gene of interest and bombarded onto the plant tissue. This technique is used for nuclear as well as chloroplast transformation

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fragments that code for viral proteins to dissect the role of the candidate sequence [[128, 129]. Interestingly, de Mesa et al. used the gene gun to launch gold particles coated with A. tumefaciens to transform strawberry and observed increased transformation efficiency than the routine A. tumefaciens-mediated transformation [130]. Further, Chen et al. demonstrated RNA delivery by bombardment and studied gene silencing in rice [131].

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Biolistics-Mediated Chloroplast Transformation Plant cells carry chloroplasts and mitochondria, while animal and fungal cells carry only mitochondria. These organelles are semiautonomous with their genetic material and machinery to carry out replication, transcription, and translation, and resemble molecularly the prokaryotic organisms [132]. Agrobacterium has evolved to deliver and integrate its T-DNA into the plant nuclear genome. However, physical means of DNA delivery paved the way for delivery of exogenous DNA into the organellar genome. Boynton et al. reported the first successful chloroplast transformation in Chlamydomonas reinhardtii [133]. However, Nicotiana tabacum was the first higher plant species to be chloroplast transformed by biolisticmediated DNA delivery approach [122]. Biolistics is the preferred approach for organellar transformation, while occasionally protoplasts were subjected to polyethylene glycol (PEG) treatment to achieve transient as well as stable chloroplast transformations [134, 135]. Several plant species were optimized and reported for successful chloroplast transformation [122, 136, 137]. Unlike chloroplast, the mitochondrial transformation was achieved only in two single cellular organisms, Saccharomyces cerevisiae and C. reinhardtii [138–140]. Plant, as well as animal mitochondrial transformation, has not been successful yet [141]. However, an in organello genetic modification was demonstrated by Mileshina et al. [142]. Chloroplast transformation has several advantages over nuclear transformation such as high gene expression, targeted gene integration, containment of transgenes, uniparental passage, and absence of ploidy and position effect. The high copy number of chloroplasts per cell allowed for harvesting the transgene product up to 70% of total soluble protein and hence preferred for plantbased antigen production [143, 144]. The exogenous DNA integrates into the chloroplast genome exclusively by homologous recombination [143]. As chloroplasts resemble prokaryotic cell at the molecular organizational level, multiple genes can be inserted and expressed as polycistronic operons [145, 146].

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Bioactive Beads-Mediated Gene Transfer This method involves immobilization of DNA fragments onto calcium alginate beads. In this procedure, the DNA containing beads are incubated with protoplasts, followed by routine washing, selection, and regeneration. Calcium alginate beads are positively charged and can electrostatically interact with the negatively charged DNA molecules and the cell membrane (Fig. 3). Drop mixing of DNA containing calcium chloride solution into emulsified sodium alginate solution often followed by mixing or sonication leads to the formation of calcium alginate beads with DNA fragments [147, 148]. DNA fragments up to the size of 280 kb can be immobilized [149]. Sone et al. reported the first application of calcium alginate beads in plant genetic transformation using tobacco protoplast [147]. Later, the transformation of several other model plant species was achieved [150–152]. Wada et al. reported that bioactive beads-mediated transformation resulted in transgenics with relatively low transgene copy number [148]. However, occasional multiple insertions and transgene rearrangements were also observed while using large DNA fragments up to 100 kb [153]. The usage of DNA–Lipofectin® complex was shown to increase the transformation efficiency fourfold with naked DNA [148]. Wada et al. also observed similar transformation efficiency by using uniform smaller (~3 μm) beads [148].

Fig. 3 Pollen tube pathway; this method exploits pollen tube for the delivery of exogenous DNA containing a transgene into the developing embryo sacs

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Pollen and Pollen Tube Pathway-Mediated Plant Transformation The long-term in vitro maintenance of plant tissues often induces somaclonal variations [154]. Pollen tube-mediated transformation (PTT) is a special method of plant transformation as it differs in steps in obtaining transgenic plant without routine tissue culture steps. The PTT resembles the floral dip method; however, does not involve Agrobacterium (Fig. 4) [155]. Hess and Ohta claimed the usage of pollens to develop transgenic plant solely based on the phenotypic data [156, 157]. However, their results were not reproducible [158, 159]. The likely explanation of these contradictory results was the degradation of exogenous DNA by pollen nucleases. Subsequently, Zhou et al. reported the transformation of cotton by PTT approach [160]. These authors performed PTT by removing the stigma from the postanthesis flowers and application of the DNA fragments of interest to the cut surface [160–162]. The exogenous DNA was proposed to pass through the pollen tube and integrate into the genome of an undivided nascent diploid zygote. PTT efficiency can be greatly influenced by various factors such as the candidate plant species, flower size, the timing of the postanthesis DNA delivery, and concentration of exogenous DNA

Fig. 4 Transformation techniques using plant protoplasts. (a) Electroporation; electric pulse creates pores in the cell membrane of plant protoplast through which the exogenous DNA enters the plant cell and expresses. (b) Microinjection; the exogenous DNA is delivered to the nucleus of the plant cell through an injection needle. (c) Bioactive bead-mediated gene transfer; large DNA molecules can be efficiently transferred by embedding them into a bead. (d) Silicon carbide whisker-mediated transformation; small needle-like silicon carbide whiskers with high tensile strength and gene of interest are mixed with plant protoplasts. The mixture is vortexed and protoplasts are plated on selection plates for the assessment of DNA insertion

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in the solution. Delayed delivery of DNA solution may lead to the formation of chimeras as the cell division would start before the DNA delivery [163]. Several model and crop plants have been reported to be optimized for PTT [164, 165]. Liu et al. modified the PTT (ovary drip method) by cut-opening ovary and applying exogenous DNA directly onto the ovules and reported increased efficiency of transformation. Apart from PTT, pollen itself can be transformed prior to pollen fertilization and can be allowed to fertilize the egg to generate the transgenic seed [164]. Matthews et al. demonstrated the electroporation of pollen and transient expression of foreign genes [166]. The first transgenic plant using electroporated pollen was reported by Smith et al. [167]. Particle bombardment-mediated pollen transformation failed to develop transgenic plants, and the expression reported was only transient [168, 169]. Neuhaus et al. reported the development of transgenic embryos using pollen microinjected with exogenous DNA; however, the embryos were chimeric in nature [170].

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Silicon Carbide Whisker- and Nanoparticle-Mediated Plant Transformation Kaeppler et al. first demonstrated silicon carbide (SiC) whiskermediated plant transformation (SCWPT) [171]. This procedure involves the mixing of cell suspension cultures with silicon carbide fibers (whiskers), which physically penetrate the cell, causing cell perforation and abrasion, thereby allowing for the entry of exogenous DNA (Fig. 3). Several model and crop species have been successfully transformed by SCWPT method [172]. Wheat, with low transformation efficiency for Agrobacterium, showed increased transformation efficiency upon SiC treatment [173]. A similar observation was reported earlier by Nagatani et al. in rice [174]. The SCWPT requires a sophisticated protocol for plant regeneration from cell cultures and thus is preferred least for plant transformation. Several nanoparticulated molecules are studied and applied to deliver molecules into the cell. Torney et al. reported the delivery of DNA into the cell by silica nanoparticles [126]. Zhao et al. transformed pollens using magnetic particles and forwarded to the next generation to generate transgenic plants [175]. Using carbon nanotubes, Demirer et al. delivered the exogenous DNA into the plant cell and reported transient expression [176]. The mechanism of nanoparticle-mediated genetic transformation is, however, largely unclear and proposed that the nanoparticles protect the DNA from cellular enzymatic attack until delivery [177, 178]. Studies on nanoparticle-mediated plant transformation are emerging and yet to be expanded (for further discussion on this subject consult Chapter 8 in this volume).

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Electrophoretic Transfection and Electroporation-Mediated Plant Transformation The electroporation-mediated plant transformation (EPT) is a similar technique as used in the bacterial transformation. Shimamoto et al. reported the first successful transformation of rice protoplasts and transgenic plants recovery by EPT [179]. Later, a similar approach was performed in barley [180]. Similar to other techniques, EPT needs established protocol to regenerate plants from protoplasts. However, this method can be applied to study transient analysis and to study gene functions at the cellular level. Ahokas used barley kernels to transfect electrophoretically; however, the expression was transient (Fig. 3) [181]. Later, Chowrira et al. made an approach similar to floral dip [182]. The nodal meristem was electroporated with exogenous DNA, and allowed to develop flowers and offspring with the transgene. Using maize and potato mitochondria, in organelle electrophoretic transfection was also performed, and gene integration characterized [142].

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Microinjection-Mediated Plant Transformation Similar to particle bombardment, microinjection-mediated plant transformation is also a method of direct DNA delivery and does not rely on host plant species as in Agrobacterium-mediated transformation. However, unlike particle bombardment, in microinjection, a single cell is transformed and with lower efficient recovery. The exogenous DNA is injected into the cytoplasm or nucleus of the isolated recipient cell that is immobilized onto low-meltingpoint agar. Microinjection was originally used for genetic modification of animal cell lines (Fig. 3). The first study in plants was reported in tobacco mesophyll cells [183]. Following this, Neuhaus et al. developed haploid rapeseed plants obtained from microinjection-mediated transformed microspores [170]. Similarly, Jones-Villeneuve et al. attempted microinjection in uninuclear microspores of rapeseed; however, the regenerated putative transgenic plants did not carry a stable integration [184]. Holm et al. used zygotic protoplasts from barley and demonstrated a successful transformation of barley by microinjection [185]. Baskaran et al. used shoot apical meristem of cucumber and microinjected it with A. tumefaciens containing the binary vector and demonstrated regeneration of plants from the microinjected shoot apical meristems [186]. The experiment resulted in obtaining more transgenic plants than routine A. tumefaciens-mediated transformation. In consideration of the facts, such as poor transformation efficiency, the transformation of the small number of cells, the labor involved, and availability of relatively more established methods, microinjection-mediated plant transformation method is often

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not preferred by plant biologists in their routine research, unless specifically needed. A comparison of different plant genetic transformation methods reported in this chapter is presented in Table 1.

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Progress in Recalcitrant Plant Transformation Plant transformation depends on genetic engineering of plant cells followed by regeneration of plants from those transformed cells [187]. The plant species that are optimized for plant transformation and regeneration are ever expanding for the past 35 years. However, several plant species remain recalcitrant for plant transformation and in particular for regeneration processes. One possible way is developing alternative methods that are devoid of regeneration steps involving tissue culture such as floral dip method reported by Clough and Bent in Arabidopsis thaliana, Verma et al. in Brassica napus, and Fang et al. in an important cut flower, Lisianthus (Eustoma grandiflorum) [155, 188, 189]. Additionally, a vacuum infiltration-assisted floral dip method was shown to be successful in Camelina [190]. However, not all species are as receptive as A. thaliana; therefore, the tissue culture-mediated plant transformation method remains to be the only choice in such cases and needs to be optimized. Genotypic variation in crop plants also throws a great challenge in plant tissue culture. Plant regeneration efficiency can be increased by exposure to various stresses and manipulating hormonal concentrations such as auxins and cytokinins [191–193]. Modulating the expression of genes coding for transcription factors such as WUS and BABY BOOM can be attempted to increase transformation efficiency [194– 196]. Further, plants’ susceptibility to Agrobacterium can be increased by downregulating infection-responsive genes such as MYB transcription factors [197]. Tissue browning, which reduces transformation efficiency, can be averted by supplementing medium with selected antioxidant compounds such as α-lipoic acid [198]. Comparatively, monocots are more recalcitrant than dicots, and this could possibly be because of the difference in wound response [199]. Several modifications have been performed throughout history and currently a wide range of monocot crop species are available with optimized transformation methods [200, 201]. Increasing the virulence of Agrobacterium strains with additional copies of vir genes may also be tried to get efficient transformation. Komari developed “super-binary vector” that carried a segment of vir region from A281 strain [202]. This vector was used by several groups to develop transgenic monocots such as maize, wheat, barley, and sorghum [203–206]. Using tomato as a system, Someya et al. demonstrated that reducing ethylene production can increase the transformation efficiency [207]. Further, using embryogenic calli dramatically increased the transformation

With and without TCR

With and without TCR

With and without TCR

Requires plant regeneration from protoplast

Requires plant regeneration from protoplast

With TCR

With and without TCR

Agrobacterium

Biolistic

Electroporation

Microinjection

Lipofection

Silicon carbide fibers

Virus based methods

Tissue culture and Transformation regeneration (TCR) method requirement

The most preferred method for stable gene integration and nuclear transformation

Limited to virus host specificity

Unrestricted

Cells, tissues and whole plant

Wide range of cell types

High; transient

Moderate

Quick, affordable, and easy to set up

Quick, affordable, and easy to set up

High in PEG Requires optimization with regenerable based method suspension cell culture

Protoplasts

Restricted to species amenable to protoplasting

High skill and experience needed, whole chromosome can be potentially transformed

Very low

Protoplasts

Requires tedious optimization process

Low/Moderate; Desired technique for chloroplast stable transformation and transient expression assays. Risk of high-copy integration and gene rearrangement

High; stable

Transformation Efficiency Notes

Protoplasts, pollen Low grains, and meristems

Any intact tissue, explants or microspores

Cells, tissues, and whole plant

Tissue type

Restricted to species amenable to protoplasting

Unrestricted

No issues with plant species range

Wide range of dicots and few monocots

Plant species range

Table 1 A comparative illustration of plant genetic transformation methods and their features

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efficiency, especially in monocots such as rice, maize, and sugarcane [42, 208, 209]. However, shoot apical meristem stood good choice for pearl millet [210]. Hiei et al. reported that embryo pretreatment with heat and centrifugation can increase the Agrobacteriummediated transformation efficiency in rice [211]. Cocultivation period and temperature greatly influenced the transformation efficiency in Chrysanthemum [212].

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Gene Integration Through Homologous Recombination Homologous recombination was originally utilized for the removal of marker genes from transformants; later it was primarily used for developing transgenics with multigene stacking, site-specific integration, or site-specific mutagenesis [213, 214]. In these transgenics, the stable integrations are often flanked by the AT-rich DNA sequences [215, 216]. Random integration often resulted in variable transgene expression and even positional effect silencing, often influenced by integration site, termed as “chromosomal position effect” [217, 218]. Multiple T-DNA copies are often observed to integrate as inverted T-DNA repeats, potentially triggering transgene silencing [219]. Transgene integrations in actively transcribing genic regions and strong promoters of transgenes at the vicinity of transcriptionally repressed endogenous genes often results in unpredicted phenotypes [220, 221]. Multigene insertions in multiple loci pose an enormous challenge for the introgression of those events to elite varieties by conventional crop breeding. Sequential retransformation, a time-consuming practice, demands diverse selectable markers, and well-established transformation protocols for concerned crop species [222]. Site-specific DNA breakage can be achieved by designing nucleases for transgene integration directed at specific sites. Molecular mapping and marker-assisted breeding of crop species can be more precise and targeted with the use of nextgeneration sequencing technology and rapidly accumulating omics data advanced bioinformatics tools and use of artificial intelligence [223–225]. Cre, a recombinase from E. coli bacteriophage P1, was used to eliminate marker genes that carried lox recombination sites (cre–lox recombination), later the technology was used for gene stacking in transgene locus with pre-integrated lox sites [226]. However, this technology depends on the first random integration event to be targeted [227]. Cre–lox system was soon replaced by the modern nucleases-mediated technology such as Zinc-finger nucleases (ZFN), Transcription activator-like effector nucleases (TALEN) and Clustered regularly interspaced short palindromic repeats/CRISPR associated protein 9 (CRISPR/ Cas9), which virtually allow for precise integration at any chromosomal loci.

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ZFN and TALEN are custom designed nucleases that carry bacterial FokI endonuclease cleavage domain to cleave specific site. ZFN and TALEN both contain two prominent domains, tailored DNA-binding domain and DNA-cleavage bacterial FokI domain. This DNA-binding domain binds to the specific candidate DNA sequence, while the DNA-cleavage domain cleaves the DNA to generate double-strand break (DSB). This process recruits endogenous DNA repair machinery to perform nonhomologous end-joining (NHEJ) or homologous recombination (HR) [228]. Nucleases-mediated technology allowed researchers to mutate and manipulate the target loci to get the desired phenotype. Lloyd et al. and Wright et al. were the first to employ ZFN to develop transgenic Arabidopsis and tobacco, respectively, with sitespecific mutations [229, 230]. The DNA-binding domain of ZFN carries three to six ZF repeats, which respectively recognize 9–18 specific base pairs. The DNA-binding sequences can be tailored by modifying the ZF repeats [231]. However, ZFNs are also shown to fail in achieving the expected outcome [232]. TAL effectors are secreted by Xanthomonas type III secretion system during the plant infection process. The DNA-binding domain carries a conserved 33–34 amino acid (aa) sequence with variable 12th and 13th amino acid dubbed Repeat Variable Di-residues (RVDs), that recognizes the cleaving site. This RVD can be altered to make custom designed TALEN [233]. TALENs exhibit relatively higher specificity than ZFN technology [234]. The use of the CRISPR/Cas9 system in plant genetic engineering is relatively more recent and widely accepted tool for genome editing than ZFNs and TALENs [235].

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CRISPR/Cas9-Mediated Genome Editing and Beyond The CRISPR-Cas system has evolved in prokaryotic bacteria and archaea against invading alien genetic elements as an RNA-guided adaptive immune system to evade entities like viruses, transposons, and plasmids. The host genome carries DNA repeat sequences, dubbed the CRISPR (clustered regularly interspaced short palindromic repeats) locus which has incorporated small DNA fragments of previous invading elements, and accompanying CRISPR-associated (Cas) genes [236, 237]. Upon entry of alien elements, the selected region of the fragmentized elements (spacers) are incorporated into the CRISPR locus and flanked on either side by direct repeats; thus, the CRISPR loci act as a genetic ledger of preinfected elements, a step called immunization. Upon the second incident of infection by pre-infected element, CRISPR RNAs (crRNAs) are generated from the CRISPR array, with spacer at the 50 end, assembled with Cas proteins, forming crRNA–effector complexes. Hybridization of spacer region of the crRNAs with incoming DNA/RNA elements leads to cleavage of the candidate

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element, in a sequence-specific manner. The target recognition often requires the presence of 2- to 5-bp conserved Protospacer Adjacent Motif or PAM [238]. CRISPR/Cas systems are grouped into six (I–VI) types [239, 240]. The type II CRISPR/Cas9 system from Streptococcus pyogenes, which utilizes single Cas (Cas9) protein in place of multiple Cas members is employed widely in modern research applications. Further type II CRISPR/Cas system requires a small noncoding RNA, called the trans-activating CRISPR RNA (tracrRNA) to hybridize to the CRISPR RNA (crRNA), and form a double-stranded RNA dubbed guide RNA (gRNA) for the target recognition and cleavage [237, 238]. The crRNA and tracrRNA can be combined to form single-guide RNA (sgRNA) [241]. Tailoring the sgRNA and delivering into host cell can lead to targeted DNA cleavage with blunt-ended double-strand break (DSB) in the host genome, which then follows a DNA repair mechanism either by NHEJ with small random insertions and/or deletions (indels) or by homology-directed DNA repair (HDR) [238, 242], thus resulting in editing the genome at the targeted locus. The CRISPR/Cas9 from the prokaryotic system was tested successfully in eukaryotic human and mouse mammalian systems simultaneously by two groups [243, 244]. In quick succession, several groups demonstrated CRISPR/Cas9-mediated genome editing in model plant systems (Arabidopsis, rice, Nicotiana tabacum, and N. benthamiana) for developing transgenic plants, calli, protoplast, and in planta using various genetic transformation technologies [245–249]. Later, model plants, N. benthamiana, N. tabacum, N. attenuate, and Marchantia polymorpha, crop plants such as Oryza sativa, Triticum aestivum, Sorghum bicolor, Zea mays, Solanum lycopersicum, S. tuberosum, Brassica oleracea, Glycine max, Hordeum vulgare, Lactuca sativa, Cucumis sativus, and tree species such as Citrus sinensis, Populus tomentosa, P. tremula, and Medicago truncatula were reported to be genome edited with CRISPR/Cas9 technology for trait improvement or functional genomic studies [242, 248–265]. Recently, metabolic engineering by editing multiple genes using CRISPR/Cas9 was achieved in plants such as Papaver somniferum and tomato [266, 267]. CRISPR-Cas9 system has also been employed to generate plants for resistance against plant viral pathogens [268]. CRISPR interference (CRISPRi) is a further technological modification using CRISPR, through which mutations in promoter regions that block transcription initiation and elongation, and precise gene silencing can be achieved [269, 270]. Recently, this technology has been successfully implemented in plants for precise nucleotide base editing [271]. Similarly, gene targeting by homology-directed repair in plants has proven to be successful using the CRISPR/Cas9 system [187].

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Conclusions The fundamental understanding of plant molecular biology has advanced remarkably through the current progress in the functional genomics. However, translating this basic knowledge to crop improvement is significantly constrained by the inefficient genetic transformation methods. Although in planta transformation method is a smart solution, which eliminates the need for tissue culture and regeneration, so far it is imitated/restricted to a few plant species. Therefore, future research is needed to develop novel tissue culture independent universal methods of plant genetic transformation, which are cost-effective, technically less demanding, and unrestricted so that these could be implemented in the laboratories in the resource-deprived countries. Without the innovative and smart solutions to overcome the bottleneck of plant genetic transformation, the implementation of recent advances in plant genome editing and synthetic biology will stay a distant dream. Usage of plant genes involved in developmental reprogramming and usage of tissue culture-independent nanoparticle-based genetic transformation systems are few such initiatives in this direction.

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using tomato as a model. Plant Physiol 166:455–469 256. Wang S, Zhang S, Wang W, Xiong X, Meng F, Cui X (2015) Efficient targeted mutagenesis in potato by the CRISPR/Cas9 system. Plant Cell Rep 34:1473–1476 257. Lawrenson T, Shorinola O, Stacey N, Li C, Østergaard L, Patron N, Uauy C, Harwood W (2015) Induction of targeted, heritable mutations in barley and Brassica oleracea using RNA-guided Cas9 nuclease. Genome Biol 16:258 258. Jacobs TB, LaFayette PR, Schmitz RJ, Parrott WA (2015) Targeted genome modifications in soybean with CRISPR/Cas9. BMC Biotechnol 15:16 259. Chandrasekaran J, Brumin M, Wolf D, Leibman D, Klap C, Pearlsman M, Sherman A, Arazi T, Gal-On A (2016) Development of broad virus resistance in non-transgenic cucumber using CRISPR/ Cas9 technology. Mol Plant Pathol 17:1140–1153 260. Jia H, Wang N (2014) Targeted genome editing of sweet orange using Cas9/sgRNA. PLoS One 9:e93806 261. Fan D, Liu T, Li C, Jiao B, Li S, Hou Y, Luo K (2015) Efficient CRISPR/Cas9-mediated targeted mutagenesis in Populus in the first generation. Sci Rep 5:12217 262. Michno J-M, Wang X, Liu J, Curtin SJ, Kono TJ, Stupar RM (2015) CRISPR/Cas mutagenesis of soybean and Medicago truncatula using a new web-tool and a modified Cas9 enzyme. GM Crops Food 6:243–252 263. Jaganathan D, Ramasamy K, Sellamuthu G, Jayabalan S, Venkataraman G (2018) CRISPR for crop improvement: An update review. Front Plant Sci 9:985

264. Jang G, Joung YH (2019) CRISPR/Casmediated genome editing for crop improvement: current applications and future prospects. Plant Biotechnol Rep 13:1–10 265. Zhou X, Jacobs TB, Xue LJ, Harding SA, Tsai CJ (2015) Exploiting SNPs for biallelic CRISPR mutations in the outcrossing woody perennial populus reveals 4-coumarate: CoA ligase specificity and redundancy. New Phytol 208:298–301 266. Alagoz Y, Gurkok T, Zhang B, Unver T (2016) Manipulating the biosynthesis of bioactive compound alkaloids for nextgeneration metabolic engineering in opium poppy using CRISPR-Cas 9 genome editing technology. Sci Rep 6:30910 267. Li R, Li R, Li X, Fu D, Zhu B, Tian H, Luo Y, Zhu H (2018) Multiplexed CRISPR/Cas9mediated metabolic engineering of γ-aminobutyric acid levels in Solanum lycopersicum. Plant Biotechnol J 16:415–427 268. Ali Z, Abulfaraj A, Idris A, Ali S, Tashkandi M, Mahfouz MM (2015) CRISPR/Cas9-mediated viral interference in plants. Genome Biol 16:238 269. Gilbert LA, Larson MH, Morsut L, Liu Z, Brar GA, Torres SE, Stern-Ginossar N, Brandman O, Whitehead EH, Doudna JA, Lim WA, Weissman JS, Qi LS (2013) CRISPR-mediated modular RNA-guided regulation of transcription in eukaryotes. Cell 154:442–451 270. Qi LS, Larson MH, Gilbert LA, Doudna JA, Weissman JS, Arkin AP, Lim WA (2013) Repurposing CRISPR as an RNA-guided platform for sequence-specific control of gene expression. Cell 152:1173–1183 271. Marzec M, Hensel G (2018) Targeted base editing systems are available for plants. Trends Plant Sci 23:955–957

Chapter 4 Choice of the Promoter for Tissue and Developmental Stage-Specific Gene Expression Olga G. Smirnova and Alex V. Kochetov Abstract Transgenic technologies belong to important tools of reverse genetics and biotechnology in plants. Targeted genetic modifications can reveal functions of genes of interest, change metabolic and regulatory pathways, or result in accumulation of valuable proteins or metabolites. However, to be efficient in targeted genetic modification, the chimeric gene construct should be designed properly. In particular, the promoters used to control transgene expression need to be carefully chosen. Most promoters in widely used vectors belong to strong and constitutively expressed variants. However, in many cases transgene expression has to be restricted to certain tissue, stage of development, or response to some internal or external stimuli. In turn, a large variety of tissue-specific promoters have been studied and information on their characteristics may be recovered from the literature. An appropriate promoter may be selected and used in genetic construct to optimize the transgene transcription pattern. We have previously designed the TGP database (TransGene Promoters, http://wwwmgs.bionet.nsc.ru/mgs/dbases/tgp/home.html) collecting information from the publications in this field. Here we review the wide range of noncanonical tissue-specific and developmentally regulated promoters that might be used for transgene expression control. Key words Promoter, Tissue-specific, Developmentally regulated, Transgenes, Genetic engineering, Expression control

1

Introduction Plant research using transgenic approach frequently demands selection of promoters that provide appropriate transcription activities. The set of promoters commonly used in chimeric gene construction is rather limited, and provides only a restricted variation of gene expression patterns. Moreover, the usage of two identical promoters in a complex construct can induce transgene silencing. Although this problem can be solved by using a variety of plant gene promoters with experimentally verified characteristics, it requires a time-consuming analysis of literature data. We have recently developed the TransGene Promoters Database (TGP; http://wwwmgs.bionet.nsc.ru/mgs/dbases/tgp/ home.html) to annotate plant promoters with known

Sachin Rustgi and Hong Luo (eds.), Biolistic DNA Delivery in Plants: Methods and Protocols, Methods in Molecular Biology, vol. 2124, https://doi.org/10.1007/978-1-0716-0356-7_4, © Springer Science+Business Media, LLC, part of Springer Nature 2020

69

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Olga G. Smirnova and Alex V. Kochetov

characteristics [1, 2]. TGP collects data on the promoter nucleotide sequences and their specific activities in transgenic plants. Although it is still far from being complete, the database has been periodically updated and the list of annotated plant promoters is constantly expanding. TGP could be useful for selection of candidate promoters with characteristics appropriate for particular experimental needs. In this chapter, we present a brief overview of available literature data concerning a wide range of tissue-specific and developmentally regulated plant promoters, whose activities have been experimentally evaluated with transgenic plants. Frequently, the promoter expression pattern is determined by the evaluation of β-glucuronidase (GUS) or green fluorescent protein (GFP) reporter gene expression. For each promoter, we give the size, position, level of expressions, and plant tissues in which the promoter has been tested. Some of them that have already been annotated in the TGP database are referenced by the corresponding identifiers, which allow to get the information about the promoter activity and the promoter sequence from the TGP database. We do not include in the chapter a number of widely known promoters whose information is generally available and has previously been presented in a number of reviews [3–6]. Rather, we have included information on less-known tissue-specific and developmentally regulated promoters isolated from exotic species in some cases, which may have the potential for use in transgenic research. Some of these promoters have conservative regulatory elements and can be used to control the expression of target genes in heterologous species.

2

Materials

2.1 Fruit-Specific Promoters

Many fruit-specific promoters have been characterized and shown to direct ripening-specific expression of reporter genes (Table 1). Tomatoes are often used for determination of a fruit-specific promoter activity from species with long life cycle or that are difficult for transformation. The tomato fruit has several stages of ripeness: green, breaker, turning, pink, and red [7]. At a breaker stage, a color starts to be visible from exterior on blossom end of the fruit. At a turning stage, fruits are 10–30% red. Polygalacturonase is one of the enzymes involved in fruit ripening. A promoter of a polygalacturonase (PGA) gene from kiwifruit (Actinidia chinensis) was studied in tomato (Lycopersicon esculentum cv. UC82B). CkPGA promoter fragments (
467 to +189,
860 to +189, and
1296 to +189 from the translation initiation site, TIS) directed fruit ripening-specific GUS gene expression in transgenic tomatoes [8]. No histochemical staining was detected in leaves, roots, flowers, abscission zones, or

1489 bp

1359 bp

0.7 kb

1111 bp

2919 bp 403 bp

3025 bp

3159 bp

no

1148 bp

1800 bp, 872 bp

2065 bp, 1258 bp

1032 bp, 600 bp

4.2 kb

CkPGA

AcAct

MaACO1

MaACS

PpACO1

FaEG1

FaEG3

FaAPX

FaGalUR

MdPG

MdACO

EgDes

CuMT45

CvAGPL1 1573 bp, 1301 bp

Size

Promoter

Tomato

Tomato

Tomato

Strawberry

Strawberry

Strawberry

Strawberry

Tomato Tomato

Banana

Banana

Petunia

Tomato

Tested in

Watermelon

Tomato, watermelon

Orange

Satsuma mandarin Arabidopsis

Oil palm

Apple

Apple

Strawberry

Strawberry

Strawberry

Strawberry

Peach

Banana

Banana

Kiwifruit

Kiwifruit

Origin

Table 1 Promoters for fruit- and flower-specific expression

Mid

Mid

Strong

Strong

Strong

Strong

Strong Mid

Green and ripe fruits

Fruit (juice sac)

Silique

Strong

Mid

Mature green fruit, ripe Strong fruit

Ripening fruit

Ripening fruit

Ripe fruit

Turning fruit

Fruit

Ripening fruit

Ripening fruit Constitutive in fruit

Ripening fruit

Ripening fruit

Receptacles, placenta of Strong mature seed pod

Flower, seed, rosette leaves Leaf

Leaf

Leaf

Leaf, stem, flower, root

Vascular system of leaves, petals, stems, and roots

Activity Other tissues

Ripening fruit, breaker Mid stage

Main organ/tissue

Brazzein

FaAGPS(
), FaCel1(
)

Target gene

(continued)

Eg:Des

Md:ACO

Md:PG

Fa:GalUR

Pp:ACO1

TGP ID

Tissue-Specific and Developmentally-Regulated Plant Promoters 71

2305 bp

1.5 kb

601 bp

1.2 kb

1038 bp

CaCCS

MdANS1

PpACO1

AtFUL

LePDS

733 bp

908 bp

ShCYC-B

Tomato

Arabidopsis

Peach

Apple

Pepper

Solanum habrochaites

Arabidopsis

3873 bp 682 bp

Cucumber

AtFUL

2.1 kb

CsLS

Cucumber

Melon

1.5 kb

CsExp

Origin

Cucumisin 1181 bp, 234 bp

Size

Promoter

Table 1 (continued)

Green and ripe fruit

Valve, silique

Immature green fruit

Fruit

Ripe fruit

Main organ/tissue

Tobacco

Tomato

Arabidopsis

Tomato

Tobacco

Mid

Mid

Strong

Mid

Mid

Strong

Petal, anther, corolla, red fruit Petal, ovary

Inflorescence, floral meristem

Flower

Strong

Strong

Mid

Mid

Petal

Ovary

Stem, root, fruit

Vegetative organs

Leaf

Stamen, leaf

Inflorescence

Leaf, stem

Activity Other tissues

Strong Flower, sepal, petal, receptacle, immature seed

Pepper, tomato Green and ripe fruit

Tomato

Arabidopsis, false flax

Melon

Cucumber

Cucumber

Tested in

CrtO

ClLS

Target gene

Le:PDS

Pp:ACO1

Ca:CCS

TGP ID

72 Olga G. Smirnova and Alex V. Kochetov

Tissue-Specific and Developmentally-Regulated Plant Promoters

73

developing fruit prior to breaker stage. In tomato fruit at breaker stage staining was observed throughout the inner and outer pericarp as well as in the columella and seeds. The strongest staining was observed in the vascular strands. At later stages in ripening, staining was typically restricted to the inner pericarp and seeds. At these stages, the
1296 CkPGA promoter fragment provided weaker GUS staining than either the
860 or
467 fusions. Actinidin is the most abundant soluble protein in kiwifruit. The actinidin mRNA is rare in leaves and roots and increases in abundance during later stages of fruit development. The 50 -flanking region (
1301 to +58 from the transcription start site, TSS) of a kiwifruit actinidin gene (AcAct) directed high-level gene expression in receptacles and placenta of mature seed pods of transgenic petunia plants cv. Mitchell [9]. Significant GUS activity was induced in the placenta of seed pods approximately 2 weeks after pollination, and GUS staining reached a high level 4 weeks after pollination, when the seed pods were starting to turn brown. Placental tissues rapidly underwent senescence thereafter and dried out. Some GUS expression was also detected in the vascular system in leaves, petals and stems. In the root system GUS activity is preferentially detected at the root branching points. A shorter AcAct promoter fragment consisting of nucleotides
115 to +58 conferred similar spatial and temporal regulation in some of the transgenic petunia plants. Kiwifruit actinidin promoter is able to direct expression of a heterologous gene during the late stages of seed pod development in petunias. Detailed study of AcAct promoter activity during “ripening” was not possible with petunia because the tissues dehydrate rapidly as they change color and senesce. GUS expression in transgenic tomato plants was qualitatively similar to the patterns found in petunia. Suppression of GUS expression observed in some T1 tomato plants and their progeny prevented to obtain accurate quantitative data for GUS expression during tomato fruit ripening [9]. ACC oxidase (ACO1) and ACC synthase (ACS) genes specifically expressed in banana (Musa acuminate AAA Group) ripening fruits. The results of transient gene expression assay showed that the 0.7 kb of the MaACO1 promoter was able to direct fruitspecific gene expression in banana [10]. Some positive regulatory elements may exist within the region from
822 to
468. For the MaACS promoter the regulatory region for fruit specificity was possibly located in the region from
1111 to +1 and a positive regulatory region may locate between nucleotide
1111 to
608 [11]. A PpACO1 gene encodes 1-aminocyclopropane-1-carboxylic acid oxidase from peach (Prunus persica cv. Loring). The fulllength PpACO1 promoter (2919 bp upstream of TIS) directed GUS expression in the green stage of tomato (cv. Pixie) fruit development, and increased GUS expression as fruit matured

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Olga G. Smirnova and Alex V. Kochetov

([12]; TGP Pp:ACO1). A regulatory region between
2919 and
2141 controls the temporal expression of the promoter in tomato fruits. GUS staining in fruit first appeared in vascular bundles regardless of developmental stage. In the low expression lines, staining was confined to the vascular bundles and the columella, whereas in the high expressers, staining was extended to the placental tissue and the pericarp. The locular tissue was never stained as much as the rest of the fruit tissues and no staining was visible in the epidermis. The full-length promoter provided significant staining detected in leaf, flower, stem, and root, but at apparently lower levels than in the fruit tissue. The
403 bp PpACO1 promoter had significant constitutive activity at all stages of tomato fruit development (from green to red) with little or no expression in other tissues. The
403 bp PpACO1 promoter activity in tomato fruit is comparable to that of the most popular constitutive 35S promoter obtained from the Cauliflower Mosaic Virus (CaMV35S) [12]. “Pp:ACO1” is the identifier of the PpACO1 promoter in the TGP database. Two endo-b-1,4-glucanases, also known as cellulases, are involved in softening of fruits and highly expressed during ripening of strawberries (Fragaria  ananassa) [13]. Full-length promoters of both genes (FaEG1 and FaEG3) were highly expressed in strawberry fruits in transient assays. The strength of the largest 50 fragments of the promoters (
2980 to +45 from TSS for FaEG1 and
3025 to +134 from TSS for FaEG3) was greater than that of the CaMV35S promoter. In the case of FaEG3, the 30 region had a downregulating effect on the expression of GUS, and this might account for the lower amount of FaEG3 mRNA usually observed in ripe fruits compared to that of FaEG1. The fruit ripening specificity and the high rate of expression of FaEG1 make its promoter a good candidate for biotechnological uses that require a fruit and ripening specific expression of a target gene in strawberry. Early expression of FaEG3 in fruits might offer a tool to modulate the timing of fruit ripening [14]. A strawberry cytosolic ascorbate peroxidase (FaAPX) is strongly expressed in fruit at the turning (1/2 red) stage and weakly in leaf, root, and petiole, but not in seed [15]. To increase fruit firmness, an antisense strawberry endo-b-1,4-glucanase gene FaCel1(
) and an antisense strawberry cDNA of ADP-glucose pyrophosphorylase small subunit gene FaAGPS(
) were expressed under the control of fruit-specific FaAPX promoter [16, 17]. Unfortunately, the authors did not specify the size of the promoter. In transgenic strawberry the transcription of FaCel1 and FaAGPS was reduced in fruits at all the stages of development, and almost completely inhibited in the red stage. Fruit-specific downregulation of the FaAGPS and FaCel1 genes by using the FaAPX promoter is an effective technique for increasing fruit firmness in strawberries [16, 17].

Tissue-Specific and Developmentally-Regulated Plant Promoters

75

GalUR gene is involved in the biosynthesis of vitamin C in strawberry fruits. The activity of the FaGalUR promoter (
1149 to
2 from TIS) and its deletion variants were transiently assayed in strawberry cv. Chandler [18]. The promoter activity is restricted to ripe fruit not to leaves and is strictly dependent on light. The activity of the FaGalUR promoter in ripe strawberry fruit is comparable to that of the CaMV35S promoter (TGP Fa:GalUR). 1-aminocyclopropane-1-carboxylate oxidase (ACO) and polygalacturonase (PG) mRNAs were upregulated during apple (Malus  domestica) fruit ripening. Expression of the MdACO and MdPG genes was not detected in developing fruit or flowers. MdPG promoter fragments (
1460 to +340 and
532 to +340 from TSS) and MdACO1 promoter fragments (
1966 to +99 and
1159 to +99 from TSS) conferred ripening-specific expression in fruits of transgenic tomato cv. UC82B [19]. At breaker and subsequent ripening stages, expression was observed predominantly in the inner pericarp, vasculature and seeds. No expression was observed in columella or locular tissues of ripe fruit. No GUS activity was found with these promoter fragments in green fruit, roots, flowers, and leaves. The shortest
450 MdACO1 promoter fragment provided fruit, but not ripening specific expression. Tissue-specific GUS expression was downregulated by the longest
2356 MdPG promoter fragment, suggesting the presence of a negative regulatory element for the fruit-specific expression between positions
1460 and
2356 (TGP Md:ACO). Stearoyl-ACP desaturase (DES) is responsible for the production of fatty acids in an oil palm (Elaeis guineensis). EgDes promoter constructs were able to direct GUS expression in the mature green and ripe red stages of the transgenic tomato fruits and in the seeds [20]. Uniform staining was observed in the vascular, endocarp, mesocarp, columella, and placental tissues, not in the exocarp tissue of the tomato fruits. No GUS expression was detected in any of the floral or vegetative tissues from the transgenic tomato plants. The truncated (
590 to +10 from TSS) EgDes promoter showed the highest expression in most tissues compared with the full-length
1022 promoter and the
304 promoter. The GUS activity driven by the
590 EgDes promoter in fruits was 4.5-fold higher than that expressed via the CaMV35S promoter. The EgDes promoter was subjected to spatial regulation in tomato fruits, supporting the conclusion that EgDes promoter-mediated GUS expression is primarily specific to fruit (TGP Eg:Des). A type 3 metallothionein-like gene expression is abundant in Satsuma mandarin (Citrus unshiu) fruit. A full-length CuMT45 promoter (
4291 to
58 from TIS) activity was higher in juice sacs rather than in leaves of Valencia orange (Citrus sinensis) [21]. The CuMT45 promoter and the CaMV35S promoter showed similar levels of activity in juice sacs. The CuMT45 promoter conferred quantitatively preferential expression in siliques of transgenic

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Arabidopsis. These results might indicate that the cis-elements required for preferential expression in citrus fruits are functionally conserved in heterologous Arabidopsis plants and common regulatory mechanisms of gene expression operate during fruit development of these species. A large subunit of ADP-glucose pyrophosphorylase (AGPL) expressed specifically in fruits, not in leaves in watermelon (Citrullus vulgaris). A 1573 bp CvAGPL1 promoter (
1140 to +433 from TSS) can direct fruit-specific expression of GUS gene in transient expression assay in watermelon [22] and in transgenic tomato [23]. The CvAGPL1 promoter fragments of 1573, 1202, and 905 bp were able to direct GUS expression in immature green, mature green and red fruits, but not in leaves, stems and roots of tomato [24]. Frequently used a LeE8 promoter (
1532 to +24 from TSS) showed 28% less activity than the full-length CvAGPL1 promoter (
1244 to +433 from the TSS). The CvAGPL1 promoter region (
868 to +433 from the TSS) gave maximum promoter activity, 1.5-fold higher than that of the full-length promoter. A positive regulatory element is present in the region between
868 and
769. Two negative regulatory regions, from
986 to
959 and from
472 to
424, were identified in the CvAGPL1 promoter region by fine deletion analyses. Removal of both regions led to constitutive expression in epidermal cells. A TCCAAAA motif was identified in these two regions that functions as a fruit-specific element by inhibiting gene expression in leaves [24]. Sweet-taste tomato fruits were produced by expressing a Brazzein gene under control of the CvAGPL1 promoter. Other parameters of fruit quality were largely unchanged in CvAGPL1Brazzein plants [25]. In earlier studied, the CvAGPL1 promoter was referred to as WSP or Wml1promoter [22, 23]. An expansin gene from cucumber (Cucumis sativus) (CsExp) is highly and specifically expressed in ripened cucumber fruit. A 1.5 kb promoter region of CsExp displays a significant activity in fruits, not in leaves of cucumber in a transient expression assay [26]. The CsExp promoter activity in fruit is comparable to that of the CaMV35S promoter. A lumazine synthase gene (LS) from cucumber is highly expressed in cucumber fruit. The expression level of CsLS in the fruit was at least ten times higher than that in the stems and leaves. The 2.1-kb CsLS promoter was able to drive the high-level expression of the GUS gene in cucumber fruit, but not in stems or leaves [27]. Cucumisin, a subtilisin-like serine protease, is expressed at high levels in the fruit of melon (Cucumis melo). In transient expression analysis, a 50 -upstream region (1181-bp from TSS) of the Cucumisin gene was sufficient for the high level of GUS expression in immature melon fruits (10 days after pollination, DAP), but provided the very low level in leaves and stems [28]. The
234

Tissue-Specific and Developmentally-Regulated Plant Promoters

77

promoter fragment showed comparable level of activity to the fulllength
1181 promoter in fruits. GUS activity driven by the Cucumisin promoter was 2.5-fold lower than that driven by the CaMV35S promoter. A positive regulatory region is located between nucleotides
234 and
214. It contained a TGTCACA enhancer element involved in fruit-specific expression of the Cucumisin gene. The MADS-box domain transcription factor FRUITFULL (FUL) is important in development of all cell layers of the valves and normal siliques in Arabidopsis. A
3873 AtFUL promoter (
3938 to
66 from TIS) was inserted in front of a minimal 35S promoter and used to drive GUS expression in Arabidopsis [29]. Transcriptional activity of the full-length AtFUL promoter generally agreed with data from FUL mRNA in situ hybridizations assays. The GUS expression was detected in the inflorescence meristems, and throughout the developing carpel primordia before valve differentiation. A promoter deletion analysis was performed to identify promoter fragments for tissue-specific expression in silique. The 682 bp region, between positions
2952 and
2271, is required for AtFUL valve-specific expression in Arabidopsis [29] and in oilseed crop false flax (Camelina sativa) [30]. The AtFUL promoter fragment was successfully utilized to induce the expression of limonene synthase gene (LS) from lemon (Citrus limon) in false flax. Limonene synthase induced by the AtFUL promoter leads to higher limonene production in fruits than when the CaMV35S promoter was used to trigger the expression. The AtFUL promoter can be successfully utilized to induce the expression of the transgenes of interest in fruits of false flax [30]. A lycopene beta-cyclase promoter from a green-fruited Solanum habrochaites genotype EC520061 (ShCYC-B,
908 bp from TIS), showed low activity in leaves and stamens, and its activity was upregulated fourfold in fruits of transgenic tomatoes [31]. Deletion
908 to
437 bp 50 to ATG led to significant increase in GUS activity in flowers and leaves of the transgenic plants. The ShCYC-B promoter activity was lower than that of the CaMV35S promoter. A capsanthin–capsorubin synthase gene (CCS) regulates the development of red coloration in pepper (Capsicum annuum) fruits. Activity of the CaCCS promoter (2305 bp from TIS) has been found in fruit tissues but not in leaves in pepper [32] and transgenic tomato plants [33]. The CaCCS promoter was strongly upregulated during tomato fruit ripening in a manner similar to the induction of this gene in pepper fruits. Only low activity was found from ovaries of pollinated flowers up to the late immature green stage. A strong increase in activity was observed at a late mature green stage or later stages. Ethylene and water loss positively influenced the promoter activity in tomato [33]. Analysis of three tandem repeat units of the CaCCS promoter demonstrated that

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Olga G. Smirnova and Alex V. Kochetov

the constructs containing at least one repeat were associated with GUS expression similar to that provided by the full-length promoter in pepper and tomato fruits at the green ripening stage (TGP Ca:CCS) [34]. 2.2 Flower-Specific Promoters

An apple anthocyanidin synthase gene (MdANS) preferentially expressed in red-colored fruit skin. High level of the MdANS1 promoter activity in the reproductive organs, but low in the vegetative organs, of transgenic tobacco is consistent with the reproductive organ-preferential expression of MdANS1 [35]. GUS activity was detected in sepals of floral buds, petals of mature flowers, receptacles, and developing seeds of transgenic tobacco. In the sepals and the mature petals, GUS activity was found mainly in the trichomes and only weakly in the epidermal tissues. In the petals, vascular tissues also showed strong activity. In the developing fruits, activity was significant in the crenulated epidermal cells of immature seeds, but weak in the placenta. Strong activity was detected in the centers of the receptacles. Different types of tobacco and apple fruit development may explain weak GUS activity detected in the fruit skin of transgenic tobacco. Tobacco fruit is a dehiscent capsule that develops from an ovary while apple fruit is indehiscent flesh, being hypanthium developed to fruit parenchyma. Transformation of apple plants would provide a direct answer for the exact location of MdANS1 expression, and developing skin-specific promoter will be useful to enhance such biological traits as fruit color in the skin tissues (Table 1) [35]. A strong flower-specific enhancer may located between
610 and
403 bases of the peach PpACO1 promoter [12], because the
610 lines have very high expression in tomato flowers relative to all the PpACO1 promoter deletion variants and even relative to other tissues of tomato (stem, root, fruit) (TGP Pp:ACO1). A 1.2 kb promoter fragment of the AtFUL gene (
1255 to
66 from TIS) is capable of activating transcription in the inflorescence meristem and floral meristem in Arabidopsis [29]. A promoter of a phytoene desaturase gene of tomato (LePDS,
1549 to +564 from TSS) is able to drive developmentally regulated GUS expression in transgenic tomato and tobacco plants. In tomato, high levels of GUS expression are found in organs and at stages of development where chromoplasts are formed: petals, anthers, corolla, and ripening and red fruits. Tobacco petals and fruits, which do not contain chromoplasts, show instead low levels of GUS expression [36]. A cDNA of the CrtO gene from the alga Haematococcus pluvialis, encoding beta-carotene ketolase, was transferred to tobacco under the regulation of the 1038 bp LePDS promoter (
459 to +579 from TSS), which produced the highest level of expression in the petals [37]. Using metabolic engineering, the carotenoid biosynthesis pathway in tobacco (Nicotiana tabacum) was modified to produce astaxanthin, a red

Tissue-Specific and Developmentally-Regulated Plant Promoters

79

pigment of considerable economic value. Chromoplasts in the nectary tissue of transgenic plants accumulated astaxanthin and other ketocarotenoids, changing the color of the nectary from yellow to red. A truncated 733 bp LePDS promoter where 305 bp was removed from the 50 end demonstrated the highest level of expression in tobacco ovaries (TGP Le:PDS). 2.3 Anther/ Pollen-Specific Promoters

Anther/pollen-specific promoters may be used for genetically engineering male sterility in plants and to study the functions of pollen/ tapetum-specific genes (Table 2). A PsEND1 promoter (
2752 to
22 from TIS) was isolated from the anther-specific gene from pea (Pisum sativum cv. Alaska). The PsEND1 promoter is fully functional only in the anthers of transgenic Arabidopsis (cv. Columbia), tobacco (cv. Samsun), and tomato (cv. UC82b) plants [38]. GUS activity was detected from the very early stages of anther development and in the same anther tissues in which the END1 gene is usually expressed in pea: epidermis, connective, endothecium, and middle layer cells. The staminal filament, the central vascular cylinder, tapetum, the mother cells of the pollen, and adult pollen did not show any GUS activity. GUS activity was not observed in flowers, leaves, stems and roots of transgenic plants [38]. In transgenic bread wheat line Bobwhite,

Table 2 Promoters for anther/pollen-specific expression Promoter

Size

Origin

Tested in

Organ/tissue

Target gene

TGP ID

PsEND1

2731 bp Pea

Tomato, tobacco, Arabidopsis, rape, wheat, kalanchoe, pelargonium

Young pollen

barnase

Ps:END1

Ntg10

1407 bp Tobacco Tobacco

Pollen, tapetum

Zm908

2126 bp Maize

Tobacco

Pollen

LlPR10g

1233 bp Lily

Arabidopsis

Young pollen, tapetum

barnase

LGC1

946 bp

Lily

Tobacco, lily, onion

Pollen

DT-A

OsRTS

1.2 kb

Rice

Rice, creeping bentgrass, Tapetum Arabidopsis

Os03g11350 1805 bp Rice

Rice

Young pollen

Os10g22450 2 kb

Rice

Mature pollen

Rice, Arabidopsis

Pollen, tapetum

OsP12

Rice

1943 bp Rice

Nt:G10

barnase, RTS (
), LLA1271

BnCysP1

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Olga G. Smirnova and Alex V. Kochetov

GUS activity was also found in mature pollen, after anthesis [39]. The PsEND1 promoter was fused to the cytotoxic ribonuclease gene, barnase, from Bacillus amyloliquefaciens to induce specific ablation of the cell layers where the PsEND1 is expressed and consequently to produce male-sterile plants. Expression of the chimeric PsEND1-barnase gene in tobacco, tomato, Arabidopsis, rape (Brassica napus) [40], kalanchoe (Kalanchoe blossfeldiana), and pelargonium (Pelargonium zonale) [41, 42] impairs anther development from very early stages and produces complete malesterile plants (TGP Ps:END1). The use of the PsEND1 promoter instead of a tapetum-specific promoter arrests the microsporogenesis before differentiation of the microspore mother cells and no viable pollen grains are produced. This strategy represents an excellent alternative to generate genetically engineered male-sterile plants, which have proven useful in breeding programs for the production of hybrid seeds. A promoter (
1190 to +217 from TSS) of tobacco pectate lyase gene Ntg10 directed preferential GUS expression in pollen, with bimodal peaks of expression just before and during pollen mitosis I, and in mature anthers in tobacco [43]. Transient expression analysis defined the minimal Ntg10 promoter region capable of directing expression in pollen as
86 to +217 (TGP Nt:G10). A promoter (
2068 to +58 from TSS) from pollenpredominant small-peptide gene from maize Zm908 provided high GUS staining intensity in mature pollen grains, whereas no apparent GUS staining was detected in the other tissues in transgenic tobacco (cv. Xanthi-nc) [44]. The promoter deletion variants
1612 and
1126 showed strong GUS staining in the pollen as well as in seedling leaves. The results indicate the full-length Zm908 promoter is pollen-specific. A LlPR10g promoter (
1183 to +50 from TSS) of a tapetum/ microspore-specific pathogenesis-related 10 gene from lily (Lilium longiflorum) strongly and specifically expressed in young anthers, but not in the anther of mature flowers and in seedlings and vegetative organs in transgenic Arabidopsis [45]. The LlPR10gbarnase transgenic lines exhibited complete male sterility because of the disruption of the tapetum and the deformation of microspore/pollen. The decisive fragment required for anther specificity of the LlPR10g promoter is located between
1183 and
880 bp. A LGC1 promoter (
811 to +140 from TSS) of generative cellspecific gene from lily (L. longiflorum) provided pollen-specific GUS expression in transgenic tobacco [46]. No GUS expression was observed in the uninucleate microspores, vegetative cell in the mature pollen or any other vegetative and floral tissue. Transgenic tobacco plants carrying the diphtheria toxin A-chain- (DT-A)-coding region under the control of the LGC1 promoter showed normal phenotype except for anthers that contained sterile and aborted pollen. Truncations of the LGC1 promoter up to
242 showed a

Tissue-Specific and Developmentally-Regulated Plant Promoters

81

generative cell-specific expression similar to the full-length promoter. A tapetum-specific gene of rice, OsRTS, is predominantly expressed in tapetum during meiosis and disappears before anthesis [47]. A 1.2 kb OsRTS promoter, when fused to the barnase gene or the antisense of the RTS gene, is able to drive anther-specific expression of both genes in rice, creeping bentgrass (Agrostis stolonifera), and Arabidopsis, conferring male sterility to the transgenic plants [47]. Expression of an anther-specific gene LLA1271a from lily under the control of the OsRTS (TAP) promoter in Arabidopsis showed association of LLA1271a with exine formation during anther development [48]. LOC_Os03g11350 gene from rice encodes UDP-glucosyltransferase and showed immature panicle-preferred expression. The activity of the 1805 bp Os03g11350 promoter was restricted to young pollen at the uninucleate stage [49]. No GUS activity was observed in leaves, roots, or any other nonreproductive tissues from transgenic rice plants. In contrast, when Os10g22450 promoter (2 kb) was used, the GUS activity was detected only in mature pollen but not in anther walls, pollen tubes, lemma and palea of rice florets [49]. LOC_Os10g22450 encodes inositol-3-phosphate synthase, which mainly expressed in more than 10 cm panicles. A unique male-sterility and fertility-restoration system was developed in rice by combining a B. napus cysteine-protease gene (BnCysP1) with an anther-specific P12 promoter of rice (
1937 to +6 from TSS) for facilitating production of hybrid varieties. The OsP12 promoter was studied in Arabidopsis and rice systems [50]. 2.4 Seed-Specific Promoters

During seed development, seed-specific proteins are expressed in high levels. Many seed-specific promoters have been identified, such as those of the genes coding for glutenin and gliadin in wheat, zein in maize, lectin and legumin in pea, phaseolin and phytohemagglutinin in bean, glycinin and β-conglycinin in soybean, and glutelin and prolamin in rice. Arabidopsis nitrate transporter NRT2.7 plays a specific role in nitrate accumulation in seeds. AtNRT2.7 transcripts accumulate in seeds undergoing desiccation. High expression of the gene was detected in dry seeds. The AtNRT2.7 promoter (
2000 to
1 from TIS) provided high levels of GUS or GFP expression in transgenic Arabidopsis embryos while low expression was detected in the endosperm seed coat (Table 3) [51]. Anthocyanidin reductase encoded by the BANYULS (BAN) gene is the core enzyme in proanthocyanidin biosynthesis. In Arabidopsis, proanthocyanidins are found only in the seed coat, where they confer a brown color to mature seeds after oxidation. AtBAN promoter (
2332 to +44 from TSS) activity was detected specifically in developing Arabidopsis seed coat [30]. Genetic ablation of

Rice

Rice

Rice

2376 bp 324 bp

495 bp

2335 bp

1474 bp

836 bp

931 bp

981 bp

1249 bp

AtBAN

PhFBP7

OsGluB-1

OsGluB-4

OsPro10

OsPro16

OsGLB1

OsOLE18

Os02g15090 1839 bp

Os06g31070 1678 bp

Tested in

1249 bp, Wheat 688 bp

TaHMWGS 1Dx5 Wheat, tobacco

Wheat

2175 bp

TaHMWGS 1Bx7

Wheat

Wheat, rice, barley

Rice

Rice

Rice

Rice

Rice

Rice

Rice

Rice

Petunia Strawberry

TaHMW1892 bp, Wheat GS 1Bx17 1.3 kb

Rice

Rice

Rice

Rice

Rice

Petunia

Arabidopsis Arabidopsis, rape, false flax

Arabidopsis Arabidopsis

2000 bp

AtNRT2.7

Origin

Size

Promoter

Table 3 Promoters for seed-specific expression

Mid

strong

Endosperm

Endosperm

Endosperm

Seed

Endosperm, embryo

Embryo, aleurone

Strong

Strong

Strong

Strong

Strong

Weak

Endosperm, aleurone Strong

Endosperm, aleurone Strong

Endosperm, aleurone Strong

Endosperm, aleurone Strong Vegetative tissues

Endosperm, seed coat

Os:OLE18

Os:GLB1

Os:PROLAMIN16

Os:PROLAMIN10

Os:GLUB4

Os:GLUB1

TGP ID

TaLMW-GS, TaHMW-GS 1Dx5, y1, XynA, Fae, SSA

Ta:HMWGlu1Dx5

Ta:HMWGlu1Bx7

AmA1, CTB, HIVDR, Ta:HMWGlu1Bx17 SSA, TaHMW-GS 1Bx17

RINO1

barnase

barnase, ClLS

Activity Other tissues Target gene

Endosperm, aleurone Strong

Seed coat Flower, fruit, root, petiole

Seed coat

Embryo

Main organ/tissue

82 Olga G. Smirnova and Alex V. Kochetov

646 bp

875 bp

2147 bp

2669 bp 2164 bp

2158 bp

1591 bp

1545 bp

1144 bp

2032 bp

2360 bp

TaEm

TaALP type-B

TdPR60

TdPR61

PsPSL

CtMS

β-phaseolin

BnNapA

Linin

CuMFT1

Mandarin

Flax

Rape

Bean

Guar

Pea

Durum wheat

Durum wheat

Wheat

Wheat

Endosperm transfer cells Endosperm

Endosperm

Orange, mandarin, Arabidopsis

Arabidopsis

Arabidopsis

Alfalfa Arabidopsis

Alfalfa

Tobacco

Seed

Seed

Seed

Embryo Seed

Endosperm

Seed

Wheat, barley, Endosperm transfer rice cells, embryo

Rice

Wheat, barley

Tobacco

Wheat, barley, Embryo, aleurone rice Tobacco Embryo

Strong

Strong

Strong

Strong

Mid

Strong

Mid

Mid

Mid

Leaf, juice sac, style of flower

Endosperm

Embryo

Endosperm transfer cells

Endosperm

Root

PgFADX

PgFADX

PgFADX

Ps:PSL

Td:PR60

Ta:Em

Tissue-Specific and Developmentally-Regulated Plant Promoters 83

84

Olga G. Smirnova and Alex V. Kochetov

proanthocyanidin-accumulating cells targeted by the AtBAN promoter fused to barnase led to the formation of normal plants that produced viable yellow seeds that had no obvious defects in endosperm and embryo development. The 324 bp AtBAN promoter fragment (
280 to +44 from TSS) was successfully utilized to induce the expression of limonene synthase gene from lemon (Citrus limon, ClLS) in an oilseed crop false flax (Camelina sativa). Limonene synthase induced by the AtBAN promoter leads to higher limonene production in seeds than when the CaMV35S promoter was used [30]. The AtBAN promoter can be successfully utilized to induce the expression of the transgenes of interest in seeds of false flax. The full-length AtBAN promoter triggers the expression of the GUS reporter in seed coat of rape (B. napus) [52]. An ovule-specific petunia (Petunia  hybrida) MADS box gene PhFBP7 ( floral binding protein7) is expressed in carpel, not in leaves, sepals, petals, or stamens [53, 54]. In transgenic petunia, the 495 bp PhFBP7 promoter activity is restricted to the seed coat of developing seeds and it is completely silent in the embryo and endosperm. The activity of the FBP7 promoter was also monitored by fusing it to the bacterial barnase gene. The PhFBP7 promoter activity coincides exactly with the pattern of PhFBP7 gene expression during ovule and seed development [54]. In transgenic strawberry cv. Gariguette, the PhFBP7 promoter activity was found in floral and fruit tissues of all developmental stages tested, in root and petiole [55]. In red strawberry tissue, the CaMV35S promoter is at least sixfold stronger than the PhFBP7 promoter. Seed storage glutelin promoters GluB-1 (
2291 to +44 from TTS), GluB-4 (
1461 to +13 from TSS) and prolamin promoters 10 kDa (OsPro10,
836 to
1 from TIS), 16 kDa (OsPro16,
931 to
1 from TIS) of rice directed strong GUS expression in endosperm, aleurone and subaleurone, but not in the embryo in transgenic rice plants [56]. 26 kDa alpha globulin gene promoter OsGLB1 (
981 to
1 from TIS) directed GUS expression in the inner starchy endosperm tissue [56]. During seed maturation, these promoters exhibited a high level of activity. The average GUS activities for these promoters in maturing seeds (17 DAF) were about 21, 45, 39, 27, 29 pmol (4 MU/min/μg protein), respectively. GUS activity expressed from the maize ubiquitin promoter, which was used as a control, was about 7 pmol (4 MU/min/ microg protein). GUS activity was also detected in vegetative tissues in the transgenic rice carrying the OsPro10 promoter [56]. The OsPro10 gene required both the 50 and 30 flanking regions for intrinsic endosperm-specific expression (TGP Os:GLUB1, Os: GLUB4, Os:PROLAMIN10, Os:PROLAMIN16, Os:GLB1). GUS expression driven by the promoter of the rice 18-kDa oleosin gene OsOle18 (
1249 to
1 from TIS) was restricted to the embryo and aleurone with no expression in the endosperm [56]. The activity of the OsOle18 promoter in maturing seeds was

Tissue-Specific and Developmentally-Regulated Plant Promoters

85

relatively low, about 5 pmol (4 MU/min/μg protein) (TGP Os: OLE18). A rice 1D-myoinositol 3-phosphate synthase gene (RINO1) is highly expressed in developing seed embryos and in the aleurone layer, where phytic acid is synthesized and stored. To suppress phytic acid biosynthesis effectively, antisense RINO1 cDNA was expressed under the control of the OsOle18 promoter [57]. Seed phytic acid was reduced by 68% and free available phosphate was concomitantly increased in the generated transgenic rice plants. No negative effects on seed weight, germination, or plant growth were observed. Higher repression of RINO1 gene expression was achieved using the antisense RINO1 cDNA fused to the promoter of the OsOle18 than that fused to the OsGluB-1 promoter. Os02G15090 and Os06g31070 promoters of rice (1839 and 1678 bp, respectively) drove GUS expression in a seed-preferred manner in transgenic rice plants [49]. No GUS activity was observed in leaves, stems, roots and panicles. GUS expression patterns driven by the promoters were similar to the expression patterns of their endogenous genes. The gene LOC_Os02g15090 encodes glutelin and LOC_Os06g31070 encodes a prolamin precursor. A promoter (
1892 to
1 from TIS) of a gene that encodes 1Bx17 subunit of high molecular weight storage protein glutenine of wheat (Triticum aestivum, TaHMW-GS) was tested by a transient expression assay in immature wheat endosperm (cv. Bobwhite) and in stable transgenic rice plants (cv. Taipei 309) [58] (TGP Ta:HMWGlu1Bx17). A short fragment of the TaHMW-GS 1Bx17 promoter (173 bp) connected with the first intron of the rice actin gene has the same expression level as the long 1892 bp TaHMW-GS 1Bx17 promoter without the intron [58]. Expression of an amaranth (Amaranthus hypochondriacus) albumin gene AmA1, encoding the protein with a high content of essential amino acids, under the control of the endospermspecific TaHMW-GS 1Bx17 (1.8 kb) essentially increased amino acid content and some parameters associated with functional quality in bread wheat (cv. Cadenza) [59]. A high level of synthetic cholera toxin B subunit gene (CTB) was expressed under control of the wheat TaHMW-GS 1Bx17 promoter in the endosperm tissue of the transgenic rice plants. The successful expression of CTB gene (2.1% of total soluble protein) in transgenic plants makes it a powerful tool for the development of a plant-derived edible vaccine [60]. By the use of HIVDR (human immunodeficiency virus diagnostic reagent) gene construct with the wheat TaHMW-GS 1Bx17 promoter, high-level expression of a fully functional antiglycophorin single-chain antibody fused to an epitope of HIV was obtained in seeds of barley [61]. The full-length TaHMW-GS 1Bx7gene promoter (
2175 to
1 from TSS) significantly increases the GUS expression compared

86

Olga G. Smirnova and Alex V. Kochetov

to the partial promoter in transient expression assay of un-matured, 20 DAF, wheat seeds cv. Chinese Spring [62]. Therefore, the distal region of the TaHMW-GS 1Bx7 promoter contains key regulatory elements (TGP Ta:HMWGlu1Bx7). Endosperm-specific promoter TaHMW-GS 1Dx5 (
1191 to +58 from TSS) [63], was used for the expression of three genes encoding low-molecular-weight (LMW) glutenin subunits, C-terminally tagged by the c-myc epitope, in pasta wheat (Triticum turgidum subsp. durum cv. Svevo and Ofanto) [64]. The use of a specific antibody to the c-myc epitope tag allowed the transgene products to be detected readily in the complex mixture of LMW subunits (TGP Ta:HMWGlu1Dx5). Transgenic wheat has been generated by expressing the maize y1 gene encoding phytoene synthase driven by the endosperm-specific TaHMW-GS 1Dx5 promoter in the elite wheat variety EM12, together with the bacterial phytoene desaturase crtI gene from Erwinia uredovora under the constitutive CaMV35S promoter control [65]. A clear increase of the carotenoid content (up to 10.8-fold) was detected in the endosperms of transgenic wheat that visually showed a light yellow color. A deletion variant of the TaHMW-GS 1Dx5 promoter (
649 to +39 from TSS) showed endosperm-specific activity in tobacco (cv. Petit Havana, mutant SR1) that was approximately ten times greater than the activity of the CaMV35S promoter in tobacco seeds [66]. The 277 bp immediately upstream of TSS of the TaHMW-GS 1Dx5 gene is sufficient for temporal and tissue-specific regulation in tobacco. Expression of the TaHMW-GS 1Dx5 gene under the control of its own promoter resulted in increased dough strength and stability in pasta wheat [67]. The XynA endoxylanase gene from Bacillus subtilis or the Fae ferulic acid esterase gene from Aspergillus niger, both coding cell wall degrading enzymes, were successfully expressed in bread wheat under the control of the endosperm-specific TaHMW-GS 1Dx5 promoter in order to improve the functional properties of the wheat grain for baking and animal feed [68]. Sulfur amino acid composition is an important determinant of seed protein quality. A chimeric gene encoding sunflower sulfur-rich seed albumin (SSA) under control of either 1.3 kb of the TaHMWGlu1Dx5 promoter [63] or 1.3 kb of the TaHMWGlu1Bx17 promoter [69] was introduced into rice (Oryza sativa) in order to modify cysteine and methionine content of the seed. Analysis of a transgenic line expressing SSA at approximately 7% of total seed protein revealed that the mature grain showed little change in the total sulfur amino acid content compared to the parental genotype. This result indicated that the transgenic rice grain was unable to respond to the added demand for cysteine and methionine imposed by the production of SSA. Early methionine (Em) protein is the most abundant polypeptide in mature wheat embryos. TaEm promoter (
554 to +92 from TSS) drove GFP expression in the embryo in transgenic tobacco.

Tissue-Specific and Developmentally-Regulated Plant Promoters

87

Transformed tobacco seeds had very low GUS activity prior to 16 days after anthesis (daa) but accumulated GUS to extremely high levels by 20 daa, before complete desiccation (29,000 pmol 4 MU/min/μg protein) [70]. The TaEm promoter directed expression of GFP in the embryo and aleurone, but not in the endosperm, floret and leaf in transgenic rice, barley, and wheat [71, 72]. In transgenic rice, the TaEm promoter activity was also detected in the roots (TGP Ta:Em) [71]. Genes for avenin-like proteins (ALP) represent a new family of wheat storage protein genes. A TaALP type-B promoter (
785 to
1 from TIS) provided GUS activity only in the endosperm, while GUS expression was not observed in any other organ, such as leaf, root, stem or flower of transgenic tobacco plants [73]. The 785 bp TaALP type-B promoter activity (9 nmol 4 MU/h/mg protein) was one half of that of the CaMV35S promoter. The full-length promoter (1664 bp) had the lowest expression levels. TdPR60 promoter (
2147 to
1 from TIS) of durum wheat provided GUS expression in bread wheat (cv. Bobwhite) and barley (Hordeum vulgare cv. Golden Promise) endosperm transfer cells only at nine DAP and could be detected until at least 40 DAP [74]. The weak GUS activity was also detected in adjacent layers of starchy endosperm cells. In rice the TdPR60 promoter was activated in a different way than in wheat and barley. The GUS activity was found mainly inside the starchy endosperm and the weak activity was found also in the layer of endosperm transfer cells (TGP Td:PR60). A lipid transfer protein gene promoter of T. turgidum (TdPR61,
2669 to
1 from TIS) provided strong GUS expression in the endosperm transfer cells, the embryo surrounding region, and in the embryo in transgenic bread wheat (cv. Bobwhite), barley (cv. Golden Promise) and rice (cv. Nipponbare) [75]. The TdPR61 promoter deletion variants were tested in developing wheat embryo using transient expression assays. The activity of the
2164 TdPR61 promoter deletion variant was roughly the same as the activity of the full-length promoter. The TdPR61 promoter will be a useful tool for improving grain quality by manipulating the quality and quantity of nutrient/lipid uptake to the endosperm and embryo. A pea lectin gene (PsPSL) encodes an abundant seed protein. The PsPSL promoter (
2116 to +42 from TSS) provided highlevel seed-specific expression pattern in transgenic tobacco plants (7125 pmol 4 MU/min/mg protein) [76]. Progressive 50 promoter deletions resulted in a gradual decrease of transcriptional activity in tobacco seed. A fragment of 115 bp still conferred seed-specific expression albeit at a low level (TGP Ps:PSL). A guar (Cyamopsis tetragonoloba) mannan synthase promoter (CtMS,
1591 to
1 from TIS) drives strong GUS expression (125 pmol 4 MU/min/μg) specifically in endosperm in transgenic

88

Olga G. Smirnova and Alex V. Kochetov

alfalfa (Medicago sativa R2336) [77]. The potential strength and specificity of the CtMS promoter was compared with those of the constitutive CaMV35S promoter and the seed specific β-phaseolin promoter. CaMV35S promoter drove almost undetectable GUS expression in endosperm of alfalfa. GUS expression driven by the β-phaseolin promoter in endosperm of alfalfa was approximately three times lower than that of the CtMS promoter. Thus, the CtMS promoter can be generally useful for directing endospermspecific expression of transgenes in legume species. A 1144 bp promoter of B. napus napin gene (NapA), a 1548 bp promoter of bean (Phaseolus vulgaris) phaseolin gene, a 2032 bp promoter of Linum usitatissimum linin gene and a 1119 bp promoter of L. usitatissimum conlinin gene were used for overexpression of pomegranate (Punica granatum) fatty acid conjugase in the seed oil of Arabidopsis [78]. Among the four promoters of genes encoding seed storage proteins, the linin promoter led to the highest accumulation of punicic acid (13.2% of total fatty acids in the best homozygous line). Analysis of the relative expression level of PgFADX in developing seeds suggested the linin promoter to be the best choice in Arabidopsis and possibly in other oilseed crops such as B. napus. A FLOWERING LOCUS T homologue gene (MFT1) promoter (
2396 to
37 from TIS) was isolated from Satsuma mandarin (Citrus unshiu cv. Miyagawa-wase) [79]. The CuMFT1 promoter activity was more than 15-fold higher in seeds than in leaves and juice sacs in Citrus. In transgenic Arabidopsis GUS staining was observed only in seeds, and no staining was detected in leaves, stems, and roots. Slight staining was observed in the style of flowers after bloom. 2.5 Root-Specific Promoters

Root-preferential promoters can be used to drive root-specific gene expression to improve the root resistance to pathogens, pests, nutritional and water deficiency, and salt, cold, or drought stress. The Arabidopsis NRT2.1 gene encodes a nitrate transporter involved in high-affinity uptake by the roots. The AtNRT2.1 promoter (
1201 to
1 from TIS) activity was predominantly located in the cortical zone of the older parts of the roots in transgenic Arabidopsis [80]. In the shoots, the AtNRT2.1 promoter activity was very low and was restricted to leaf hydathodes. The AtNRT2.1 promoter activity was upregulated in the presence of sucrose and nitrate, and repressed by downstream N metabolites. A 150 bp sequence (from
245 to
95) is required for these regulations. In transgenic tomato plants, the AtNRT2.1 promoter (
1154 to
65 from TIS) activity was observed in vascular regions of the roots but was conspicuously absent in the leaves. Expression of a synthetic chitinase gene (NIC) under the control of the AtNRT2.1 promoter showed increased levels of resistance against root pathogen Fusarium oxysporum compared to nontransgenic tomato plants [81] (TGP At:NRT2.1) (Table 4).

1062 bp, 852 bp

774 bp

GmPRP2

AbH6H

Arabidopsis, soybean

Arabidopsis

Tobacco

Tobacco

Belladonna Henbane, belladonna, tobacco

Soybean

Red Swiss chard

1670 bp

Root

Root

Root

Main organ/ tissue

Hairy roots

Root, hypocotyl

Root, petiole

Root

Root

Root Constitutive

Tobacco, alfalfa, Root tomato

Rice

Strawberry Strawberry Tobacco

BvcPPO

2930 bp

FaRB7

Tobacco

Eastern white pine

706 bp, 636 bp

TobRB7

Rice

865 bp

1.3 kb, 2002 bp

RCc3

Lotus Tobacco japonicus

PsPR10

1029 bp

LjNRT2

Tested in

Arabidopsis Arabidopsis, tomato

Western white pine

1200 bp, 1089 bp

AtNRT2.1

Origin

PmPR10–1.14 1744 bp

Size

Promoter

Table 4 Promoters for root-specific expression

Mid

Strong

Strong

Strong

Strong

Strong

Mid

Strong

Activity

Tapetum, pollen

Petiole, main veins

Decrease in petioles after flowering

Seedling

Leaf, stem, flower

Petiole

Leaf

Leaf hydathode

Other tissues

Lj:NRT2

At:NRT2.1

TGP ID

(continued)

Gm:PRP2

Bvc:PPO

Pst:PR10

Fa:RB7

PaCS, AtThi2.1, MjTis11, Nt:RB7 Sarcotoxin IA

OsNAC10, OsNAC9, OsNAC5, OsCKX4, ScYCF1, OsABCC1

NIC

NIC

Target gene

Tissue-Specific and Developmentally-Regulated Plant Promoters 89

Size

1441 bp, 363 bp

more than 1 kb

1164 bp

1167 bp

2413 bp

2 kb

1.5 kb

1867 bp

Promoter

AbPMT1

HnTr1

SbPRP1

MeGBSSI

SlREO

ZmRCP-1

At1g74770

OsHPX1

Table 4 (continued)

Tested in

Plantain

Tomato

Cassava

Tobacco, cowpea

Henbane

Rice

Rice

Arabidopsis Arabidopsis

Maize

Tomato

Cassava

Soybean

Henbane

Belladonna Belladonna

Origin

Root

Root

Root

Root

Tuberous root

Root

Hairy roots (endodermis, pericycle)

Root

Main organ/ tissue

Strong

Strong upon nematode infection

Strong

Strong

Mid

Strong

Mid

Activity

Anther, leaf

Root, stem, leaf

Leaf

Other tissues

splicing factor

Target gene

Le:REO

TGP ID

90 Olga G. Smirnova and Alex V. Kochetov

Tissue-Specific and Developmentally-Regulated Plant Promoters

91

LjNRT2 promoter (1029 bp) from Lotus japonicus also expressed NIC transgene in a root-specific manner in tobacco. Transgenic tobacco plants showed increased resistance against F. oxysporum [81] (TGP Lj:NRT2). These results suggest that NIC gene triggered by the root-specific AtNRT2.1 and LjNRT2 promoters successfully expressed only in the roots and conferred increased levels of resistance against the root pathogen F. oxysporum. RCc3 mRNA was highly expressed only in root tissues of rice seedlings [82]. Transgenic rice plants harboring OsCKX4 under the control of 2002 bp root-specific RCc3 promoter displayed enhanced root development without affecting their shoot parts, suggesting that this strategy could be a powerful tool in rice root engineering [83]. Root-specific overexpression of OsNAC10, OsNAC9, OsNAC5, and OsNAC6 genes under control of the 1.3 kb RCc3 promoter enlarges roots, enhancing drought tolerance of transgenic rice plants, which increases grain yield significantly under field drought conditions [84–87]. Transgenic rice plants expressing two different vacuolar arsenic sequestration genes, ScYCF1 and OsABCC1, under the control of the RCc3 promoter, along with a bacterial γ-glutamylcysteine synthetase driven by the maize UBI promoter, were developed. The transgenic plants exhibited a 70% reduction in arsenic accumulation in the brown rice without jeopardizing agronomic traits. This technology could be used to reduce arsenic intake in populations suffering from arsenic toxicity and thereby improve human health [88]. A tobacco tonoplast intrinsic protein gene (TobRB7) is expressed at high levels in root meristematic and immature central cylinder regions. TobRB7 mRNA is not detected in expanded leaf, stem, or shoot apex tissue. Root-specific GUS activity driven by the TobRB7 promoter (
636 to +70 from TSS) was observed in transgenic tobacco seedlings as early as 2 days postgermination [89]. In lateral roots, expression began at initiation and remained in the meristem and developing central cylinder. GUS activity driven by the
636 promoter was threefold higher than that driven by the
813 promoter. Construct including
299 bp promoter sequence exhibited a loss in GUS expression in roots (TGP Nt:RB7). Transgenic alfalfa (Medicago sativa) was engineered by introducing the Pseudomonas aeruginosa citrate synthase gene (CS) controlled by the TobRB7 root-specific promoter to help achieve aluminum tolerance [90]. Transgenic tomato plants expressing an Arabidopsis thionin gene (Thi2.1) driven by the TobRB7 promoter conferred significant levels of enhanced resistance to bacterial wilt and Fusarium wilt. In the transgenic lines, AtThi2.1 expression was detected in roots and incidentally in leaves, but not in fruits. Potential thionin toxicity in fruits was negated by the use of a fruit-inactive TobRB7 promoter to drive the AtThi2.1 gene [91]. A fragment of Sarcotoxin IA gene, encoding an antibacterial protein of

92

Olga G. Smirnova and Alex V. Kochetov

Sarcophaga peregrina, was ligated to the 30 end of the 450-bp TobRB7 promoter [92]. Transgenic tomato plants expressing the sarcotoxin IA gene exhibited strong inhibition of parasite broomrape (Orobanche aegyptiaca) growth and significantly increased yield as compared with nontransgenic ones. A tonoplast intrinsic protein gene RB7 was isolated from strawberry (Fragaria  ananassa). A FaRB7 promoter (
2843 to +87 from TIS) was shown to direct strong, near root-specific expression in transgenic strawberry plants with expression patterns very similar to that of the endogenous gene [93]. Strong blue GUS staining was observed in young and old root tissues, light-blue staining was observed in petioles and no staining was observed in leaves, floral organs, and fruit of transgenic strawberry plants. The FaRB7 promoter was found to confer constitutive expression, comparable to that produced by the CaMV35S promoter, in tobacco (TGP Fa: RB7). A pathogenesis-related gene PR10–1.14 was expressed in lateral roots and needles of western white pine (Pinus monticola) and its promoter established a root-specific expression in transgenic tobacco [94]. GUS activities, driven by the full-length PmPR10–1.14 promoter (
1675 to +69 from TSS), were hundred times higher in the roots than that in the leaves, stems and flower organs, including sepals, petals, stamens and pistils. The distal 50 deletion of the promoter to
311 did not decrease the expression level in the roots significantly. A promoter of a pathogenesis-related PsPR10 gene was isolated from eastern white pine (Pinus strobus) [95]. The
796 PsPR10 promoter (
796 to +70 from TSS) drove GUS expression at high levels in the transgenic tobacco roots (threefold higher than the activity of the CaMV35S promoter in the roots). In plantlets, GUS staining was first detected in the hypocotyl and embryo. More GUS staining was observed in leaves, stems and roots at two and three leaves stages. At five leaves stage and in mature plants, in situ GUS staining was only detected in the primary root, no GUS staining was observed in leaves and stems. GUS activity driven by the
796 promoter in transgenic tobacco roots was similar to that of the
1681 promoter and higher than those of the
513 and
323 promoters (TGP Pst:PR10). A 1670 bp 50 -flanking region (
1648 to +22 from TSS) of the polyphenol oxidase gene (PPO) was isolated from red Swiss chard (Beta vulgaris subsp. Cicla), a betalain-producing plant. The BvcPPOP promoter could direct GUS expression in vegetative organs with root and petiole preference, but not in reproductive organs including inflorescences shoot, inflorescences leaf, flower, pod and seed [96]. Measured at 15 days old, the GUS enzymatic activities in the root and petiole were 13- and tenfold higher than that in the leaf, respectively. At flowering and after anthesis, the GUS staining in the petiole and the main veins of the rosette leaf

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faded progressively, although the GUS staining in root remained strong. 50 -truncated versions of the promoter varied in the driving strength, but not obviously in expression pattern, and even the shortest version (
225 to +22) retained the root- and petiolepreference (TGP Bvc:PPO). A proline-rich protein gene (PRP2) expression level was the highest in the root, second in the hypocotyl, seed and stem and much lower in the leaf and flower in soybean (Glycine max). A GmPRP2 promoter (
1062 to
1 from TIS) showed a rootpreferential expression in transgenic Arabidopsis [97]. GUS activity was mainly detected in roots and hypocotyls. GUS staining was also detected in the petiole during the reproductive growth stage and in the main veins in transgenic lines with the
852 promoter construct, which exhibited high expression levels. GUS staining was not detected in the cotyledons, apical roots, flowers and seeds. GUS activity was higher in transgenic Arabidopsis and soybean hairy roots with
1062 and
852 GmPRP2 promoter constructs than with the shorter promoter constructs. The promoter is responsive to salt, drought and hormone treatments (TGP Gm:PRP2). Hyoscyamine 6 beta-hydroxylase (H6H) participates in scopolamine synthesis in belladonna (Atropa belladonna). AbH6H RNA was detected in cultured root, native root and anther, but not in stem, leaf, pistil, petal, and sepal tissues. An AbH6H promoter (
671 to +103 from TSS) was sufficient for pericycle-specific expression in hairy roots of belladonna and henbane (Hyoscyamus niger), which both produce scopolamine [98]. GUS expression driven by the AbH6H promoter was generally stronger in henbane roots than in belladonna roots. The aerial parts, except for flowers, of transgenic belladonna plants did not show any GUS staining. In opened mature flowers, GUS activity was detected in tapetum and in pollen grains. In transgenic tobacco hairy roots, which do not produce scopolamine, GUS staining was not restricted to the pericycle. Some, but not all, cells in cortex and epidermis were clearly stained, along with cells in the pericycle and root apex. Putrescine N-methyltransferase (PMT) catalyzes the first committed step in the biosynthetic pathways of tropane alkaloids in belladonna. An AbPMT1 promoter (
1373 to +68 from TSS) drove GUS expression specifically in root pericycle cells, not in the leaves, stems, and flowers in transgenic belladonna [99]. The strength of the AbPMT1 promoter appeared to decrease in accordance with the progressive 50 truncation, but the 295-bp 50 -upstream region of the AbPMTl promoter was still sufficient for the pericycle-specific expression. More than 1 kb 50 -upstream sequences of the tropinone reductase genes HnTR1 and HnTR2 from henbane conferred similar patterns on the reporter gene expression in henbane transgenic hairy roots [100]. Both HnTR1 and HnTR2 promoters were active in the root endodermis and pericycle. The reporter activities were

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high in mature roots, and gradually decreased toward the younger roots branched from the mature root. The HnTR1 promoter appeared to have about twice as high transcriptional activities as the HnTR2 promoter. Although different activity levels have been observed for the HnTR promoters in the henbane root, the HnTR genes are transcribed at a similar rate. SbPRP1 is a member of the soybean proline-rich cell wall protein family and is expressed at high levels in root tissue. GUS expression driven by the SbPRP1 promoter (
1084 to +5 from TSS) in transgenic tobacco roots was 4.5-fold higher than that found in the leaves [101]. 50 -Deletion analysis of the SbPRP1 promoter indicated that a minimal promoter for GUS expression in roots is located within the first 262 bases of the SbPRP1 upstream sequence. The SbPRP1 promoter is expressed most strongly in the epidermis in the apical and elongating region of both primary and lateral roots. A similar localization pattern was found in transformed cowpea (Vigna aconitifolia) hairy roots. A promoter of a granule-bound starch synthase (GBSSI) gene was isolated from cassava (Manihot esculenta) [102]. The MeGBSSI promoter (
1107 to +57 from TSS) drove very low firefly luciferase gene expression in leaves, stems and roots, but very high expression in tuberous roots in the transgenic cassava. The luciferase activity in the tuberous roots was 800–8000 times higher than in the other tissues. These results corroborate the results of cassava GBSSI gene expression [103]. Using the MeGBSSI promoter to drive expression of target genes should result in stable and high expression of transgenes in cassava tuberous roots [102]. A promoter (
2413 to
1 from TIS) of a tomato Fe (II)dependent dioxygenase gene (SlREO) was shown to direct expression in the root cortex of transgenic tomato plants [104]. GUS staining was not detectable in leaves or flowers. GUS activity in the roots was 118-fold greater than the activity in the leaves. The SlREO promoter has properties ideal for strong and specific gene expression in tomato roots (TGP Le:REO). A maize root cap-specific protein-1 promoter (ZmRCP-1, 2 kb upstream of TIS) delivered GUS expression in roots but not in leaves of transgenic plantains [105]. In mature old roots, GUS expression becomes limited to the root cap. Invasion by the nematode Radopholus similis does not modify the ZmRCP-1 promoter activity. The ZmRCP-1 promoter can provide root tip-specific expression of transgenes in a monocot plant, such as plantain. An 1.5 kb nematode-responsive and root-specific promoter AtNRRS, derived from At1g74770 gene, provided strong GUS activity throughout the root upon nematode (Meloidogyne incognita) infection [106]. Transgenic Arabidopsis plants expressing dsRNA of a nematode splicing factor gene under the AtNRRS promoter exhibited up to a 32% reduction in number of galls as compared to 71% reduction with use of CaMV35S promoter.

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However, the use of CaMV35S promoter has to be exercised with caution since dsRNA is produced in all tissues all the time and can lead to undesirable effects in transgenic plants. A rice heme peroxidase gene (OsHPX1) expressed predominantly in roots. A OsHPX1 promoter (1867 bp from TIS) is predominantly active in the root elongation region during the vegetative stage of growth of transgenic rice plants [107]. GFP fluorescence levels, driven by the OsHPX1 promoter, were high in root elongation regions but not in root apex and cap of the transgenic plants. Very low levels of GFP fluorescence were observed in anthers and leaves. The OsHPX1 promoter activities were 16- to 190-fold higher in roots than in leaves of transgenic rice plants. 2.6 Developmentally Regulated Promoters

A promoter of Arabidopsis senescence-associated gene AtSAG12 is the most popular and well-studied developmentally regulated plant promoter (TGP At:SAG12). SAG12 did not express in young leaves and its expression increased continuously as Arabidopsis leaves progressed through senescence [108, 109]. The AtSAG12 promoter (
2073 to +106 from TSS) conferred senescence-specific expression in Arabidopsis [110], tobacco [111–114], rice [115], lettuce (Lactuca sativa cv. Evola) [116], petunia [117], tomato [118, 119], wheat [120], creeping bentgrass [121, 122], cassava [123], gerbera [124], pelargonium [42]. In transient expression assays, the AtSAG12 promoter (
1345 to +87 from TSS) drove strong GUS expression in senescence leaves and flower, and weak expression in young leaves, fresh flowers and roots from petunia. The promoters can also drive GUS expression in senescence corollas of daffodil, daylily, and orchid, although the level of GUS expression was relatively low in orchid [125]. An AtSAG12-IPT gene construct, which expressed the Agrobacterium tumefaciens cytokinin biosynthesis IPT gene in response to senescence, was introduced into different plant species to delay senescence (Table 5). An expression of BnSAG12–2 gene from B. napus reached a maximum level in the early stage of leaf senescence and decreased slightly in the middle and in the final stage of senescence. This is in contrast to the expression of AtSAG12 that is increased continuously as Arabidopsis leaves progressed through senescence. In transgenic Arabidopsis, the 3.11 kb BnSAG12–2 promoter had an expression level, which is weaker than that of the AtSAG12 promoter (1345 bp from TIS), but was regulated identically with the native AtSAG12 gene [126]. Senescence-specific BnSAG12–2-GUS expression was also evident in floral parts of transgenic Arabidopsis. Therefore, BnSAG12–2 is senescence specifically regulated in Arabidopsis. The metallothionein gene, LSC54, shows increased expression during leaf senescence in B. napus. GUS expression levels, driven by the 1055 bp BnLSC54–2 promoter, increased during Arabidopsis leaf development of approximately 750-fold between young green

Petunia, carnation

914 bp

2379 bp Rice

1976 bp Maize

2305 bp Arabidopsis Arabidopsis, tomato

1400 bp Arabidopsis Arabidopsis

1850 bp Arabidopsis Arabidopsis

2.1 kb

840 bp

2296 bp Pepper

MjXB3

OsSGR

ZmSEE1

AtBFN1

AtCAT2

AtCAT3

PpACO2

PvSARK

CaFIB

Cotton, maize

Peach

Tomato, pepper, potato, Arabidopsis, tobacco

Tobacco, rice, peanut,

Tomato

Maize, ryegrass

Rice

Arabidopsis

BnLSC54.2 1055 bp Rape

Four o’clock

Arabidopsis

Rape

Tested in

BnSAG12.2 3.11 kb

Origin

Arabidopsis Arabidopsis, tobacco, tomato, lettuce, creeping bentgrass, petunia, cassava, wheat, rice, gerbera, pelargonium

Size

AtSAG12

Promoter

Table 5 Promoters for developmentally regulated expression

Mid

Strong

Ripening fruit

Leaf

Senescent leaf

Leaf

Leaf

Leaf

Leaf

Leaf

Strong

Weak

Strong

Flower, petal Mid

Leaf, stigma, Strong root

Leaf, flower

Leaf

Leaf, stem, and cotyledon aging

Immature and ripe fruit; fruit and leaf abscission zone

Stem, root, sepal

Vascular tissues (stem, root), flower, anther, stigma, fruit, seed

Green silique

Not in seeds

Main organ/ tissue Activity Other tissues

At:BFN1

At:SAG12

EcDXS Ca:FIB

IPT

IPT, kn1

Target gene TGP ID

96 Olga G. Smirnova and Alex V. Kochetov

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and senescent leaves, which confirmed the senescence-enhanced expression of the BnLSC54–2 promoter [127]. In senescence leaves, extensive blue GUS staining was detected in the mesophyll cells. The BnLSC54–2 promoter expression levels were low in green siliques, somewhat higher in flowers and were extremely high in mature Arabidopsis roots. Strong GUS activity was also detected at the stigma extending to cells around the vascular tissue in the style. Little GUS activity was detected in other tissues. A gene encoding a putative E3 ubiquitin ligase XB3 from garden four o’clock (Mirabilis jalapa) is expressed in roots, stems and senescence flowers. In petals, MjXB3 expression is increased >40,000-fold during the onset of visible senescence. The MjXB3 promoter (
914 from TSS) did not drive obvious GUS expression in roots, leaves and fresh petals except the senescing petals in petunia cv. Mitchell Diploid in a transient expression assay [125]. In petals of petunia and carnation (Dianthus caryophyllus cv. Imperial White), the MjXB3 promoter activity was lower than that of the AtSAG12 promoter. The MjXB3 promoter did not drive gene expression in the senescing corollas of the ethyleneindependent daylily (Hemerocallis  hybrida cv. Stella d’Oro) and daffodil (Narcissus pseudonarcissus cv. King Alfred), nor in the senescing petals of the ethylene-dependent Dendrobium orchid cv. Emma White, although CaMV35S and AtSAG12 promoters did. It seems that although the MjXB3 promoter is effective in heterologous dicotyledons, it is not active in monocotyledonous species. Since AtSAG12 did drive some GUS expression in fresh petunia and carnation corollas, it seems that the AtSAG12 promoter is not as senescence-specific in flowers as the MjXB3 promoter [125]. Stay green (SGR) gene of rice is involved in activation of chlorophyll degradation and its expression is induced along with senescence [128]. An upstream region of OsSGR (2379 bp from TIS) is able to direct senescence-specific GUS expression in rice leaves. Transgenic rice plants that express exoglucanase EXG1 gene under the control of the OsSGR promoter showed enhanced saccharification efficiencies of cellulosic biomass without affecting plant growth [129]. These results indicate that expression of cell wall degrading enzymes by a senescence-inducible promoter is one of the ways to enhance the saccharification ability of cellulosic biomass for efficient production of biofuels. A ZmSEE1 promoter (1.98 kb upstream from TIS) of maize cysteine protease gene showed enhanced expression during senescence in ryegrass (Lolium multiflorum) and maize. IPT gene expression under the control of the ZmSEE1 promoter displayed a delayed-senescence phenotype in leaves of ryegrass and maize [130, 131]. This study demonstrates that the delayed-senescence trait can be engineered into a monocot crop using a homologous senescence-enhanced promoter.

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A senescence-associated type I nuclease BFN1 is involved in senescence and programmed cell death associated with different development processes in Arabidopsis. A fragment of the 50 promoter sequence of AtBFN1 (
2357 to
52 from TIS) is able to drive GUS expression in transgenic Arabidopsis and tomato [132]. In both, the AtBFN1 promoter is activated in senescence leaves (up to eightfold), differentiating xylem and the abscission zone of flowers. In tomato, specific expression was observed in the leaf and the fruit abscission zones. The AtBFN1 promoter was also active in other tissues, including developing anthers and seeds, and in floral organs after fertilization (TGP At:BFN1). Oxygen free radicals play an essential role in senescence. The activities of different isoforms of the hydrogen peroxide (H2O2)scavenging enzyme catalase (CAT) were analyzed during senescence of Arabidopsis. There is a correlation between AtCAT2 (1400 bp upstream of TIS) and AtCAT3 (1850 bp upstream of TIS) promoter activities and the corresponding enzyme activities in all tissues. The AtCAT2 promoter provides a decrease in GUS staining from very intense in 4-week-old rosette leaves to very low in 8-week-old rosette leaves. Respectively, CAT2 activity decreased and H2O2 content increased with bolting time. The age-dependent increase in the AtCAT3 promoter activity is clearly shown in seedlings and in a leaf of a 7-week-old plant. Thus, the AtCAT2 promoter is downregulated and the AtCAT3 promoter is upregulated during leaf aging in Arabidopsis [108]. A promoter of a peach 1-aminocyclopropane-1-carboxylic acid oxidase gene (PpACO2) is able to drive the expression in senescent leaf blade, not in young leaves. The PpACO2 promoter provided a low level of activity in fruit and leaf abscission zones, and in vascular bundles of immature and ripe fruit of transgenic tomatoes cv. Micro-Tom [133]. These data are in contrast with those obtained for peach ACO2 mRNA. In peach, PpACO2 mRNA was detected only in immature fruit, epicotyl and root of seedling. The discrepancy might be due to a lesser stability and translation of PpACO2 mRNA in comparison to that of chimeric one. To generate the PpACO2-GUS construct, a fragment of 2.1 kb, representing the PpACO2 promoter region, first exon, first intron, and 6 aa of the second exon was used. Transcription of senescence-associated receptor protein kinase gene from bean (PvSARK) is induced during late maturation and decreased during the development of senescence [134]. A PvSARK promoter (an 840-bp fragment) is induced toward tobacco leaf maturation and during drought treatment. A suppression of drought-induced leaf senescence in PvSARK-IPT tobacco plants resulted in outstanding drought tolerance as shown by vigorous growth after a long drought period that killed the control plants. The transgenic plants maintained high water contents, retained photosynthetic activity and displayed minimal yield loss during

Tissue-Specific and Developmentally-Regulated Plant Promoters

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the drought [135]. Expression of the IPT gene under the regulation of the PvSARK promoter helps improve productivity under water deficit conditions in rice, peanut, cotton, and maize [136– 139]. Fibrillin (FIB) is a plastid lipid-associated protein, which is accumulated in fibrillar-type chromoplasts in ripening pepper fruit, and in leaf chloroplasts under stress conditions. A CaFIB promoter (2296 bp from TIS) was strongly upregulated during tomato fruit ripening in a manner similar to the induction of this gene in pepper fruits. Induction occurred at the mature green stage preceding ripening. In addition to stress-related induction, a progressive increase in the CaFIB promoter activity is noticed during aging in various tomato photosynthetic tissues (TGP Ca:FIB) [32, 33, 140]. A progressive increase in promoter activity was observed during leaf development, reaching a c. eightfold increase in the oldest leaves with respect to young leaves. Similarly, higher CaFIB promoter activity levels were observed in older stems and cotyledons [32]. The CaFIB promoter is found to be active in various dicot species (tobacco, pepper, potato, tomato and Arabidopsis), but not in monocots (maize, barley) [32, 140]. The CaFIB promoter directed strong GUS expression in pepper leaves but, on average, signals obtained with the constitutive CaMV35S promoter were more intense than those with CaFIB [32]. Transgenic tomato plants containing Escherichia coli 1-deoxy-D-xylulose-5-phosphate synthase (DXS) gene, under the control of the CaFIB promoter targeted to the plastid with the tomato DXS transit peptide, resulted in an increased carotenoid content (1.6-fold). Carotenoid content was inherited in the next generation and exhibited the greatest increase of 2.2-fold [141].

3

Notes 1. The tissue-specific and developmentally regulated promoters presented here have a wide variety of activities and expression patterns, thus providing suitable applications in plant biotechnology. It is necessary to choose carefully the size of the promoter because different promoter deletion variants may have different tissue specificity. 2. A number of promoters provide the expression of transgenes in various plant species, in both monocotyledonous and dicotyledonous plants, pointing to a potential for a broad exploitation of these promoters for the tissue-specific expression of target genes. 3. In addition to the promoters described here, numerous other tissue-specific promoters have also been isolated (for leaves, trichomes, vascular bundle cells, stomata, etc.). TGP may be used as data source for their selection.

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root formation. Plant Physiol 165:1035–1046 84. Jeong JS, Kim YS, Baek KH, Jung H, Ha SH, Do Choi Y, Kim M, Reuzeau C, Kim JK (2010) Root-specific expression of OsNAC10 improves drought tolerance and grain yield in rice under field drought conditions. Plant Physiol 153:185–197 85. Redillas MC, Jeong JS, Kim YS, Jung H, Bang SW, Choi YD, Ha SH, Reuzeau C, Kim JK (2012) The overexpression of OsNAC9 alters the root architecture of rice plants enhancing drought resistance and grain yield under field conditions. Plant Biotechnol J 10:792–805 86. Jeong JS, Kim YS, Redillas MC, Jang G, Jung H, Bang SW, Choi YD, Ha SH, Reuzeau C, Kim JK (2013) OsNAC5 overexpression enlarges root diameter in rice plants leading to enhanced drought tolerance and increased grain yield in the field. Plant Biotechnol J 11:101–114 87. Chung PJ, Jung H, Choi YD, Kim JK (2018) Genome-wide analyses of direct target genes of four rice NAC-domain transcription factors involved in drought tolerance. BMC Genomics 19:40. https://doi.org/10.1186/ s12864-017-4367-1 88. Deng F, Yamaji N, Ma JF, Lee SK, Jeon JS, Martinoia E, Lee Y, Song WY (2018) Engineering rice with lower grain arsenic. Plant Biotechnol J 16:1691–1699 89. Yamamoto YT, Taylor CG, Acedo GN, Cheng CL, Conkling MA (1991) Characterization of cis-acting sequences regulating root-specific gene expression in tobacco. Plant Cell 3:371–382 90. Barone P, Rosellini D, Lafayette P, Bouton J, Veronesi F, Parrott W (2008) Bacterial citrate synthase expression and soil aluminum tolerance in transgenic alfalfa. Plant Cell Rep 27:893–901 91. Chan YL, Prasad V, Sanjaya CKH, Liu PC, Chan MT, Cheng CP (2005) Transgenic tomato plants expressing an Arabidopsis thionin (Thi2.1) driven by fruit-inactive promoter battle against phytopathogenic attack. Planta 221:386–393 92. Radi A, Dina P, Guy A (2006) Expression of sarcotoxin IA gene via a root-specific tob promoter enhanced host resistance against parasitic weeds in tomato plants. Plant Cell Rep 25:297–303 93. Vaughan SP, James DJ, Lindsey K, Massiah AJ (2006) Characterization of FaRB7, a near root-specific gene from strawberry (Fragaria  ananassa Duch.) and promoter

activity analysis in homologous and heterologous hosts. J Exp Bot 57:3901–3910 94. Liu JJ, Ekramoddoullah AK (2003) Rootspecific expression of a western white pine PR10 gene is mediated by different promoter regions in transgenic tobacco. Plant Mol Biol 52:103–120 95. Xu X, Guo S, Chen K, Song H, Liu J, Guo L, Qian Q, Wang H (2010) A 796 bp PsPR10 gene promoter fragment increased rootspecific expression of the GUS reporter gene under the abiotic stresses and signal molecules in tobacco. Biotechnol Lett 32:1533–1539 96. Yu ZH, Han YN, Xiao XG (2015) A PPO promoter from betalain-producing red Swiss chard, directs petiole- and root-preferential expression of foreign gene in anthocyaninsproducing plants. Int J Mol Sci 16:27032–27043 97. Chen L, Jiang B, Wu C, Sun S, Hou W, Han T (2014) GmPRP2 promoter drives rootpreferential expression in transgenic Arabidopsis and soybean hairy roots. BMC Plant Biol 14:245 98. Suzuki K, Yun DJ, Chen XY, Yamada Y, Hashimoto T (1999) An Atropa belladonna hyoscyamine 6beta-hydroxylase gene is differentially expressed in the root pericycle and anthers. Plant Mol Biol 40:141–152 99. Suzuki K, Yamada Y, Hashimoto T (1999) Expression of Atropa belladonna putrescine N-methyltransferase gene in root pericycle. Plant Cell Physiol 40:289–297 100. Nakajima K, Oshita Y, Kaya M, Yamada Y, Hashimoto T (1999) Structures and expression patterns of two tropinone reductase genes from Hyoscyamus niger. Biosci Biotechnol Biochem 63:1756–1764 101. Suzuki H, Fowler TJ, Tierney ML (1993) Deletion analysis and localization of SbPRP1, a soybean cell wall protein gene, in roots of transgenic tobacco and cowpea. Plant Mol Biol 21:109–119 102. Koehorst-van Putten HJ, Wolters AM, Pereira-Bertram IM, van den Berg HH, van der Krol AR, Visser RG (2012) Cloning and characterization of a tuberous root-specific promoter from cassava (Manihot esculenta Crantz). Planta 236:1955–1965 103. Salehuzzaman SNIM, Jacobsen E, Visser RGF (1993) Isolation and characterization of a cDNA encoding granule-bound starch synthase in cassava (Manihot esculenta Crantz) and its antisense expression in potato. Plant Mol Biol 23:947–962 104. Jones MO, Manning K, Andrews J, Wright C, Taylor IB, Thompson AJ (2008) The

Tissue-Specific and Developmentally-Regulated Plant Promoters promoter from SlREO, a highly-expressed, root-specific Solanum lycopersicum gene, directs expression to cortex of mature roots. Funct Plant Biol 35:1224–1233 105. Onyango SO, Roderick H, Tripathi JN, Collins R, Atkinson HJ, Oduor RO, Tripathi L (2016) The ZmRCP-1 promoter of maize provides root tip specific expression of transgenes in plantain. J Biol Res (Thessalon) 23:4 106. Kakrana A, Kumar A, Satheesh V, Abdin MZ, Subramaniam K, Bhattacharya RC, Srinivasan R, Sirohi A, Jain PK (2017) Identification, validation and utilization of novel nematode-responsive root-specific promoters in Arabidopsis for inducing host-delivered RNAi mediated root-knot nematode resistance. Front Plant Sci 8:2049 107. Park SH, Jeong JS, Han EH, Redilas MCFR, Bang SW, Jung H, Kim YS, Kim J-K (2013) Characterization of the root-predominant gene promoter HPX1 in transgenic rice plants. Plant Biotechnol Rep 7:339–344 108. Noh YS, Amasino RM (1999) Identification of a promoter region responsible for the senescence-specific expression of SAG12. Plant Mol Biol 41:181–194 109. Zimmermann P, Heinlein C, Orendi G, Zentgraf U (2006) Senescence-specific regulation of catalases in Arabidopsis thaliana (L.) Heynh. Plant Cell Environ 29:1049–1060 110. Huynh le N, Vantoai T, Streeter J, Banowetz G (2005) Regulation of flooding tolerance of SAG12:ipt Arabidopsis plants by cytokinin. J Exp Bot 56:1397–1407 111. Gan S, Amasino RM (1995) Inhibition of leaf senescence by autoregulated production of cytokinin. Science 270:1986–1988 112. Ori N, Juarez MT, Jackson D, Yamaguchi J, Banowetz GM, Hake S (1999) Leaf senescence is delayed in tobacco plants expressing the maize homeobox gene knotted1 under the control of a senescence-activated promoter. Plant Cell 11:1073–1080 113. Jordi W, Schapendonk A, Davelaar E, Stoopen GM, Pot CS, De Visser R, Van Rhijn JA, Gan S, Amasino RM (2000) Increased cytokinin levels in transgenic PSAG12-IPT tobacco plants have large direct and indirect effects on leaf senescence, photosynthesis and N partitioning. Plant Cell Environ 23:279–289 114. Cowan AK, Freeman M, Bjorkman PO, Nicander B, Sitbon F, Tillberg E (2005) Effects of senescence-induced alteration in cytokinin metabolism on source-sink relationships and ontogenic and stress-induced transitions in tobacco. Planta 221:801–814

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115. Fu Y, Ding Y, Liu X, Sun C, Cao S, Wang D, He S, Wang X, Li L, Tian W (1998) Rice transformation with a senescence-inhibition chimeric gene. Chin Sci Bull 43:1810–1815 116. McCabe MS, Garratt LC, Schepers F, Jordi WJ, Stoopen GM, Davelaar E, van Rhijn JH, Power JB, Davey MR (2001) Effects of P (SAG12)-IPT gene expression on development and senescence in transgenic lettuce. Plant Physiol 127:505–516 117. Chang H, Jones ML, Banowetz GM, Clark DG (2003) Overproduction of cytokinins in petunia flowers transformed with P(SAG12)IPT delays corolla senescence and decreases sensitivity to ethylene. Plant Physiol 132:2174–2183 118. Swartzberg D, Kirshner B, Rav-David D, Elad Y, Granot D (2008) Botrytis cinerea induces senescence and is inhibited by autoregulated expression of the IPT gene. Eur J Plant Pathol 120:289–297 119. Swartzberg D, Hanael R, Granot D (2011) Relationship between hexokinase and cytokinin in the regulation of leaf senescence and seed germination. Plant Biol (Stuttg) 13:439–444 120. Sykorova B, Kuresova G, Daskalova S, Trckova´ M, Hoyerova´ K, Raimanova´ I, Motyka V, Tra´vnı´ckova´ A, Elliott MC, Kamı´nek M (2008) Senescence-induced ectopic expression of the A. tumefaciens ipt gene in wheat delays leaf senescence, increases cytokinin content, nitrate influx, and nitrate reductase activity, but does not affect grain yield. J Exp Bot 59:377–387 121. Xu Y, Gianfagna T, Huang B (2010) Proteomic changes associated with expression of a gene (ipt) controlling cytokinin synthesis for improving heat tolerance in a perennial grass species. J Exp Bot 61:3273–3289 122. Merewitz EB, Gianfagna T, Huang B (2011) Photosynthesis, water use, and root viability under water stress as affected by expression of SAG12-ipt controlling cytokinin synthesis in Agrostis stolonifera. J Exp Bot 62:383–395 123. Zhang P, Wang WQ, Zhang GL, Kaminek M, Dobrev P, Xu J, Gruissem W (2010) Senescence-inducible expression of isopentenyl transferase extends leaf life, increases drought stress resistance and alters cytokinin metabolism in cassava. J Integr Plant Biol 52:653–669 124. Lai QX, Bao ZY, Zhu ZJ, Qian QQ, Mao BZ (2007) Effects of osmotic stress on antioxidant enzymes activities in leaf discs of PSAG12-IPT modified Gerbera. J Zhejiang Univ Sci B 8:458–464

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125. Xu X, Jiang CZ, Donnelly L, Reid MS (2007) Functional analysis of a RING domain ankyrin repeat protein that is highly expressed during flower senescence. J Exp Bot 58:3623–3630 126. Noh YS, Amasino RM (1999) Regulation of developmental senescence is conserved between Arabidopsis and Brassica napus. Plant Mol Biol 41:195–206 127. Butt A, Mousley C, Morris K, Beynon J, Can C, Holub E, Greenberg JT, BuchananWollaston V (1998) Differential expression of a senescence-enhanced metallothionein gene in Arabidopsis in response to isolates of Peronospora parasitica and Pseudomonas syringae. Plant J 16:209–221 128. Sato Y, Morita R, Nishimura M, Yamaguchi H, Kusaba M (2007) Mendel’s green cotyledon gene encodes a positive regulator of the chlorophyll-degrading pathway. Proc Natl Acad Sci U S A 104:14169–14174 129. Furukawa K, Ichikawa S, Nigorikawa M, Sonoki T, Ito Y (2014) Enhanced production of reducing sugars from transgenic rice expressing exo-glucanase under the control of a senescence-inducible promoter. Transgenic Res 23:531–537 130. Li Q, Robson PR, Bettany AJ, Donnison IS, Thomas H, Scott IM (2004) Modification of senescence in ryegrass transformed with IPT under the control of a monocot senescenceenhanced promoter. Plant Cell Rep 22:816–821 131. Robson PR, Donnison IS, Wang K, Frame B, Pegg SE, Thomas A, Thomas H (2004) Leaf senescence is delayed in maize expressing the Agrobacterium IPT gene under the control of a novel maize senescence-enhanced promoter. Plant Biotechnol J 2:101–112 132. Farage-Barhom S, Burd S, Sonego L, PerlTreves R, Lers A (2008) Expression analysis of the BFN1 nuclease gene promoter during senescence, abscission, and programmed cell death-related processes. J Exp Bot 59:3247–3258 133. Rasori A, Bertolasi B, Furini A, Bonghia C, Tonuttia P, Ramina A (2003) Functional analysis of peach ACC oxidase promoters in

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Chapter 5 Choice of Explant for Plant Genetic Transformation Nibedita Chakraborty, Priyanka Chakraborty, Moutushi Sen, and Rajib Bandopadhyay Abstract Particle bombardment or biolistic transformation is an efficient, versatile method. This method does not need any vector for the gene transfer and is not dependent on the cell type, species, and genotype. The success of any transformation technique depends on the starting experimental materials or the explants. Here, we describe the factors that have influenced the choice of explants in biolistic transformation. Many general factors in the selection of explants in the development of transgenic plants are presented here. Therefore, this chapter provides extensive guidelines regarding the choice of explants for researchers working on various plant genetic transformation techniques. Key words Explants, Particle bombardment, Plant genetic transformation, Transgenic plants, Tissue culture

1

Introduction A tissue culture system is required for successful genetic transformation which gives rise to regenerated whole plants from single cells. Genetic transformation is a method to transfer the gene of interest to those cells that have the regeneration capacity. The transformation and regeneration are highly influenced by the explants type, such as hypocotyls, cotyledons, leaves, stems, and roots all used as starting materials in the process of transformation. The following sections discuss with the selection of explants and the different factors influencing the choice of explants.

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Plant Genetic Transformation

2.1 General Strategies for Transgene Delivery

Plant genetic transformation is a scientific approach of transferring DNA from an organism into the genome of another organism of interest. The foreign gene incorporates into the host plant genome and inherits through generations after generations. The transferred

Sachin Rustgi and Hong Luo (eds.), Biolistic DNA Delivery in Plants: Methods and Protocols, Methods in Molecular Biology, vol. 2124, https://doi.org/10.1007/978-1-0716-0356-7_5, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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gene is called transgene and the resulting plant is known as transgenic plant. Once the explant has been transformed, the plant cells containing the transgene are cultured in the nutrient medium under controlled condition and regenerated back into whole plants. This can happen because of totipotent nature of plant cells. Therefore, the gene of interest is present in every single plant cell. Several methods for exogenous gene introduction into a plant genome have been described in literature [1, 2]. In plants, the methods of gene transfer are classified into two groups; (a) vector-mediated gene transfer and (b) direct or vectorless gene transfer. The commonly used methods of gene transfer are represented in the flowchart (Fig. 1). Vector-mediated gene transfer is performed either by Agrobacterium-mediated transformation or by using plant viruses as vectors. Agrobacterium is a natural living soil bacterium which is capable of infecting a wide range of plant species and causing crown gall diseases. It has natural transformation abilities. When A. tumefaciens infects a plant cell naturally through wound or other ways, it transfers a copy of its T-DNA, which is a small section of DNA carried on its Ti (Tumor inducing) plasmid. The term direct or vectorless gene transfer is used when a naked DNA of interest is transferred to the plant genome. The direct gene transfer methods can be categorized into three groups: (a) physical methods including electroporation, particle bombardment, microinjection, liposome encapsulation, and silicon carbide fibermediated gene transfer. (b) Chemical gene transfer method

Plant Transformation methods

Physical

Chemical

DNA imbibition

Biological

1. Polyethylene glycol

1. Agrobacterium

2. Biolistic or Prticle

(PEG)-mediated

tumefaciens

bombardment

2. Diethyl amino ethyl

2. A. rhizogenes

(DEAE) dextran-

3. plant viruses

1. Electroporation

3. Microinjection

4. Liposome encapsulation 5. Silicon carbide fibres

mediated 3. Calcium phosphate precipitation

Vector- or virusmediated gene transfer

Direct or vector less gene transfer

Fig. 1 Schematic representation of the commonly used gene transfer methods in plants

Explants for Transformation

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Plant Excised explants (leaf, mature or immature embryo, callus, meristem) Physical or chemical or biological method

Transgene delivery Transformed cells/tissues Shoot Embryo culture Callus culture regeneration Shoot induction

Regeneration and selection

Transgenic plants PCR or Southern blotting

Screening of transgenic plants

Fig. 2 A general overview of the development of transgenic plants

containing polyethylene glycol (PEG)-mediated, diethyl amino ethyl (DEAE) dextran-mediated, calcium phosphate precipitation and (c) Imbibition of DNA by cells, tissues or organs. Majority of the direct gene transfer methods are very simple and efficient and are applied in the development of several transgenic plants. An outline of general schematic representation of the development of transgenic plants is provided in Fig. 2. The following section is focused on some details of the particle bombardment-mediated physical gene transfer method. 2.2 Biolistics or Particle Bombardment Method

One of the most effective direct gene transfer methods is biolistics or particle bombardment method. This is the physical method since it does not use any bacteria for the transformation process and become versatile and effective for the transfer of DNA in mammalian cells, microorganisms and also in monocot plants [3]. This method was first described in plants by John Sanford at Cornell University [4]. In this method, the recombinant DNA molecules are coated with microscopic tungsten or gold particles (microcarriers) known as microprojectiles which are accelerated by macrocarriers (macro projectiles). These macrocarriers are inserted into the firing chamber of the gene gun and an explosive force fires or compressed gas is used to drive the macrocarriers. At the end of the gene gun there is a small stopping plate which does not permit the macrocarriers but allow the microcarriers coated with DNA to pass through at a high speed to the cell wall into the cytoplasm of the target cells. At this point, DNA dissociates from the microcarriers and integrates into the plant genome.

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The biolistic transformation occurs in two stages: At the preintegration stage, DNA enters into the cells, and at a later stage it integrates into the plant genome. DNA delivery in particle bombardment method is very efficient compared to DNA integration. After DNA transfer, a small proportion of the cells receive DNA and actually become stably transformed, while in the rest of the cells the DNA enters and may be expressed for a short period which is termed as transient expression. Here, the DNA is not integrated into the plant genome and is eventually degraded by plant nucleases [5]. Transient expression occurs immediately after DNA/gene transfer and after particle bombardment the transfer gene expressed with a reporter gene, such as gusA or gfp, which is used for the identification and comparison of the protein construct and the protein’s appropriate activity. The success of efficient particle bombardment method involves the selection of potential explants with distinct morphogenic properties. The explants can be different parts of a plant (e.g., leaves, shoots, stems, roots, flowers, petals, pollen, endosperm, callus) and various types of immature or mature cells. The explants should be totipotent in nature which has the ability to dedifferentiate into a whole plant. In plant tissue culture, explants are aseptically cut and placed in a nutrient culture medium, where they start to grow and continue their function under sterile conditions. The types of explants in plant genetic transformations vary from species to species.

3

Different Types of Explants Used in Plant Genetic Transformation A successful transformation is generally influenced by the correct choice of explants used in plant tissue culture. The choice of explants depends on (1) the type of culture, (2) the purpose of the proposed culture, and (2) the plant species from which the explants need to be used. A brief introduction of different types of explants used in plant tissue culture is provided below: Cotyledon: Cotyledon is the first embryonic leaves of seedlings after seed germination. The classification of the flowering plants is determined by the number of cotyledons. Plant species with one embryonic leaf or cotyledon are termed monocotyledonous and those with two embryonic leaves are termed dicotyledonous. Maize is one of the most commonly used monocotyledonous grains for transformation technology with particle bombardment method [6]. Cotyledon culture needs to be performed with aseptically grown seedlings for efficient shoot regeneration. The excision of cotyledon from embryo axes depends on the age of the cotyledon used for efficient regeneration studies.

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Leaf: In leaf culture, excised young leaf primordia or young immature leaves are used as explants. The explants can be collected from plant species aseptically grown for tissue culture. The excision of the leaf explants depends upon the maturity stage to maintain the growth potential in the tissue culture medium. Shoot tip/meristem: Shoot tip/meristem culture is the culture of terminal (0.1–1.0 mm) portion of a shoot including the meristem (0.05–0.1 mm) along with primordial and emerging leaves and adjoining stem tissues. Meristem culture is the in vitro culture of the shiny special dome-like structure less than 0.1 mm in length excised from the shoot apex. Once the shoots grow directly from the excised explant materials like shoot tip/meristem, they can be used for nodal propagation by separating shoot-containing nodes into small segments and culturing them on a nutrient medium. Flowers: The culture of excised aseptic floral buds on a nutrient medium to produce full flowers and subsequently mature seeds is termed flower culture. In flower culture, flowers can be excised at different developmental stages, for example, primordial stage, preor postpollination stage, or bud stage. Flower primordium or flower bud culture is mainly used for studying the fundamentals of floral development, and flower culture at postpollination stage is applied in studies of fruit development. At the prepollination stages of a flower, the fruit development usually does not occur; however, parthenocarpic fruits can be produced by supplementing the nutrient medium with auxin. Roots: In the case of root culture, radicle root tips are excised from aseptically grown germinated seeds and cultured on the nutrient medium under optimum conditions. A replica of excised roots can also be cultured by cutting and rooting the main root tips or lateral root tips and subculturing them on a fresh medium under controlled conditions. Biolistic transformations have been widely used for a wide range of cell and tissue explants which are capable of producing healthy and regenerable cells through nuclear division [5]. These explants mainly include embryos collected from seeds, shoot apices, microspores, and immature embryos. One of the main drawbacks of using immature embryo is the unavailability of explant materials in suitable stages of development [6]. A list of transgenic plants with their source of explants used in particle bombardment method is given in Table 1. Several different aspects related to the choice of explants used in plant genetic transformation and regeneration are discussed in this chapter. Although this chapter mainly emphasizes on the choice of explants for particle bombardment method, many factors discussed also influence the choice of explants for Agrobacteriummediated transformation. Therefore, the chapter is equally useful to researchers working on either of the two methods, providing them with new experimental guidelines and platforms for plant genetic transformation.

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Table 1 A list of transgenic plants with their explants used in particle bombardment method Plants

Source of the explants

References

Alfalfa

Embryonic callus

[7]

Banana

Embryogenic suspension cells, banana buds

[8, 9]

Barley

Microspore, immature embryo,

[10, 11]

Cotton

Meristems of embryonic axes

[12]

Maize

Immature embryo

[13]

Mulberry

Hypocotyl and cotyledon explants

[14]

Peanut

Embryo axes

[15]

Potato

Nonembryogenic tissues (nodes, leaves, and microtubers)

[16, 17]

Rice

Embryonic callus, immature embryos

[18–21]

Sorghum

Immature embryo

[22]

Soybean

Meristematic regions from either mature or immature embryo axes

[23]

Tobacco

Suspension cell

[24]

Tomato

In vitro germinated seedlings

[25]

Watermelon

Cotyledon

[26]

Wheat

Immature caryopses, scutella from immature embryo, callus

[27, 28]

4 4.1

Factors Influencing the Choice of Explants Explant Age

The age of explants is an important factor in plant tissue culture. The transformation and regeneration efficiencies are greatly controlled by the age of explants [29]. The explants collected at a controlled developmental stage from a healthy donor plants developed under optimal condition would reproduce an efficient transformation with phenotypically healthy, fertile, transgenic plants. Pastori et al. [27] showed the significant correlation (r2 ¼ 0.946) between the transformation efficiency and the age of the donor plants in the wheat varieties. In this study, the best bombardment achieved higher transformation efficiency in both of the wheat varieties (Cadenza and Canon) when the embryos collected from approximately 70-day-old plants in comparison to 80- to 82-day-old plants [27]. Another study also reported that the effect of explant age on successful biolistic transformation in watermelon is significant [26]. The cotyledon explants obtained from 4-day-old seedlings of watermelon showed higher percentage (98%) of transformation in comparison to 5-day-old seedlings (90%). In case of Agrobacterium-mediated transformation, leaf explants that were collected from 30-day-old Populus sp. showed higher

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transformation efficiency than those collected from sixty- and ninety-day-old plants [30]. This is because Agrobacteriummediated transformation efficiency depends on the surface of the leaf explants, since surface of younger leaves have more binding sites that can be easily attached by Agrobacterium [31]. In contrast, mature leaves contain intact epidermis cells at their leaf surface having limited number of attachment sites and resulting in a decrease in transformation efficiency [32]. Similarly, leaf explants from younger seedlings (three weeks old) of Lycium barbarum showed higher transformation efficiency than those from 4- or 5-week-old seedlings [33]. An alternative clarification by Villemont et al. [34] states that a few number of cells divide in mature seedlings, resulting in reduction in T-DNA integration in host cells, thereby causing a decrease in transformation and shoot regeneration efficiency [34]. Similar phenomena have been evidenced in several plant species (e.g., Petunia [34], Citrus [35–37] and Poncirus [38]). Likewise, the explants from shoot tip showed higher transformation efficiency when collected at a young age. Apical tissues collected from 4 to 6 days germinated Zea mays seedlings showed higher susceptibility (90–100%) toward Agrobacterium culture than older seedlings [39]. Similar results have been reported in brinjal (Solanum melongena) where shoot tip from 7-day-old seedlings showed higher transformation efficiency (19.8%) than 14- and 21-day-old seedlings (15.7% and 10.4%, respectively) [40]. The higher transformation efficiency in the younger shoot tip has been attributed to the high rate of cell division in young explants which helps in the uptake of T-DNA at a higher rate [40]. In a similar way, explants with juvenile and actively dividing cells are commonly the best target tissue for a successful bombardment transformation [41]. The shoot apices collected from 3-, 5-, and 9-day-old cotton (Gossypium spp.) seedlings showed significant differences in cotton shoot elongation process [42]. The highest regeneration efficiency was evidenced in shoot apices excised from 5-day-old seedlings (76.29%) than 3- and 9-day-old cotton seedlings (54.29% and 68.93%, respectively) [42]. Similar results have also been shown in cotyledonary leaf explants in that the explants excised from fresh germinating seeds showed significantly higher transformation and regeneration efficiency than the explants collected from 1- and 2-week-old leaves [43]. Nodal explants excised from 50- to 60-day-old in vitro seedlings (Withania somnifera) showed higher GUS expression along with higher regeneration efficiency [44]. The regeneration efficiency has also been observed to depend on the age of petiole explants [45]. The significantly higher percentage of regeneration (65.78%) with higher number of regenerated shoot buds (6.76) were observed in the petioles isolated from second leaf of Jatropha curcas rather than first (20.82%), third (53.33%), fourth (30.07%), and fifth (12.31%) leaves, counted from the top of the shoots

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[45]. The young age-influenced regeneration efficiency has also been evidenced with root explants [46]. Among the root explants excised from three-, five-, seven-, and nine-day-old seedlings of Indica rice genotypes, the 3-day-old root explants showed higher regeneration efficiency than roots collected from older seedlings [46]. 4.2

Explant Size

4.3 Explant Excise Position

The size of explants is an essential factor for plant regeneration efficiency. In plant tissue culture, the large size of explants is more suitable than small size because smaller explants are too difficult to culture and large explants can reserve more nutrient and plant growth regulator to maintain their growth in the culture condition [47]. In maize, 1–2 mm size of the immature embryo has been reported as the optimum explants for embryogenic callus regeneration in biolistic transformation [48]. Similarly, in their analysis of the size of embryos, Pastori et al. [27] reported it should lie between 0.5 and 1.5 mm for their use as explants in biolistic method [27]. Stem segments of citrus ~1 cm in length were more efficient in shoot regeneration than shorter segments (0.2–0.5 cm). The cut ends from larger stem segments can regenerate shoots with little or no callus formation [35]. Likewise, longer size (1.5 cm) of hypocotyl explants of eggplant showed optimum response in shoot regeneration compared to short- (0.5 cm), medium-sized (1 cm) or very long-sized (2 cm) explants [49]. In rubber-producing dandelion species, regeneration and transformation efficiency was influenced by the size of the root, where roots greater than 1 mm, but not those 60 m2/cm3) [55]. Regardless of definition, particularly for plant transformation applications requiring internalization into the cell through the cell wall and lipid membrane, nanomaterial carrier size is one of the most important factors that determine whether particles internalize passively or will require assistance from biolistics. Nanoparticles are increasingly used as delivery vehicles for applications in plant science; therefore, it is essential to understand plant-nanomaterial interactions. In particular, it is important to understand how nanomaterials interact with plants, if there could be adverse or toxic effects, or if nanomaterials could affect endogenous plant biology. Several research groups have begun studying these plant–nanomaterial interactions [7, 56–59]. For instance, an early study revealed that silica nanoparticles do not adversely affect the wheat seed germination, emergence, or growth of seedlings

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[60]. Another follow-up study determined the effects of functionalized and non-functionalized SWCNTs on root elongation of six different crop species: cabbage, carrot, cucumber, lettuce, onion, and tomato. Results showed that both types of CNTs enhanced root elongation in onion and cucumber, cabbage and carrot were not affected by either form of nanotubes, root elongation in lettuce was inhibited with functionalized CNTs, and tomato was found to be most sensitive for CNTs with significant root length reduction [61]. It was also demonstrated by another study that SWCNTs enhanced germination of rice and zucchini seeds and did not show adverse effects on root elongation [62]. A few studies reported the uptake, translocation, and specific localization of magnetic iron oxide nanoparticles in pumpkin plants, with no observable toxicity on plant growth [63]. Despite progress in assessing plant-nanomaterial interactions, it is worthwhile to note that interactions will be specific to plant species and tissue type, plant age, nanoparticle type, nanoparticle surface chemistry, and other parameters, and that more research in this multiparameter space will be necessary to fully understand nanomaterial interactions with plants. Nanomaterial-based delivery vehicles show promise to bring unique advantages to plant genetic transformations given the success in animal studies over the past few decades. Nanomaterials can provide controlled, target-specific, and stimuli-responsive release of genetic cargoes into plant cells and can also be targeted to subcellular locations such as plastids. In this chapter, we discuss the synthesis and characterization of the few instances of nanoparticles used for nanobiolistics, information gained from animal nanobiolistic studies, and compare nanobiolistic to microbiolistic delivery.

2

Discussion

2.1 Synthesis and Characterization of Nanoparticles for Nanobiolistics

The most commonly used nanoparticles for nanobiolistic delivery are mesoporous silica nanoparticles (MSNs) and gold nanoparticles (AuNPs). MSNs contain a highly porous structure that permits internal loading of biomolecules—such as DNA, RNA, and proteins—and subsequent biolistic delivery to animal and plant tissues. AuNPs are chemically identical to the standard 0.6 μm gold projectile traditionally used in biolistic delivery; however, their smaller size offers several advantages. The remainder of this section will focus on the synthesis and characterization of these two common nanoparticles used in plant biolistics, and examples from the literature of their successful application in animal and plant transformations.

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2.1.1 Synthesis of Mesoporous Silica Nanoparticles

MSN synthesis proceeds by a modified Sto¨ber process, which relies on alkyl silicate polymerization in the presence of an ionic surfactant to produce porous spherical particles [64]. The most commonly used alkyl silicate precursor and surfactant are tetraethyl orthosilicate (TEOS) and cetyltrimethylammonium bromide (CTAB), respectively. Alkyl silicates undergo sequential hydrolysis in acidic conditions and condensation in basic conditions to produce 3-dimensional siloxane networks. Ionic surfactants form micelles with positively charged surfaces that associate with silicate molecules and provide a template around which the polymer network is formed. The surfactant is then removed by calcination, dialysis, or solvent extraction, resulting in a siloxane mesostructure with particle diameters on the order of tens to hundreds of nanometers and pores on the order of several nanometers. Precise control over particle diameter is achieved by tuning the reaction pH and precursor-surfactant ratio, whereas pore diameter can be tuned by changing the surfactant chain length, adding organic swelling agents, or by using block copolymer cotemplates [65]. Furthermore, functionalized MSNs can be synthesized by chemically modifying the alkyl silicate precursor prior to polymerization. For example, Slowing et al. synthesized fluorescent-labeled MSNs through the coupling of fluorescein isothiocyanate (FITC) to an aminosilane APTMS, which then underwent co-condensation with TEOS to form FITC-MSNs, allowing cellular internalization to be monitored by fluorescence microscopy [66]. Additionally, co-condensation with a mercaptosilane generates thiolated MSNs [67] that are useful for downstream functionalization such as bioconjugation or pore-capping for improved delivery efficiency. For example, Lai et al. synthesized pore-capped MSNs through disulfide linkage with cadmium sulfide (CdS) nanoparticles, allowing for controlled release of small molecules upon the introduction of a disulfide-reducing agent [68].

2.1.2 Synthesis of Gold Nanoparticles

AuNP synthesis typically proceeds in situ by Au(III) reduction from chloroauric acid (HAuCl4) precursor in the presence of a stabilizing agent to form colloidal Au. The major synthetic routes for spherical AuNPs are the Turkevich–Frens method [69], where surfaceadsorbed citrate acts as a stabilizer, and the Brust–Schriffin method [70], where covalently bonded thiols act as a stabilizer. Many advancements have been made to these standard protocols, allowing control over particle diameter through tuning of the Au–stabilizer ratio or use of alternate reducing and stabilizing agents [71]. The Brust–Schriffin synthesis is preferred for downstream bioconjugation applications as thiolated surface groups are amenable to a wide variety of biocompatible linker chemistries. Lastly, cylindrical gold nanorods may be synthesized electrochemically by deposition onto polycarbonate or alumina pore templates or in the presence of rod-inducing surfactants [72].

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2.1.3 Nanoparticle Characterization

Many techniques have been developed to measure critical parameters of MSNs and AuNPs such as size, morphology, dispersity, colloidal stability, and porosity. Zeta potential (ZP), or the electric potential at the interfacial double layer, is a convenient and wellestablished indicator of colloidal stability that is simple and inexpensive to measure. Commonly accepted values for ZP are >30 mV and 80%, and hence this is a suitable system to adopt. This chapter will outline two important determinants, target tissue and biolistic variables, to achieve the highest TE of sugarcane cells. A range of target tissue including mature leaves, furled leaf sections, embryogenic callus, and single cell suspension has been used for biolistic experiments in sugarcane [15]. Putative embryos (also known as an embryogenic callus) are the most commonly reported tissue used for transformation in sugarcane. Depending on the variety, transformation can be achieved within 7–10 days of initiation using furled leaf sections [22] versus 2–3 months when using embryogenic callus [2]. While using furled leaf sections is attractive, some varieties are more recalcitrant and require embryogenic callus as the target tissue. A prerequisite to obtaining high TE irrespective of the target tissue is subculturing within a week of the planned bombardment. It is to ensure that cells are in an active state of division. Pre and post bombardment osmotic treatments are important irrespective of the type of target tissue. The tissue must be exposed to both pre and post bombardment treatments which involve placing the selected tissue on an osmotic medium that consists of Murashige and Skoog (MS) based medium [23] supplemented with osmotic reagents (0.2 M sorbitol and 0.2 M mannitol solutions) for several hours. This step induces plasmolysis of the cells, thus minimizing the risk of cell rupture upon bombardment [24]. Following the post bombardment osmotic treatment, the bombarded tissue is transferred to the dark on to MS growth medium containing the necessary growth regulators, but no selection agent (recovery phase). The recovery phase can vary from 3 to 7 days, to enable rapid multiplication of cells, before being transferred to selection medium containing the selection agent at the optimum concentration as per the dose–response growth curve, developed previously. Cells are left on selection in the dark for either another 3 weeks (embryogenic callus) or transferred to light (furled leaf). Production of plantlets is monitored, and selection pressure is maintained throughout the period in light. This selection scheme is to minimize the escapes of untransformed cells. Once plantlets are >3 cm, they can be tested for the presence of the transgenes, using standard DNA extraction and PCR analysis using transgene-specific primers.

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Materials Target Tissue

2.2 Stock Solutions and Media

1. In the Australian sugarcane varieties, embryogenic callus is initiated from transverse sections of immature furled leaf whorls and maintained on MS medium containing 1–3 mg/L of 2,4-Dichlorophenoxy acetic acid (2,4-D) depending on the variety (Fig. 2) (see Notes 1 and 2). It is a prerequisite that the production of embryogenic callus has been optimized in the variety that is being transformed. 1. MS medium: Prepare Murashige and Skoog medium using M5519 from Sigma, which contains the macro-, micronutrients, and vitamins of the original classic formulation. Use 4.4 g of M5519 for a liter of medium; add 30 g of sucrose and 1–3 mg of 2,4-D. Adjust pH (5.6–5.8) of the medium. Add agar directly into the 1 L Schott bottle (7 g/L), before adding the MS liquid medium and autoclave it at 121
C for 20 min. 2. 2,4-D: Make a stock solution of 2,4-D at 1 mg/mL. Weigh 40 mg of 2,4-D into a beaker and add 1 M KOH slowly, till the powder dissolves. Add distilled water to make the final volume to 40 mL. This solution can be passed through a 0.22 μm filter in the sterile hood, and stored in 1 mL aliquots in 1.5 mL Eppendorf tubes at 20
C. Alternatively, it can be stored in the refrigerator and added to the medium prior to autoclaving, as it is not heat-labile.

Fig. 2 Production of Embryogenic callus in sugarcane. (a) Cane top with leaves trimmed. (b) Transverse sections on callus induction medium. (c) Embryogenic callus production after 10–12 weeks

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2.3 DNA for Bombardment into Sugarcane Cells

221

1. Amount of DNA: The prep contains 50 ng to 10 μg which is enough for 6 shots (see Note 3). 2. Type of DNA: Circular or linearized plasmid (see Note 4). 3. The number of plasmids to be cobombarded: (1) Plasmids containing DNA to be introduced into cells can be either intact (circular) or linearized and carry the gene of interest, selectable marker gene (e.g., neomycin phosphotransferase or other) and/or reporter genes (see Note 5). (2) When cobombarded several plasmids (up to four) can be mixed before shooting. We recommend the use of the equimolar quantity of each plasmid.

2.4 Basic Equipment and Reagents for Bombardment

1. Microcarriers: (1) Particle type: Gold or tungsten (see Note 6). (2) Particle size: Gold—0.6, 1 or 1.6 μm; Tungsten—0.4, 0.7, 1.1, 1.3 or 1.7 μm. The choice of particles is subjective. Gold is more expensive, and the smallest size available is 0.6 μm. Tungsten, on the other hand, is cheaper and comes in a smaller size (0.4 μm), than the smallest gold particles. However, we obtained more consistent and reproducible results using gold particles in sugarcane. 2. Rupture disk (psi 450–2200): Rupture disks also come in a range of thicknesses and provides the choice of shooting pressure. The higher shooting pressures are preferred when transforming organelles, which requires penetration of the DNA carrying particles across two layers of membranes. 3. Sterile macrocarrier. 4. Stopping screens. 5. 0.1 M Spermidine solution: A 0.92 g/mL density spermidine solution can be purchased from Sigma-Aldrich. Add 15.8 μL of spermidine solution to sterile distilled water to make a total volume of 1 mL. Store at 20
C in 25 μL aliquots. 6. 2.5 M Calcium chloride: Prepare the solution using distilled water and filter sterilize using a 0.22 μm syringe filter. Perform this in the sterile hood. Store at 4
C in 55 μL aliquots (see Note 7). 7. Ethanol analytical grade.

3

Methods

3.1 Preparation of Target Tissue

1. On the day of bombardment, place embryogenic callus pieces in a circle (~2.5 cm diameter) (Fig. 3) on MS containing osmoticum medium and incubate for several hours (see Note 8).

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Fig. 3 A plate showing the positioning of embryogenic calli before particle bombardment

3.2 Preparation of Gold Particles

1. Weigh out 50 mg gold (0.6 μm) in microfuge tube. 2. Add 1 mL 100% ethanol. 3. Vortex thoroughly for 2–5 min. 4. Stand for 15 min. 5. Pellet gold by centrifugation at 16,500  g for 10 s. 6. Remove ethanol, wash particles three times in water (add 1 mL water, vortex, pellet, and repeat the process twice). 7. Resuspend particles in 1 mL sterile 50% glycerol. 8. Dispense 50 μL aliquots (enough for 6 bombardments) into 1.5 mL tubes and store at 20
C (for up to 3 months).

3.3 Microcarrier Preparation for Bombardment

This mix is sufficient for 6 bombardments (the experiment is performed under sterile conditions in laminar flow and during the whole procedure the tubes are kept on ice). 1. Vortex 50 μL gold aliquot thoroughly (homogenize to remove clumps). 2. Add 10 μL DNA adjusted to 1 μg/μL concentration and vortex. If cobombarding with the selectable marker, then add equimolar ratio of both constructs. 3. Add 50 μL 2.5 M CaCl2 to the mixture and vortex. 4. Add 20 μL 0.1 M spermidine (free base) to the mixture and vortex. It is important to add CaCl2 and spermidine while gold particles are still suspended—have these ready to be added in two pipettors. There should be no delay between CaCl2 and spermidine additions. 5. Let the DNA precipitate on gold particles for 5 min on ice.

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6. Pellet gold particles by centrifugation at 950  g for 10 s. 7. Remove supernatant. 8. Add 35 μL of 100% ethanol (analytical grade) and resuspend gold particles by vortexing. 9. Visually confirm dispersal of gold particles and immediately apply 5 μL of suspension to the center of macrocarrier and spread evenly. Allow it to dry. 3.4 Microprojectile Bombardment

1. Sterilize the hood and particle gun chamber with ethanol. 2. Sterilize macrocarrier holders, macrocarriers, rupture disks, stopping screens and macrocarrier seeding tools in 70% isopropanol (soak for 30 min and air dry). (Perform steps 1 and 2 an hour prior to bombardment). 3. Place macrocarrier into the macrocarrier holder and ensure proper loading using macrocarrier seeding tool. 4. Place rupture disk into the rupture disk retaining cap, screw it into the place and tighten with wrench. 5. Place stopping screen at the bottom of microcarrier launch assembly. 6. Place macrocarrier holder containing gold particles coated macrocarrier into the assembly, with gold particles coated side facing stopping screen, and secure by screwing. Place the microcarrier launch assembly into the top shelf of PDS-1000. 7. Place tissue for bombardment onto selected shelf (second shelf from bottom). 8. Close the door and turn on vacuum switch. 9. Ensure helium valve is adjusted to approximately 200 psi above the prescribed pressure holding capacity of the rupture disk. 10. When vacuum reaches 28 psi, flip switch to hold and hold down fire switch till the disk ruptures. 11. Release vacuum. 12. Remove bombarded rupture disk.

tissue,

spent

macrocarrier

and

13. Place tissue on to osmotic medium for a further 3 h. 3.5 Post Bombardment

1. Transfer calli back on to MS medium with 2,4-D for the recovery period (up to 1 week), with no selection pressure and in the dark (see Note 9). 2. Replace culture medium with the one containing selection agent, and continue to culture in the dark for another 3 weeks.

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Fig. 4 Stages of plant regeneration in light on selection media. (a) Little green leaves emerged after 4 weeks in the light. (b) Ten week old well-established plantlets. (c) Plant ready for transfer to the humid chamber in soil

3. Subculture onto fresh medium with selection agent, but no 2,4-D and transfer into light (3650 lux) (Fig. 4a). 4. Repeat step 3 every third week, until the plantlets are visible (see Note 10) (Fig. 4b). 3.6 Molecular Analysis of Regenerants

1. Test regenerating plants for presence of transgene when they are at least 3 cm tall (Fig. 4c). 2. Extract DNA using published methods or kits. We use Qiagen plant DNA kits when we have a few samples to test. In case of large number of samples, we use the rapid and simple method of Thomson and Dietzgen [25]. After extraction DNA is diluted fivefold and 1 μL is used as template in the 25 μL PCR reaction. 3. Design transgene-specific primers and test DNA for the presence of transgene. 4. Optimize PCR reactions and annealing temperature as well as the thermocycler conditions for each primer pair. 5. Transfer plants containing transgenes to large tissue culture jars and continue to grow them in light on MS medium with selection agent. 6. When plants have a well-established root system, trim leaves and roots and then transfer plants to soil and grow in the glasshouse/growth chamber initially in a humid chamber for about a week, and then open vents gradually over 3 weeks to harden off plants, and finally remove the cover (Fig. 5a). 7. Spray plants with water to keep moist while in the humid chamber (Fig. 5b). 8. Transfer hardened plants to pots in a physical containment glasshouse after plants show signs of good growth.

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Fig. 5 Transgenic plant culture and Southern blot analysis. (a) Regenerated plants within the humid chamber and in soil. (b) After the cover has been removed. (c) Southern blot analysis of DNA from plants transformed using biolistic approach. The bands appeared after the hybridization of the DIG (digoxigenin) labeled probe. Variation in the number of bands show a variable number of the transgene integrations

9. Extract DNA from whole young leaves for Southern blot analysis as per published protocols (Fig. 5c). 10. Identify plants showing presence of transgenes and determine copy number. Both quantitative real time PCR or Droplet Digital PCR can be used [26].

4

Notes 1. Optimization of culture conditions to obtain the most suitable tissue to bombard is critical. The researcher will need to first test various 2,4-D concentrations to induce and maintain embryogenesis in the sugarcane variety they want to transform. If unsuccessful, then the effect of adding other growth regulators in different combinations will require testing, as there is not one specific culture medium that will produce embryogenic calli in all varieties of sugarcane. 2. For studies on gene/construct functionality, use young newly emerged leaves. We have found that the tender leaves of tissue cultured plants are ideal for this type of study. Another source would be the youngest leaves of recently germinated setts. 3. Amount of DNA for shooting requires optimization for sugarcane variety as well as the tissue being transformed. In our experience, using linearized plasmid DNA and amounts as

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low as 50 ng/prep can lead to high TE. We have used even lower concentrations of DNA, but TE got reduced. 4. We have tested both circular whole plasmid vectors as well as linearized minimal DNA cassettes. The advantage of the latter is that only sequences carrying genes of interest and regulatory elements are introduced, without the inclusion of the bacterial vector backbone sequences carrying antibiotic resistance or bacterial site of the origin of replication. 5. We have performed dose–response curves for the antibiotics (hygromycin and neomycin) in Australian sugarcane varieties [2]. Both neomycin and hygromycin resistant marker genes were effective in the selection of transformed cells by killing them in a reproducible manner. The dose–response curves showed reduced growth and fresh weight in nontransformed susceptible cells over time, in both dark and light treatment. Thus, either of these resistance marker genes can be used for effective selection of transformed cells of sugarcane. 6. We have used both microcarriers, and found that gold particles give consistent TE. However, for organellar transformation, tungsten may be a more suitable choice as it has a greater adsorptive surface due to its shape being not perfectly round. This enables adsorption of more DNA on a smaller particle size. 7. We aliquot CaCl2 and spermidine solutions sufficient for one batch of shooting into PCR tubes, and store them at 20
C, till required, thus preventing consecutive cycles of thawing and freezing, as well as minimizing contamination. 8. It is important that the target tissue has been recently subcultured on to fresh medium within the past 3–4 days. This encourages production of new cells which are actively dividing and are highly embryogenic and show high TE (Fig. 2). 9. The medium used during the recovery period was the same as for embryogenic callus generation. 10. We subculture the bombarded tissue every 3 weeks as we have found that callus growth and plant regeneration is best when medium is changed every 3 weeks.

Acknowledgments The authors would like to acknowledge Sugar Research Australia for the financial support provided.

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References 1. Bower R, Birch RG (1992) Transgenic sugarcane plants via microprojectile bombardment. Plant J 2:409–416 2. Joyce P, Kuwahata M, Turner N, Lakshmanan P (2010) Selection system and co-cultivation medium are important determinants of agrobacterium-mediated transformation of sugarcane. Plant Cell Rep 29:173–183 3. Jackson MA, Anderson DJ, Birch RG (2013) Comparison of agrobacterium and particle bombardment using whole plasmid or minimal cassette for production of high-expressing, low-copy transgenic plants. Transgenic Res 22:143–151 4. Wu H, Saeed F, Vilarinho A, Zeng Q, Kannan B, Phipps T, McCuiston J, Wang W, Caffall K, Altpeter F (2015) Transgene integration complexity and expression stability following biolistic or agrobacterium-mediated transformation of sugarcane. In Vitro Cell Dev Biol Plant 51:603–611 5. Ramasamy M, Mora V, Damaj MB, Padilla CS, Ramos N, Rossi D, Solis-Gracia N, VargasBautista C, Irigoyen S, DaSilva JA, Mirkov TE, Mandadi KK (2018) A biolistic-based genetic transformation system applicable to a broad-range of sugarcane and energycane varieties. GM Crops Food 17:1–17 6. Basnayake S, Moyle R, Birch RG (2011) Embryogenic callus proliferation and regeneration conditions for genetic transformation of diverse sugarcane cultivars. Plant Cell Rep 30:439–448 7. Taparia Y, Gallo M, Altpeter F (2012) Comparison of direct and indirect embryogenesis protocols, biolistic gene transfer and selection parameters for efficient genetic transformation of sugarcane. Plant Cell Tissue Organ Cult 111:131–141 8. Finer JJ, Vain P, Jones MW, McMullen MD (1992) Development of the particle inflow gun for DNA delivery to plant cells. Plant Cell Rep 11:323–328 9. PDS-1000/He particle delivery system biorad brochure and training video. http://www.biorad.com/en-au/product/pds-1000-he-heptasystems?ID¼1730e08d-f43a-46ea-b7f37b35c04c36eb 10. Weng LX, Deng HH, Xu JL, Li Q, Zhang YQ, Jiang ZD, Li QW, Chen JW, Zhang LH (2011) Transgenic sugarcane plants expressing high levels of modified cry1Ac provide effective control against stem borers in field trials. Transgenic Res 20:759–772

11. Joyce P, Hermann S, O’Connell A, Dinh Q, Shumbe L, Lakshmanan P (2014) Field performance of transgenic sugarcane produced using agrobacterium and biolistics methods. Plant Biotechnol J 12:411–424 12. Gao SW, Yang YY, Xu LP, Guo JL, Su YC, Wu QB, Wang CF, Que YX (2018) Particle bombardment of the cry2A gene cassette induces stem borer resistance in sugarcane. Int J Mol Sci 19:E1692 13. Kannan B, Jung JH, Moxley GW, Lee S-M, Altpeter F (2018) TALEN-mediated targeted mutagenesis of more than 100 COMT copies/ alleles in highly polyploid sugarcane improves saccharification efficiency without compromising biomass yield. Plant Biotechnol J 16:856–866 14. Kim JY, Gallo M, Altpeter F (2012) Analysis of transgene integration and expression following biolistic transfer of different quantities of minimal expression cassette into sugarcane (Saccharum spp. hybrids). Plant Cell Tissue Organ Cult 108:297–302 15. Franks T, Birch RG (1991) Gene-transfer into intact sugarcane cells using microprojectile bombardment. Aust J Plant Physiol 18:471–480 16. Snyman SJ, Meyer GM, Richards JM, Haricharan N, Ramgareeb S, Huckett BI (2006) Refining the application of direct embryogenesis in sugarcane: effect of the developmental phase of leaf disc explants and the timing of DNA transfer on transformation efficiency. Plant Cell Rep 25:1016–1023 17. Manickavasagam M, Ganapathi A, Anbazhagan VR, Sudhakar B, Selvaraj N, Vasudevan A, Kasthurirengan S (2004) Agrobacteriummediated genetic transformation and development of herbicide-resistant sugarcane (Saccharum species hybrids) using axillary buds. Plant Cell Rep 23:134–143 18. Tilbrook K, Gnanasambandam A, Schenk PM, Brumbley SM (2010) Efficient targeting of polyhydroxybutyrate biosynthetic enzymes to plant peroxisomes requires more than three amino acids in the carboxyl-terminal signal. J Plant Physiol 167:329–332 19. Rathus C, Bower R, Birch RG (1993) Effects of promoter, intron and enhancer elements on transient gene expression in sugarcane and carrot protoplast. Plant Mol Biol 23:613–618 20. Zhang MQ, Zhuo XL, Wang J, Wu Y, Yao W, Chen R (2015) Effective selection and regeneration of transgenic sugarcane plants using

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positive selection system. In Vitro Cell Dev Biol Plant 51:52–61 21. Beyene G, Buenrostro-Nava MT, Damaj MB, Gao SJ, Molina J, Mirkov TE (2011) Unprecedented enhancement of transient gene expression from minimal cassettes using a double terminator. Plant Cell Rep 30:13–25 22. Taparia Y, Fouad WM, Gallo M, Altpeter F (2012) Rapid production of transgenic sugarcane with the introduction of simple loci following biolistic transfer of a minimal expression cassette and direct embryogenesis. In Vitro Cell Dev Biol Plant 48:15–22 23. Murashige T, Skoog F (1962) A revised medium for rapid growth and bioassays with

tobacco tissue cultures. Physiol Plant 15:473–497 24. Vain P, McMullen MD, Finer JJ (1993) Osmotic treatment enhances particle bombardment-mediated transient and stable transformation of maize. Plant Cell Rep 12:84–88 25. Thomson D, Dietzgen RG (1995) Detection of DNA and RNA plant viruses by PCR and RT-PCR using a rapid virus release protocol without tissue homogenization. J Virol Methods 54:85–95 26. Sun Y, Joyce PA (2017) Application of droplet digital PCR to determine copy number of endogenous genes and transgenes in sugarcane. Plant Cell Rep 36:1775–1783

Chapter 12 Genetic Transformation of Common Wheat (Triticum aestivum L.) Using Biolistics Caroline A. Sparks and Angela Doherty Abstract The following protocol describes the genetic transformation of wheat using the BioRad PDS/1000-He particle delivery system. Immature embryos are isolated 12–16 days post-anthesis, the embryonic axis is removed, and the immature scutella are precultured for 1–2 days prior to particle bombardment. Gold particles are coated with plasmid DNA containing the gene(s) of interest plus a selectable marker gene, in this instance bar (bialaphos resistance), and are fired into the cells to deliver the DNA. Subsequent tissue culture and regeneration steps allow recovery of plantlets, assisted by the inclusion of PPT (phosphinothricin tripeptide), the active ingredient of glufosinate-ammonium containing herbicides, to help select transformants. This updated method introduces selection earlier in the regeneration process which provides a shortened protocol while maintaining high transformation efficiencies. Key words Biolistics, Gene gun, Particle bombardment, Wheat, Genetic transformation, Immature embryo, Transgenic plants, DNA delivery, Tissue culture, Regeneration

1

Introduction Although a variety of methods of DNA delivery have been explored over the years, by far the simplest concept must be that of “shooting” DNA into cells. Transformation via biolistics had its origin in the 1980s [1, 2] with only minor adaptations having been made to the particle gun in the intervening period, for example, exchanging the bullet for helium gas, and DNA delivery using this device remains a fundamental tool in plant transformation studies. Wheat proved to be one of the more challenging crops to transform with initial success being achieved through bombardment in the early 1990s [3]. PEG (polyethylene glycol)-mediated transformation and microinjection were not viable options for wheat if stably transformed plants were required as it is not possible to regenerate from wheat protoplasts. As a consequence, particle bombardment was the primary transformation method for a number of years until mechanisms to allow Agrobacterium tumefaciens to infect

Sachin Rustgi and Hong Luo (eds.), Biolistic DNA Delivery in Plants: Methods and Protocols, Methods in Molecular Biology, vol. 2124, https://doi.org/10.1007/978-1-0716-0356-7_12, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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monocotyledonous species made possible wheat transformation by this method in 1997 [4]. Both biolistics and Agrobacteriummediated transformation continue to be widely used for research. Although there are advantages and disadvantages to both methods, particle bombardment offers much more flexibility than Agrobacterium. Co-bombardment of multiple constructs (including those designed for Agrobacterium transformation) is possible via biolistics. Alternatively, fragments of DNA, that is, isolated gene cassettes, can be introduced, thereby avoiding incorporation of vector backbone and antibiotic resistance marker genes. There is also potential to bombard entities other than DNA (e.g., RNA or proteins). This latter possibility could be particularly useful for the latest CRISPR (clustered regularly interspaced short palindromic repeats) gene-editing technology where, rather than bombarding gene cassettes which become integrated into the genome, the pre-assembled Cas9 ribonucleoprotein (RNP) complex could be introduced. Having generated gene edits, the protein complex would be naturally degraded thus plants generated using this method may have reduced offsite cleavage and avoid continued cleavage of DNA in subsequent generations. It is also possible such plants may avoid GM regulation depending on government rulings. Since even the latest transformation technologies still require some sort of delivery method to the cell, although originally a simple concept and design, the particle gun is still relevant and remains a valid and valued technique, not yet superseded by other practices. For wheat, although the main parameters for the particle gun have largely been optimized, there are still opportunities to improve the transformation process by being more precise in the choice of explants and by modifying tissue culture and regeneration steps. Scutella from immature embryos are still the explant of choice for wheat as they give the best response in tissue culture for regeneration of plants. Although there have been some reports of success with alternative explants (e.g., mature seed callus [5] and shoot apical meristems [6]), immature embryos are generally much more reliable and result in better transformation efficiencies. This chapter describes an updated version of a previously published standard wheat transformation system [7] which now delivers even higher and more consistent transformation efficiencies and is also more rapid due to earlier introduction of plant selection [8].

2 2.1

Materials Media

All stock solutions and media are prepared using reverse osmosis, polished water with a resistivity of 18.2 MΩ/cm (megaohm/cm). Unless otherwise specified, chemicals are generally tissue culture tested or analytical grade and supplied by Sigma-Aldrich,

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UK. Sterilization is carried out by autoclaving at 121
C for 15 min or by filter sterilization using a 0.22 μm syringe filter (Fisher Scientific, UK) or, for larger volumes, 0.2 μm MediaKap filters (VWR International Ltd., UK) connected to a peristaltic pump. Stock solutions can be stored at 4
C for 1–2 months or frozen at 20
C for up to a year provided no freeze-thawing has occurred (see Note 1). Petri dishes and Magenta vessels containing medium can be stored at 4
C for 3–4 weeks (see Note 2). 2.1.1 Stock Solutions

The following stock solutions are required to prepare the stock solutions of plant culture media and final plant culture media listed in Subheadings 2.1.2 and 2.1.3. 1. MS (Murashige and Skoog) Macrosalts (10): 19.0 g/L KNO3, 16.5 g/L NH4NO3, 4.4 g/L CaCl2·2H2O (see Note 3), 3.7 g/L MgSO4·7H2O, and 1.7 g/L KH2PO4. Autoclave and store at 4
C. 2. L7 Macrosalts (10): 15.0 g/L KNO3, 2.5 g/L NH4NO3, 3.5 g/L MgSO4·7H2O, 4.5 g/L CaCl2·2H2O (see Note 3), and 2.0 g/L KH2PO4. Autoclave and store at 4
C. 3. L Microsalts (100): 1.67 g/L MnSO4 (see Note 4), 750 mg/ L ZnSO4·7H2O, 500 mg/L H3BO3, 75.0 mg/L KI, 25.0 mg/L Na2MoO4·2H2O, 2.5 mg/L CuSO4·5H2O, and 2.5 mg/L CoCl2·6H2O. Autoclave and store at 4
C. 4. MS vitamins (100): 10 g/L myoinositol, 200 mg/L glycine, 100 mg/L thiamine hydrochloride, 50 mg/L pyridoxine hydrochloride, 50 mg/L nicotinic acid. Prepare 200 mL volume and store at 4
C. 5. Modified MS vitamins (100): As for MS vitamins (100) but omit the glycine. Prepare 200 mL volume and store at 4
C. 6. Amino acids, 3AA solution (25): 18.75 g/L L-glutamine (see Note 5), 3.75 g/L L-proline, 2.5 g/L L-asparagine (see Note 6). Aliquot into 40 mL volumes and store at 20
C. 7. Vitamins/Inositol (200): 40 g/L myoinositol, 2 g/L thiamine hydrochloride, 200 mg/L pyridoxine hydrochloride, 200 mg/L nicotinic acid, 200 mg/L calcium pantothenate, 200 mg/L ascorbic acid. Aliquot into 50 mL volumes and store at 20
C. 8. 2,4-Dichlorophenoxyacetic acid (2,4-D) (1 mg/mL): Dissolve 50 mg 2,4-D in 35 mL 70% ethanol until completely dissolved. Make up to 50 mL volume with distilled water. Filter-sterilize. Aliquot into 1 mL volumes and store at 20
C. 9. Indole-3-Butyric Acid (IBA) (100 mg/mL): Add 1 N NaOH dropwise to 5 mg IBA until completely dissolved. Make up to 50 mL with distilled water. Filter-sterilize. Aliquot into 10 mL volumes and store at 4
C.

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10. Picloram (100 mg/L): Add 1 N NaOH dropwise to 25 mg picloram until completely dissolved. Make up to 250 mL with distilled water. Store at 4
C. 11. Zeatin (mixed isomers) (Melford Laboratories Ltd., UK) (10 mg/mL): Add 1 N NaOH dropwise to 500 mg zeatin until completely dissolved. Make up to 50 mL with distilled water. Filter-sterilize. Aliquot into 1 mL volumes and store at 20
C. 12. Silver nitrate (AgNO3) (20 mg/mL): Dissolve 1 g AgNO3 in 50 mL distilled water. Filter-sterilize. Aliquot into 1 mL volumes and store at 20
C in the dark (see Note 7). 13. Copper sulfate (CuSO4·5H2O) (25 mg/mL): Dissolve 2.5 g CuSO4·5H2O in 100 mL distilled water. Filter-sterilize. Aliquot into 1 mL volumes and store at 4
C. 14. Ascorbic acid (100 mg/mL): Dissolve 5 g of ascorbic acid in 50 mL distilled water. Filter-sterilize. Aliquot into 10 mL volumes and store at 4
C. 15. Glufosinate ammonium (PPT) (Greyhound Chromatography and Allied Chemicals, UK) (10 mg/mL): Dissolve 500 mg glufosinate ammonium in 50 mL distilled water. Filter-sterilize (see Note 8). Aliquot into 1 mL volumes and store at 20
C. 2.1.2 Plant Culture Media Stock Solutions

1. M9% (2): Add 200 mL MS Macrosalts (10), 20 mL L Microsalts (100), 20 mL ferrous sulfate chelate solution (100), 20 mL Modified MS Vitamins (100), 40 mL 3AA Solution (25) and 180 g sucrose to 500 mL of distilled water. Make up to 1 L and adjust to pH 5.7 with 5 M NaOH. The osmolarity should be between 800 and 1100 mOsm (milliosmole). Filter-sterilize and store at 4
C. 2. R (2): Add 200 mL L7 Macrosalts (10), 20 mL L Microsalts (100), 20 mL ferrous sulfate chelate solution (100), 10 mL Vitamins/Inositol (200), and 60 g maltose to 500 mL distilled water. Make up to 1 L and adjust to pH 5.7 with 5 M NaOH. The osmolarity should be within the range 269–298 mOsm. Filter-sterilize and store at 4
C. 3. Agargel® (2): Prepare in 800 mL volumes at 10 g/L (see Note 9). Autoclave and store at room temperature, melting in a microwave before using in the final culture media (Subheading 2.1.3) (see Note 10).

2.1.3 Final Culture Media

The induction, regeneration and rooting media are prepared from the stock solutions of plant culture media (Subheading 2.1.2) which are made at double strength to allow the addition of an equal volume of double concentration gelling agent (Subheading 2.1.2) (see Notes 10 and 11). The first and second selection media

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are solidified with agarose, which is added directly to the medium prior to autoclaving (see Note 2). 1. Induction medium: Mix an equal volume of M9% (2) with sterilized, melted Agargel®. Add 0.5 mg/L 2,4-D and 10 mg/ L AgNO3. Pour into 9 cm petri dishes, approximately 28 mL per dish. Store at 4
C in the dark (see Notes 7 and 12). 2. First selection medium (P5): Add 100 mL MS Macrosalts (10), 10 mL L Microsalts (100), 10 mL ferrous sulfate chelate solution (100), 10 mL MS vitamins (100), 0.5 g glutamine, 100 mg casein hydrolysate, 750 mg MgCl2·6H2O, 1.95 g MES [2-(N-morpholino)ethanesulfonic acid], 0.5 mg/ L 2,4-D, 2.2 mg/L picloram and 40 g maltose to 600 mL distilled water. Make up to 1 L, adjust to pH 5.8 with 5 M NaOH and add 5 g agarose (Type I, low EEO). Autoclave at 121
C for 15 min, cool to ~50
C then add 100 mg/L ascorbic acid, 0.85 mg/L AgNO3 and 5 mg/L PPT. Pour into 9 cm petri dishes, approximately 28 mL per dish. Store at 4
C in the dark (see Note 7). 3. Second selection medium (P10): As for first selection medium (P5) but add 10 mg/L PPT. Pour into 9 cm petri dishes, approximately 28 mL per dish. Store at 4
C in the dark (see Note 7). 4. Regeneration Medium (RZP5): Mix equal volumes of R (2) and sterilized, melted Agargel®. Add 5 mg/L zeatin, 0.1 mg/L 2,4-D, 25 mg/L CuSO4·5H2O (see Note 13) and 5 mg/L PPT. Pour into 9 cm petri dishes, approximately 28 mL per dish. Store at 4
C. 5. Rooting Medium (RP5): Mix equal volumes of R (2) and sterilized, melted Agargel®. Add 0.1 mg/L IBA and 5 mg/L PPT. Pour into GA-7 magenta vessels (Sigma-Aldrich, UK), approximately 60 mL per vessel. Store at 4
C. 2.2 Isolation of Target Explants 2.2.1 Growth of Donor Plants

Wheat seeds (Triticum aestivum L. variety Cadenza) are sown fortnightly to provide a regular supply of material for transformation. Plants are grown, five per 21 cm pot, in Rothamsted prescription mix soil (Petersfield products, UK; see Note 14), top-watered by hand initially then watered automatically by flood benching once established. Leaves are stripped from the plants at about 5 weeks to leave only the strongest 5–6 tillers. Lighting in the controlled environment rooms is provided by banks of hydrargyrum quartz iodide (HQI) lamps 400 W (Osram Ltd., UK) to give an intensity of ~700 μmol/m2/s photosynthetically active radiation (PAR) at pot level with a 16 h day length. Temperature is maintained at 18
C day and 15
C night. Good plant husbandry and routine cleaning of controlled environment rooms keep pests and diseases to a minimum but Amblyseius caliginosus [Nursery Trades (Lea

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Valley) Ltd., UK] is used as a biological control agent against thrips and a single spray with a mildew preventative, e.g., Talius [DuPont (UK) Ltd., UK] is applied at 4–6 weeks. The condition of the donor plants is critical for determining the state of the immature embryos and therefore transformation success (see Note 15). 2.2.2 Sterilization of Immature Caryopses

1. 70% (v/v) aqueous ethanol. 2. 10% (v/v) aqueous bleach (see Note 16). 3. Sterile distilled water.

2.3 Components for Particle Bombardment

1. 0.6 μm (submicron) gold microcarriers (Bio-Rad Laboratories Ltd., UK) (see Note 17). 2. Spermidine (free base) 0.1 M: Initially prepare a 1 M solution by dissolving 1 g spermidine powder in 6.89 mL sterile distilled water. Aliquot into 50 μL volumes and store immediately at 80
C. For the working stock, dilute a 50 μL aliquot of 1 M spermidine with 450 μL sterile distilled water to give 0.1 M. Aliquot into 25 μL volumes and store immediately at 20
C (see Note 18). 3. Calcium chloride (Fisher Scientific UK Ltd., UK) 2.5 M: Dissolve 3.67 g CaCl2·2H2O in 10 mL water. Mix well by vortexing. Filter-sterilize. Aliquot into 55 μL volumes and store at 20
C. 4. Macrocarriers, macrocarrier holders, stopping screens, 650 psi (pounds per square inch) rupture discs (see Note 19) (all Bio-Rad Laboratories Ltd., UK). 5. Plasmid DNA (1 mg/mL): Prepare plasmids using a Qiagen plasmid purification kit (Qiagen Ltd., UK) or similar, to give good-quality DNA (see Note 20). Plasmids carrying the gene (s) of interest and also a selectable marker cassette (on the same or different plasmid for co-bombardment) are required (see Notes 8, 21 and 22).

3

Methods

3.1 Isolation of Target Explants 3.1.1 Collection and Sterilization of Immature Caryopses

1. Select ears from donor plants grown in controlled environment rooms (see Subheading 2.2.1) at approximately 12–16 days post-anthesis (see Note 23). Pick out the immature caryopses from the outer florets of each spikelet only, avoiding the top and bottom few spikelets of the ear (see Note 24). 2. Surface sterilize the immature caryopses by immersing in 70% (v/v) aqueous ethanol for 3–5 min, followed by 10% (v/v) aqueous bleach for 5–10 min with occasional gentle shaking.

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Fig. 1 Development of calli and regeneration of putative transgenic plantlets. (a) Example of sizes of isolated immature scutella, from top to bottom: 2.5, 2.2, 2.0, 1.7, 1.5 mm. (b) Immature scutella ~1.8 mm isolated ready for transformation. (c) Scutellum development on first selection medium (P5). (d) Embryogenic callus formation on second selection medium (P10). (e) Somatic embryos forming shoots and roots on RZP5 medium. (f) Regeneration of plantlets in Magenta vessels on RP5 medium. (g) Putative transgenic plantlets potted to soil in GM glasshouse. Scale bar ¼ 1 mm approximately

3. Remove the bleach solution and rinse the caryopses several times in plenty of sterile distilled water, draining away as much water as possible after the final wash (see Note 25). 3.1.2 Isolation of Immature Scutella

1. Isolate immature embryos from the caryopses in sterile conditions with the aid of a stereo microscope (see Note 26): the embryos should be 1.7–2.5 mm in length and translucent in appearance (see Note 27, Fig. 1a). 2. Remove the embryonic axis (see Note 28) and place the remaining scutellum with the cut surface face-down in the central region of a plate of Induction medium (see Note 29). Isolate 30 scutella per plate (see Note 30, Fig. 1b). 3. Seal the plates with Parafilm® (Fisher Scientific, UK) and incubate at 22–24
C in the dark for 1–2 days prior to bombardment (see Note 31).

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3.2 Preparation of Gold Microcarriers for Particle Bombardment 3.2.1 Preparation of Gold Microcarriers Stock

1. Weigh 40 mg 0.6 μm gold into an Eppendorf tube. 2. Working in a sterile environment, add 1 mL 100% ethanol and sonicate for 2 min. 3. Centrifuge in a benchtop microcentrifuge at top speed for a 3 s pulse. Discard the supernatant. 4. Repeat steps 2 and 3 twice more. 5. Add 1 mL sterile distilled water and sonicate for 2 min. 6. Centrifuge in benchtop microcentrifuge at top speed for a 3 s pulse. Discard the supernatant. 7. Repeat steps 5 and 6. 8. Resuspend fully in 1 mL sterile distilled water to give 40 mg/ mL concentration (see Note 32). Aliquot 50 μL amounts into sterile Eppendorf tubes, vortexing between taking each aliquot to ensure an even mix. Store at 20
C.

3.2.2 Coating of Gold Microcarriers with DNA

1. Working in a sterile environment, defrost one 50 μL aliquot of gold microcarriers (see Subheading 3.2.1) for each plasmid to be transformed (see Notes 33 and 34). Sonicate for 3 min (see Note 35) then vortex briefly. 2. Add 5 μg DNA to the side of the tube (see Notes 21, 22, 34 and 36), then vortex for 2–3 s to mix. 3. Mix together 50 μL 2.5 M CaCl2·2H2O and 20 μL 0.1 M spermidine in the lid of the Eppendorf tube. Carefully close the lid without displacing the solution then tap the tube to amalgamate with the gold + DNA and immediately vortex for 5 s to mix (see Note 37). 4. Incubate at room temperature for 2–3 min to allow the DNA to bind fully to the particles. 5. Centrifuge at top speed in a benchtop microcentrifuge for 3–5 s. Discard the supernatant. 6. Wash the gold particles in 150 μL 100% ethanol, resuspending as thoroughly as possible (see Note 38). 7. Centrifuge at top speed in a benchtop microcentrifuge for 3–5 s. Discard the supernatant. 8. Resuspend in 85 μL 100% ethanol (see Notes 39 and 40), seal the tubes with Parafilm® to prevent evaporation and maintain the particles on ice before use (see Note 41).

3.3 Transformation Via Particle Bombardment

Particle bombardment is carried out using the PDS-1000/He particle delivery system (Bio-Rad Laboratories Ltd., UK) according to the manufacturer’s instructions. This is a high-pressure system, using pressurized helium to accelerate subcellular particles to high velocity. Although helium is neither flammable nor toxic, any gas under high pressure should be handled with caution and appropriate safety precautions should be taken when operating this device.

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1. Various parameters can be altered within the particle gun apparatus. For transformation of wheat immature scutella as detailed here, the system has been optimized using the following settings: 2.5 cm gap (distance between rupture disc and macrocarrier), 5.5 cm target distance (distance between stopping screen and target plate), 0.8 cm stopping plate aperture (distance between macrocarrier and stopping screen), 28–3000 Hg vacuum, 5.0 vacuum flow rate, 4.5 vacuum vent rate. 2. Sterilize the gun chamber, rupture disc retaining cap, microcarrier launch assembly, target shelf and macrocarrier insertion tool using 90% (v/v) aqueous ethanol and allow to evaporate completely. 3. Sterilize macrocarriers, macrocarrier holders, stopping screens and rupture discs by dipping in 100% ethanol (see Note 42) and allow to dry in a sterile environment. Allow one set per shot. 4. Once dry, press a macrocarrier into its holder using the red plastic macrocarrier insertion tool (see Note 43). Place 5 μL DNA-coated gold microcarriers (see Subheading 3.2.2) per macrocarrier, vortexing between taking each aliquot (see Note 44) and spread the particles gently with the pipette tip over the central area. Allow the particles to dry slowly on a non-vibrating surface (see Notes 45 and 46). 5. Open the helium cylinder main valve and adjust the regulator to ~200 psi above the pressure of the rupture disc being used (see Note 47).

3.3.2 Assembling the Gun

1. Fit a rupture disc into the base of the rupture disc retaining cap (see Note 48). Screw the retaining cap onto the gas acceleration tube within the gun chamber and tighten fully using the mini torque wrench (see Note 49, Fig. 2a). 2. Place a stopping screen into the fixed nest of the microcarrier launch assembly (see Note 50). Invert a macrocarrier holder containing a macrocarrier loaded with DNA-coated gold particles and place over the stopping screen, screwing on the retaining ring to hold in place (see Note 51). Mount the microcarrier launch assembly onto the top shelf to give a 2.5 cm gap from rupture disc to macrocarrier (see Fig. 2b). 3. Place a petri dish with target tissues (prepared Subheading 3.1.2) on the target stage and remove the lid. Mount the stage on the shelf third down from the top to give the 5.5 cm target distance from macrocarrier to target tissue (see Fig. 2c). Close the chamber door.

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Fig. 2 PDS-1000/He particle gun to show the key component parts of the assembly 3.3.3 Firing the Gun

1. Switch on the particle gun. 2. Draw a vacuum in the gun chamber to >2800 Hg then set the vacuum switch to “Hold” (see Note 52). 3. Press and hold the “Fire” button to allow the helium to enter the gas acceleration tube and build up behind the rupture disc (see Note 53). Once the correct pressure is reached the rupture disc will burst (see Notes 54 and 55); release the “Fire” button immediately after the shot. 4. Release the vacuum in the chamber by moving the button to the “Vent” position and, once the “Fire” button is no longer illuminated, open the chamber door. 5. Remove the plate of bombarded target tissues and cover with a lid.

3.3.4 Disassembling the Gun

1. Remove the microcarrier launch assembly and disassemble: place the macrocarrier holder and stopping screen in 100% ethanol to resterilize if required and discard the used macrocarrier (see Note 56). 2. Free the rupture disc retaining cap using the mini torque wrench then unscrew it fully. Remove and discard the burst rupture disc (see Note 57). 3. Repeat the assembly/disassembly and firing process for the remaining shots (see Note 58).

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1. Close the main valve on the helium cylinder and release the regulator adjusting screw. Draw a vacuum in the chamber of ~1000 Hg such that the “Fire” switch illuminates. Press and release the “Fire” switch several times until the pressure is drained from the regulator as shown by the pressure reducing on the regulator gauge. Turn off the vacuum pump and gun. 2. Spray out the gun chamber and all component parts using 90% (v/v) aqueous ethanol and allow to evaporate. 3. Remove the macrocarrier holders and stopping screens from 100% ethanol, rinse in water and place in 10% (v/v) aqueous Savlon® to soak. Sonicate for 5 min to eradicate any residual DNA and prevent carry over.

3.4 Regeneration and Selection of Transgenic Plantlets Following Bombardment

3.4.1 Induction of Embryogenic Callus

Following the transformation step, the scutella pass through a number of tissue culture phases to induce embryogenic callus formation (Subheading 3.4.1), introduce selection (Subheading 3.4.2) increase the selection pressure (Subheading 3.4.3), regenerate plantlets (Subheading 3.4.4) and promote rooting (Subheading 3.4.5) prior to potting to soil (Subheading 3.4.6). Each step below has specified intervals but experience will also help with gauging when and which material to transfer. 1. Following bombardment, spread the scutella more evenly across the medium, dividing between two petri dishes of Induction medium, that is, 15 scutella per plate (see Note 59). 2. Seal the plates with Parafilm® and incubate in the dark at 22–24
C (see Note 60).

3.4.2 Introduction of Selection

1. After 1 week the scutella should have enlarged, appear healthy and look bright with some visible embryogenesis; scutella which are fading to yellow or have a brownish tinge are unlikely to respond further. Transfer responsive calli to the first selection medium (P5) (see Notes 61 and 62). Calli from the control plates should be moved to medium with and without selection to demonstrate the capacity to regenerate and/or the effectiveness of selection (see Note 58). 2. Seal the plates with Parafilm® and continue to incubate in the dark at 22–24
C for 2 weeks (see Note 60, Fig. 1c).

3.4.3 Increasing Selection Pressure

1. After 2 weeks on first selection medium (P5), transformed calli should have continued to grow, remaining quite bright and have larger regions of embryogenesis. Transfer calli to second selection medium (P10), ignoring any which are unresponsive, that is, wet, glutinous, or discoloring; these should be discarded. The control calli should be transferred to medium  PPT as previously (see Note 58). At this stage it should be

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possible to judge the likely success of the bombardment according to the level of response. 2. Seal the plates with Parafilm® and incubate at 22–24
C in the dark for 3 weeks (see Note 60, Fig. 1d). 3.4.4 Regeneration of Plantlets

1. Six weeks after the initial bombardment it should be obvious whether the callus is transformed; the callus pieces should have grown quite prolifically and may have small green regions visible. Transfer embryogenic calli to Regeneration medium RZP5 (see Note 63). The calli might be quite fragile at this point and may break up into pieces; keep all the pieces from one callus together and mark them so that plants from each initial callus can be tracked (see Note 64). The control calli should be transferred to medium  PPT as previously (see Note 58). 2. Seal the plates with Parafilm® and culture at 22–24
C but now exposed to light for 2 weeks (see Note 60, Fig. 1e).

3.4.5 Rooting of Plantlets

1. Following 2 weeks on Regeneration medium RZP5, the calli should have small green shoots and also some small roots although some calli may just be knobbly and green (see Note 65). Transfer any material which is embryogenic and green to Rooting medium RP5 in GA-7 magenta vessels (see Note 66, Table 1). The control calli should be transferred to medium  PPT as previously (see Note 58). 2. Culture at 22–24
C in the light for 2 weeks (see Note 60, Fig. 1f).

3.4.6 Potting Putatively Transformed Plantlets to Soil

1. When the plantlets are strong enough and have an established root system they can be potted to soil (see Note 67, Table 1). Remove the plantlets from the Magenta vessel, rinsing the roots with water to remove any Agargel® if necessary, and pot into moistened soil (Rothamsted prescription mix (see Note 14)), in 4 cm square pots [Nursery Trades (Lea Valley) Ltd., UK] (see Notes 64 and 68). 2. Top water the pots and place in a tray containing damp capillary matting to keep the soil moist. Place the tray under a propagator lid (see Note 69) and grow in an appropriate GM containment greenhouse (see Note 70), removing the propagator lid after approximately 1 week (see Fig. 1g).

3.4.7 Analysis of Plants and Further Growth

Approximately 2 weeks after potting to soil, the plantlets should have become sufficiently established to allow leaf samples to be taken for extraction of genomic DNA (see Notes 71 and 72) on which PCR analysis can be carried out for the gene of interest and the selectable marker gene and any other analysis as required. Following this, transfer confirmed transformed plants to larger

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Table 1 Timeline for transformation and regeneration of transgenic plants Day 0

Isolate immature scutella onto induction medium

Day 1

Bombard tissues and spread more evenly between 2 and 3 plates of induction medium

Day 7

Transfer to first selection medium (P5)

Day 21

Transfer to second selection medium (P10)

Day 42

Transfer to regeneration medium RZP5

Day 56

Transfer to rooting medium RP5

Day 70

Pot putative transgenic plantlets to soil

Day 84

Pot on confirmed transgenic plants

10 cm square pots [Nursery Trades (Lea Valley) Ltd., UK] with Rothamsted prescription mix soil (see Note 14) and grow to maturity in an appropriate GM containment glasshouse (see Notes 68 and 70).

4

Notes 1. Sterile stock solutions can be stored at 4
C for 1–2 months. Some settling of salts may occur during storage, therefore check the solutions and shake well prior to use in order to resuspend them if necessary. Stock solutions stored at 20
C should remain effective for at least a year, provided no freeze/ thawing has occurred. 2. Petri dishes and Magenta vessels with tissue culture media should be prepared as freshly as possible but always a few days in advance of use to allow for detection of contamination. Once prepared, petri dishes and Magenta vessels with media can be wrapped and stored at 4
C for 3–4 weeks. 3. Before mixing with other components, dissolve CaCl2·2H2O in water. 4. MnSO4 is available in various hydrated states which will alter the required weight: for MnSO4·H2O, add 1.71 g/L, MnSO4·4H2O, add 2.32 g/L, or for MnSO4·7H2O, add 2.80 g/L. 5. If the L-glutamine is difficult to dissolve, dissolve it separately at pH 9.0 prior to mixing with the other amino acids. 6. Instead of using the 3AA stock solution, 1.5 g/L L-glutamine, 0.3 g/L L-proline, and 0.2 g/L L-asparagine can be added individually to the M9% (2) medium.

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7. Silver nitrate induces stress which helps to promote embryogenesis in the initial tissue culture stages; it is photosensitive so the stock solution and any media plates containing it should be kept in the dark. 8. A commonly used selectable marker gene is bar, which confers resistance to glufosinate ammonium-based broad-spectrum herbicides such as Basta®, Challenge®, Liberty®, and Harvest®. Glufosinate ammonium (PPT), the active ingredient of these herbicides, is added as a supplement to certain tissue culture media to destroy any tissues which have not incorporated the selectable marker DNA. Other options for plant selection include antibiotics (e.g., kanamycin, hygromycin) and positive selection (e.g., phosphomannoisomerase [PMI]). 9. Generally, only 800 mL volumes of Agargel® are prepared in 1 L bottles to allow for easier mixing and to prevent boil-over when autoclaving. 10. The Agargel® solution should be shaken well, both before and after autoclaving, to avoid non-uniform solidification which leads to difficulties when remelting. Agargel® is a blend of agar and Phytagel™ that was developed to help control vitrification in plant tissue cultures. This product provides the positive attributes of both products and is superior to Phytagel, where vitrification is a problem. It also serves as an economical alternative to agar. Agargel® produces a semiclear gel which allows better detection of contamination. The product should be used at a concentration of 3.5–5.0 g/L, depending upon the desired gel strength. 11. To minimize condensation in the plates, allow the remelted Agargel® to cool to ~50
C prior to mixing with other media components and pouring the final medium. 12. This chapter describes the protocol for transformation of wheat variety Cadenza (hard, red, spring bread wheat). Over 45 other wheat or wheat-related varieties (including spring and winter bread wheat, durum wheat, and introgression lines) have been transformed using this protocol but modifications to the induction medium may be required. For example, the choice of basal salts (MS or L7), the concentration of sugars (sucrose or maltose) and the level of hormones need to be empirically determined. 9% sucrose partially plasmolyzes the cells during preculture which may increase their ability to withstand bombardment [9]. However, this is variety and species dependent; 3% or 4% sucrose is often more appropriate, for example, for scutella of T. turgidum ssp. durum. 13. Copper sulfate is a stress-inducing agent (similar to silver nitrate) used to promote shooting; an increased level of the micronutrient copper in the culture medium dramatically

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improves regeneration (see Ref. 10). 25 mg/L (100 μM) is the recommended copper sulfate concentration, but if shooting is too prolific, the concentration can be reduced by half. 14. Rothamsted prescription mix soil from Petersfield Products, UK, contains: 75% medium grade peat, 12% sterilized loam, 3% medium grade vermiculite, 10% grit (5 mm screen, lime-free), 3.5 kg Osmocote Exact 3/4 month/m3 (slow-release fertilizer, 16N/11P/11K plus micronutrients, supplied by Scotts UK Professional, Ipswich, Suffolk), 0.5 kg of PG mix/m3 (14N/16P/18K granular fertilizer plus micronutrients, supplied by Hydro Agri (UK), Ltd., Lincs.), lime ~3 kg to pH 5.5–6.0, Vitax Ultrawet at 200 mL/m3 (wetting agent). 15. The condition of the donor plants is critical to transformation success; specific light intensities, wavelengths of light, photoperiod, watering, diseases, and so on are all key factors which can influence the transformation outcome. The authors have experienced fluctuations in transformation efficiencies related to even minor alterations in growth conditions, so a failure to get transformed plants could be associated with this primary, fundamental stage. While field-grown or glasshouse-grown material can be used, this will not give the uniformity required for consistent, reliably high transformation efficiencies. 16. Commercially available thin bleach can be used for dilution; this commonly has a sodium hypochlorite content of 4–6% (v/v). 17. Gold particles of alternative size are available (e.g., Heraeus 0.4–1.2 μm diameter [W. C. Heraeus GmbH & Co. KG, Germany]), but the more uniform 0.6 μm particles have proved most successful for wheat due to the small cell size of the target tissue. 18. Spermidine solutions are hygroscopic and oxidizable, and deaminate with time, and consequently should be maintained below 20
C (preferably at 80
C) and fresh stocks should be made regularly. Once thawed any unused aliquots should be discarded. 19. Rupture discs are available from Bio-Rad Laboratories Ltd. as 450, 650, 900, 1100, 1350, 1550, 1800, 2000, and 2200 psi. The wheat bombardment protocol described here has been optimized using 650 psi rupture discs in combination with other particle gun parameters (gap, target distance, etc.). For alternative target explants or different species, empirical testing is required to determine the optimum rupture pressure. 20. Plasmid DNA needs be of good quality and purity as contaminants can lead to clumping of gold during the DNA coating process. A DNA concentration of 1 mg/mL is recommended; lower concentrations can be used but preferably not less than

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0.6 mg/mL otherwise dilution of solutions during DNA precipitation onto the gold can occur. 21. A reporter gene (such as uidA encoding β-glucuronidase or a fluorescent protein, e.g., green fluorescent protein, GFP, red fluorescent protein, RFP) can be included in order to monitor the effectiveness of shots and therefore verify transformation or help optimize the delivery parameters. 22. Particle bombardment not only provides the opportunity to co-bombard multiple constructs, it also allows for the introduction of vectors designed for Agrobacterium transformation, DNA of isolated gene cassettes free of vector-backbone DNA (i.e., no antibiotic resistance marker gene), and potentially Cas9 ribonucleoprotein (RNP) complexes for gene editing. Such possibilities make particle bombardment a versatile and less restrictive transformation system compared to Agrobacterium-mediated transformation. 23. Experience is required to determine the best ears to use; generally, the immature caryopses are just visible between the glumes, the endosperm should be solidifying but still soft (medium milk stage) and the embryo should be approximately 2 mm long. A few immature embryos can be crudely isolated when collecting the ears to gauge the size if necessary. If the ears are ready but explants cannot be isolated that day, keep the ears with the stalk in water at 4
C overnight. 24. Only caryopses from the outer florets and those in the central region of the spike are used to ensure that the embryos isolated are at a similar developmental stage. 25. It is advisable not to store caryopses once sterilized as they will not perform well if maintained in damp conditions. If the scutella will not be isolated immediately, the caryopses can be kept unsterilized at 4
C overnight. 26. A microscope eye piece graticule can be used to provide an accurate measure of embryo size. This may be useful to help select consistently sized embryos while becoming familiar with the technique but the appearance of the embryo can be more meaningful than the actual size. 27. Embryos which give the best response are usually in the size range 1.7–2.5 mm and will have a whitish, semitranslucent appearance. Opaque, creamy-yellow embryos should be avoided as these are too mature. 28. The embryonic axis is removed to prevent precocious germination which would diminish the callus-forming potential of the scutellum tissue. 29. Typically, the gun shot fires most gold particles within a central ~2 cm diameter circular area of a petri dish. Arranging scutella

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within this area maximizes particle delivery and hence transformation efficiency. 30. 30 scutella per treatment plate are usually a convenient number to isolate and also handle in later tissue culture stages. Additional plates with fewer scutella can be isolated to act as controls both with and without selection. 31. Pre-plasmolysis of the scutella tissues prior to bombardment increases their chances of survival; 1–2 days’ preculture is generally optimal. 32. The recommended concentration of gold is 40 mg/mL, but half this concentration can be just as effective. If using the lower concentration, add an equal volume of sterile distilled water to an aliquot of 40 mg/mL gold prior to coating with DNA (see Subheading 3.2.2). However, the higher gold concentration is advised for transient transformation experiments to maximize the number of cells targeted. 33. Each 50 μL aliquot of gold should result in a sufficient volume of coated gold for 10–12 shots; the amount of gold together with other components can be scaled up or down accordingly, for example, if only four shots are required, start with 25 μL gold. 34. A gold aliquot without DNA should be prepared for each experiment to act as a control in order to show the effect of bombardment with gold particles only on the tissue culture response. 35. Sonication should disperse any clumps and resuspend the gold. However, over-sonication should be avoided as there is evidence that it may have the opposite effect and lead to particle agglomeration. 36. The DNA added can be a single construct (with one or more gene cassettes) or alternatively multiple constructs can be co-bombarded. If a selectable marker gene cassette is not present in the construct alongside the gene of interest cassette, a separate selectable marker gene construct needs to be included. Provided individual constructs are not too large, up to four can be successfully co-bombarded and will reliably generate transgenic plants containing all four genes. When co-precipitating constructs onto the gold, equimolar ratios should be calculated to give equivalent amounts of each construct. Additionally, if desired, a 1.5:1 ratio of the gene of interest construct(s) to selectable marker gene construct can be used which skews toward the gene of interest; plants surviving the selection process are therefore more likely to contain the gene of interest also. Regardless of the number of constructs co-bombarded and the ratios used, the 5 μg DNA per 50 μL aliquot of gold should not be exceeded as this will result in clumping of the

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gold; the DNA is added in excess, so it is better to add less rather than more. 37. The precipitation of DNA onto the gold microcarriers is very rapid. CaCl2·2H2O and spermidine stabilize, precipitate and bind the DNA to the gold and mixing together first ensures a more uniform coating of gold with less clumping of particles. 38. Use the pipette tip to scrape the sides and base of the Eppendorf tube to resuspend the particles and remove small clumps of gold which could cause damage to the target tissues. If the microcarriers are not smooth at this point it will be difficult to remove clumps later, consequently, if clumps remain, it is better to discard the aliquot and start again, first ensuring the volume and concentration of the DNA added are correct. 39. The gold preparation should not be aspirated too much at this stage or the ethanol will evaporate and increase the overall concentration of the particles; the reduced volume will also mean fewer shots are possible. 40. For transient assays, the gold can be resuspended in a smaller final volume to increase the concentration of particles and maximize the number of cells targeted. 41. Coated gold particles should be used as soon as possible for bombardment. Alternatively, the particles can be kept at the wash stage (see Subheading 3.2.2, step 6) and resuspended in the final ethanol volume just prior to use, but this should never be for longer than an hour. 42. The rupture discs are composed of laminate layers of plastic, so they should only be dipped briefly in ethanol and not soaked, otherwise the layers may separate. 43. The macrocarriers must be properly mounted in their holders otherwise they may not release correctly during firing which would result in a failure to discharge the gold particles. 44. Vortex the gold preparation between taking each sample to ensure an equal loading of particles on each macrocarrier. 45. The macrocarriers can be examined microscopically prior to bombardment to determine the uniformity and spread of particles and ensure that they will not do too much damage to the target tissues. Greater experience with the gold coating process will ultimately be sufficient to determine if the gold will be suitable. 46. Only a few macrocarriers should be prepared at any one time and they should be used as soon as possible after the ethanol has evaporated.

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47. The regulator should be set to ~200 psi above the intended rupture pressure in order to ensure sufficient helium is allowed through the system to accumulate and burst the rupture disc. 48. The rupture disc must be perfectly seated in the rupture disc retaining cap to completely seal it onto the gas acceleration tube. Failure to do this will result in a leak in the system meaning the helium cannot build to the required pressure to burst the disc and no firing will take place. 49. Tighten the rupture disc retaining cap fully with the mini torque wrench to prevent any escape of helium gas which could also prevent firing due to lack of pressure. 50. During a shot, the macrocarrier is released from the macrocarrier holder and strikes the stopping screen thereby discharging the microcarriers. Omitting to add a stopping screen will allow the whole macrocarrier to pass through the hole in the fixed nest, potentially damaging and/or contaminating the target tissue. 51. At this point the gold particles should be on the underside of the macrocarrier, facing toward the target tissue. 52. The level of vacuum drawn is important to ensure successful transformation as it reduces air resistance thereby allowing the microcarriers to retain their speed and direction; generally, the higher the vacuum the better but it should always be >2600 Hg. Certain wheat varieties seem to be somewhat sensitive to being subjected to a vacuum in which case the vacuum drawn may need to be toward the lower end of the range. 53. If the helium pressure gauge does not register any helium entering the system, firstly check the cylinder has been opened and the regulator has been set. Assuming they have, either the rupture disc has not been seated correctly in the rupture disc retaining cap or the cap itself has not been fully tightened onto the gas acceleration tube with the mini torque wrench. Abort the shot by releasing the vacuum button to “Vent” and check the cap and rupture disc, tightening the cap or repositioning the disc as necessary. 54. If the rupture disc doesn’t burst even when the expected pressure is reached (as shown on the helium pressure gauge), two rupture discs may have been placed in together. In this case, release the “Fire” button, return the vacuum button to “Vent” and remove the rupture disc retaining cap. If two rupture discs are present, remove and discard both discs and use a new one; rupture discs already put under pressure should not be reused as they may have been weakened. 55. Monitor and record the actual pressure at which the rupture disc bursts; if this is lower/higher than the anticipated pressure

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the microcarriers may not have been released as expected which could affect the transformation efficiency. 56. The macrocarrier should show a mesh pattern where it has been released from its holder and struck the stopping screen; gold particles carrying DNA will have passed through the mesh to penetrate the target cells. 57. If the burst rupture disc is not present in the rupture disc retaining cap it may have remained attached to the gas acceleration tube. The disc must be removed, otherwise it could interfere with subsequent shots. 58. Within any experiment, controls should be incorporated. These should include non-bombarded scutella to assess the suitability of the embryos to form embryogenic callus and regenerate; scutella bombarded with gold but without DNA to assess the effect of the bombardment process on tissues and their response; some plates with and without selective agent to verify the effectiveness of the selection. 59. Scutella are bombarded within the central target region of a petri dish so in order to develop individually following bombardment they need to be spread out to provide more space and have less competition for nutrients. However, if scutella have been bombarded to detect transient expression only, for example, from a reporter gene, they need not be spread. 60. The controlled environment culture room is maintained at 22–24
C with 12 h photoperiod provided by cool white fluorescent tubes supplying ~100 μmol/m2/s photosynthetically active radiation. For incubation in the dark, solid trays are used and covered with foil. 61. The selection agent to include will depend on the selectable marker gene used; this could be either an herbicide or its active ingredient such as PPT described here, or an antibiotic, for example, G418 (see Note 8). For any selection agent, the appropriate concentration to use will need to be determined by carrying out a kill curve on the target tissue. There may even be some variation in survival between different wheat varieties due to inherent natural resistance; for example, Cadenza has quite high natural resistance to PPT. Consequently, the kill curve should be performed ideally on the actual variety to be transformed. 62. Although selection can be introduced at a later stage, for example, at regeneration, provided the correct concentration of selective agent is used, earlier selection will kill non-transformed material much sooner. The amount of calli to be transferred is therefore decreased which not only saves time but also reduces the expense of media and other consumables.

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63. Some calli may only have small areas that are embryogenic, in which case, only move these embryogenic fragments; any calli which haven’t grown or aren’t embryogenic should be discarded. 64. Plants which originate from the same initial callus may be clonal and they should be treated as such until proven otherwise. It is important therefore to group any pieces of calli together in order to trace plant origins and label plants accordingly. 65. Proper small plantlets may be recognizable at this stage but this can be dependent on the wheat variety being used. 66. Transfer not only the calli with shoots/roots but also the knobbly green calli as some of these may produce roots and shoots on the RP5 medium. 67. A strong root system is required to enable the plantlets to survive in soil; incubate the plantlets for longer than 2 weeks prior to potting up if roots are slow to develop. 68. A plant should also be potted up which originates from a control plate bombarded with gold but without DNA followed by growth on media without plant selection; this will act as a negative control plant which has been through the same experimental conditions as transgenic plants but without DNA. 69. The trays should be placed under a propagator lid as plantlets from tissue culture have an inadequate waxy cuticle and therefore require humid conditions to prevent desiccation until the cuticle forms. 70. For newly potted plants, the greenhouse temperature is maintained at 22
C day/18
C night for 3–4 weeks. Following this, GM glasshouse conditions are maintained at 18–20
C day/ 14–16
C night temperatures with a 16 h photoperiod provided by natural light supplemented with banks of Phytolux Atis-7 LED lamps which generate a light intensity of approximately 400 μmol/m2/s photosynthetically active radiation (PAR) at canopy level. 71. When sampling, never cut the most recently emerged leaf as this may kill the plant; always sample one of the older leaves. Leaf samples are frozen immediately in liquid nitrogen and can be used either straight away for DNA extraction or stored at 80
C until required. 72. Genomic DNA is generally extracted using the Promega Wizard kit (Promega, UK) or the CTAB (cetyl trimethylammonium bromide) method [11]. The authors have found such methods preferable in order to generate reasonable yields of good quality DNA with low contamination. Other kits are available which allow, for example, direct PCR from plant tissue but these have not been found to be sufficiently reliable.

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Acknowledgments Caroline Sparks and Angela Doherty (Rothamsted Research) receive grant-aided support from the Biotechnology and Biological Sciences Research Council (BBSRC) Designing Future Wheat project [BB/P016855/1]. References 1. Sanford JC, Klein TM, Wolf ED, Allen N (1987) Delivery of substances into cells and tissues using a particle bombardment process. Particulate Sci Technol 5:27–37 2. Klein TM, Wolf ED, Wu R, Sanford JC (1987) High-velocity microprojectiles for delivering nucleic acids into living cells. Nature 327:70–73. https://doi.org/10.1038/ 327070a0 3. Vasil V, Castillo AM, Fromm ME, Vasil IK (1992) Herbicide resistant fertile transgenic wheat plants obtained by microprojectile bombardment of regenerable embryogenic callus. Nat Biotechnol 10:667–674 4. Cheng M, Fry JE, Pang SZ, Zhou HP, Hironaka CM, Duncan DR, Conner TW, Wan YC (1997) Genetic transformation of wheat mediated by Agrobacterium tumefaciens. Plant Physiol 115:971–980 5. Medvecka´ E, Harwood WA (2015) Wheat (Triticum aestivum L.) transformation using mature embryos. In: Wang K (ed) Agrobacterium protocols. Methods in molecular biology, vol 1223. Springer, New York, NY 6. Hamada H, Linghu Q, Nagira Y, Miki R, Taoka N, Imai R (2017) An in planta biolistic

method for stable wheat transformation. Sci Rep 7:11443 7. Sparks CA, Jones HD (2014) Genetic transformation of wheat via particle bombardment. In: Henry RJ, Furtado A (eds) Cereal genomics: methods and protocols, Methods in molecular biology, vol 1099. Humana Press, New York, NY 8. Ishida Y, Tsunashima M, Hiei Y, Komari T (2015) Wheat (Triticum aestivum L.) transformation using immature embryos. In: Wang K (ed) Agrobacterium protocols. Methods in molecular biology, vol 1223. Springer, New York, NY 9. Brettschneider R, Becker D, Lo¨rz H (1997) Efficient transformation of scutellar tissue of immature maize embryos. Theor Appl Genet 94:737–748 10. Lazzeri PA c/o Las Tres Calas SA, RascoGaunt S (2002) Improved transformation and regeneration of wheat using increased copper levels. European Patent Office, EP1409671A1 11. Doyle JJ, Doyle JL (1987) A rapid procedure for DNA purification from small quantities of fresh leaf tissue. Phytochem Bull 19:11–15

Chapter 13 Biolistic DNA Delivery in Turfgrass Embryonic Callus Initiated from Mature Seeds Man Zhou, Junming Zhao, Dayong Li, Shuangrong Yuan, Ning Yuan, Zhigang Li, Haiyan Jia, Fangyuan Gao, Bekir San, Qian Hu, and Hong Luo Abstract We describe a protocol for the establishment and preparation of creeping bentgrass (Agrostis stolonifera L.) cultivar “Penn A-4” embryonic calli, biolistic transformation, selection, and regeneration of transgenic plants. The embryonic callus is initiated from mature seeds, maintained by visual selection under the dissecting microscope and subjected to bombardment with plasmid DNA containing a bialaphos-resistance (bar) gene. PCR, Southern, and Northern blot analyses are used to confirm the transgene integration and expression. Key words Biolistic, DNA delivery, Gene gun, Genetic transformation, Embryonic callus, Mature seed, Turfgrass

1

Introduction Biolistic transformation (also referred to particle bombardment method, the particle gun method, ballistic method, and the microprojectile method) is a process which employs high-velocity microprojectiles to deliver DNA, protein, nucleoprotein complex, or dye into cells [1, 2]. This method has been widely applied in many plant species, particularly monocot plants such as rice, maize, wheat, tall fescue, pearl millet, sugarcane, and creeping bentgrass (Agrostis stolonifera L.), to name a few [3–12]. Although Agrobacteriummediated transformation is the method of choice in many cases, biolistic-mediated transformation is versatile and effective for some cereal crops recalcitrant to Agrobacterium-facilitated transformation, or proof of concept, and functional genomics studies [10]. For biolistic transformation, embryonic suspension cultures or callus cultures induced from mature seeds or embryos can serve as

Man Zhou, Junming Zhao, Dayong Li, Shuangrong Yuan, Ning Yuan contributed equally to this work. Sachin Rustgi and Hong Luo (eds.), Biolistic DNA Delivery in Plants: Methods and Protocols, Methods in Molecular Biology, vol. 2124, https://doi.org/10.1007/978-1-0716-0356-7_13, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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suitable explants. The popular selectable markers used are similar to that used in the Agrobacterium-mediated transformation method, such as hygromycin phosphotransferase gene (hph) and phosphinothricin acetyltransferase gene (bar). The integration and expression of these genes in the transformed cells render resistance to the antibiotic hygromycin and the herbicide phosphinothricin (PPT). The utilization of selectable markers facilitates the selection and regeneration process of the transformed cells. Rice actin, maize ubiquitin, and cauliflower mosaic virus (CaMV) 35S promoters are commonly used in the gene expression cassettes [2, 7]. Turfgrass is among the most important perennial grasses providing numerous environmental, societal, and economic benefits. Like in many row crops, genetic engineering of turfgrass using transgenic approach offers the opportunity to incorporate many desirable traits which are otherwise difficult or rather impossible to achieve through the conventional breeding methods. The use of a transgenic approach in developing new cultivars will become increasingly more important as the demand for better turfgrass is continuously increasing and the feasibility of commercializing transformed cultivars becomes physically possible. Considerable efforts have been directed in turfgrass toward establishing and optimizing different transformation methods to achieve various objectives [3, 5, 7, 8, 10, 13–16]. Here, we present a biolistic transformation procedure using PPT for selection in Penn A-4, a commercial cultivar of creeping bentgrass, an important coolseason turfgrass species.

2

Materials

2.1

Plant Material

Creeping bentgrass cultivar, Penn A-4 is used to induce embryogenic calli from mature seeds (caryopses, Fig. 1a). The culture is incubated at room temperature (24  2  C) in the dark. The induced embryonic calli (Fig. 1b) are visually inspected, selected, and transferred to fresh callus initiation medium every 2 weeks to maintain them in good condition before biolistic transformation (for details, see Subheading 3.1).

2.2

Plasmids

The plasmid pSBUbibar-35SGUS is used for transformation [16, 17], which contains a selectable marker gene, bar and a reporter gene, GUS driven by the corn ubiquitin promoter and the CaMV 35S promoter, respectively. Expression of the bar gene leads to herbicide resistance in the transformed cells. QIAGEN HiSpeed Plasmid Maxi Kit (QIAGEN, Inc., CA) is recommended to prepare pure plasmid DNA for genetic transformation experiment.

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Fig. 1 The illustration of (a) mature seeds for callus initiation, (b) induced callus, (c) herbicide-resistant callus, (d) regenerated shoots on the R1 medium, (e) regenerated roots on the R2 medium, and (f) regenerated whole plants in soil grown in the greenhouse 2.3 Culture Media and Solutions 2.3.1 Chemical Stock

1. 3,6-Dichloro-o-anisic acid (Dicamba): Prepare Dicamba to a final stock concentration of 6.6 mg/mL in double distilled water (ddH2O) and store it at 4  C. Use 1 mL of stock solution per liter of media. 2. 6-Benzylaminopurine (BAP): Prepare the BAP solution to a final concentration of 0.5 mg/mL in NaOH and make the final volume using ddH2O. Store the BAP solution at 4  C for later applications. Use 1 mL of stock solution per liter of media. 3. Phosphinothricin (PPT, Duchefa): Prepare the PPT stock to a final concentration of 4 mg/mL in ddH2O. Use 1 or 2 mL of stock per liter of media. The stock is filter-sterilized before use (see Note 1) and stored at 4  C.

2.3.2 Initiation of Embryogenic Callus from Mature Seeds

1. Sterilization solution for seeds: Clorox® bleach (6% sodium hypochlorite) and Tween-20™ (Polysorbate 20; PhytoTechnology Labs, Shawnee Mission, KS, USA). For creeping bentgrass, use 10% (v/v) bleach solution and 0.2% (v/v) Tween20™. 2. See Table 1 for the creeping bentgrass callus-initiation medium (MMSG).

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Table 1 The creeping bentgrass callus-initiation medium (MMSG) Ingredients

Final concentration

Murashige and Skoog (MS) basal salts (Sigma; [18])

4.3 g/L

1000 Gamborg vitamins (Phytotechnology)

1 mL/L

Sucrose

30 g/L

Casein hydrolysate

500 mg/L

Dicamba

6.6 mg/L

BAP

0.5 mg/L

Phytagel

2 g/L

Use KOH to adjust the pH of the medium to 5.7 before autoclaving at 121  C for 20 min

Table 2 List of ingredients used for the regeneration medium R1 Ingredients

Final concentration

MS basal salts

4.3 g/L

Surcose

30 g/L

Myoinositol

100 mg/L

1000 Gamborg vitamins

1 mL/L

BAP

1 mL/L

Phytagel

2 g/L

Use KOH to adjust the pH of the medium to 5.7 before autoclaving at 121  C for 20 min. After cooling the media to 55  C in a water bath, add 8 mg/L filter-sterilized PPT

2.3.3 Plant Selection and Regeneration Media

1. Selection media: For bar gene selection, add phosphinothricin (PPT) at the rate of 4 mg/L to the MMSG medium. 2. For regeneration medium R1 used for shoot induction, see Table 2. 3. For regeneration medium R2 used for root induction, see Table 3. 4. For details of the osmotic medium, see Table 4.

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Table 3 List of ingredients used for the regeneration medium R2 Ingredients

Final concentration

MS basal salts

4.3 g/L

Surcose

30 g/L

Myoinositol

100 mg/L

1000 Gamborg vitamins

1 mL/L

Phytagel

2 g/L

Using KOH to adjust the pH of the medium to 5.7 before autoclaving at 121  C for 20 min. After cooling the media to 55  C in a water bath, add 8 mg/L filter-sterilized PPT

Table 4 List of ingredients used for the osmotic medium Ingredients

Final concentration

Murashige and Skoog (MS) basal salts (Sigma; [18]) 4.3 g/L 1000 Gamborg vitamins (Phytotechnology)

1 mL/L

Sucrose

30 g/L

Casein hydrolysate

500 mg/L

Dicamba

6.6 mg/L

BAP

0.5 mg/L

Sorbitol

0.2 M/L

Mannitol

0.2 M/L

Phytagel

2 g/L

Using KOH to adjust the pH of the medium to 5.7 before autoclaving at 121  C for 20 min

3

Methods

3.1 Initiation and Proliferation of Embryonic Callus from Mature Seeds

1. Add approximately 40 mL of 70% ethanol to a 50 mL falcon tube filled with 10 mL of creeping bentgrass seeds and shake on an orbital shaker at 175 rpm for 2 min. Rinse the seeds three times using sterile ddH2O (see Note 2). 2. Add 40 mL seed sterilization solution to the seeds washed with 70% ethanol and shake them at 175 rpm for 30 min. Repeat this step two more times. Then rinse the seeds five times using sterile ddH2O to make sure the bleach or other chemicals have been completely removed. Now the seeds are ready for culturing.

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3. Place the sterilized seeds onto callus-initiation medium (MMSG). Put seeds (approximately 50–80) gently on the medium with the help of forceps or using a pipette seated with a precut tip in a laminar flow hood (see Note 3). 4. The culture is maintained in the dark at room temperature for 2–3 weeks. Once the seeds germinate, approximately 15 germinated seedlings are transferred to fresh callus initiation media to allow more space for growing. 5. The embryogenic calli which resemble Type II callus of maize are selected by dissecting microscope and then placed onto the fresh callus-initiation medium (Fig. 1b) (cf. ref. 19, and see Note 4). 6. All cultures are incubated in the dark. Every 2–3 weeks, the embryogenic calli can be divided into 0.5-cm pieces for subculturing. Repeat this subculturing process until enough colonies are prepared for transformation. 7. About 1 week before the genetic transformation, subculture the embryonic calli again (see Note 5). 3.2 Callus Preparation Prior to Bombardment

Four hours before the bombardment, 8- to 12-week-old embryogenic calli are placed evenly onto the center of an osmotic medium in 3 cm diameter (see Note 6).

3.3 Biolistic Transformation of Creeping Bentgrass Embryogenic Calli with pSBUbibar-35SGUS

1. The DNA/gold coating procedure [7] is carried out as follows: 2 mg of 0.6 μm gold particles are added to 5 μL of plasmid DNA solution of pSBUbibar-35SGUS (1 μg/μL) to form a suspension (see Note 7). Then 220 μL of sterile distilled water, 250 μL of 2.5 M calcium chloride, and 50 μL of 0.1 M spermidine are added to the suspension with mixing upon each addition. The mixture is vortexed at 4  C for 10 min, and the supernatant is discarded after centrifugation at 850  g for 5 min. The coated DNA is pelleted and then washed with 600 μL of absolute ethanol and resuspended in 38 μL of absolute ethanol. 2. Place 7 μL of the gold–DNA solution on each microcarrier for each bombardment (see Note 8). 3. Four hours before the bombardment, the embryogenic calli are transferred to the osmotic medium (see Subheading 3.2). 4. A PDS1000/He particle delivery system (Bio-Rad, Hercules, CA, USA) is used to perform the bombardment. The parameters used in for bombardment are listed in Table 5. 5. Each plate of prepared calli on the osmotic medium is bombarded once. The bombarded calli are transferred from the osmotic medium to callus initiation medium 1–24 h after the bombardment [7].

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Table 5 List of parameters used for the particle bombardment

3.4 Selection and Regeneration of Transformed Colonies

Parameters

Value

Rupture disk pressure

1100 psi

Target distance

6 cm

Gap distance

6 mm

Stopping plate aperture

1 cm

Torr vacuum at rupture

27–28

1. Four to seven days after the biolistic transformation, the colonies are placed on selection medium with 4 mg/L filtersterilized PPT. If the selectable marker gene bar is introduced into embryogenic calli through the plasmid DNA-coated gold particles, the colonies should be resistant to PPT (Fig. 1c). 2. After 2 selective cultures at 25  C in dark, resistant calli are carefully selected and transferred to the R1 medium (Fig. 1d) and then to the R2 medium (Fig. 1e) with 8 mg L 1 PPT and incubated at 25  C under fluorescent light (60 με m 2 s 1) (see Notes 9 and 10).

3.5 Regenerated Plants Maintenance

1. Regenerated plants are grown in commercial potting mixture soil (Fafard 3-B Mix, Fafard Inc., Anderson, SC, USA) and maintained in the greenhouse under a 16 h photoperiod with supplemental lighting at 27  C day and 25  C night temperatures (Fig. 1f). 2. Plants from individual transformation events are clonally propagated from stolons and grown in pots (15 cm  10.5 cm, Dillen Products, Middlefield, OH, USA) using commercial potting mixture soil as previously described [13, 15]. 3. One month after transfer into the soil, the regenerated plants are screened for herbicide resistance with 0.5% Finale® (a commercial brand of the herbicide glufosinate, which is the ammonium salt of PPT). The glufosinate-resistant plants are maintained in the greenhouse with regular fertilization, mowing, and irrigation (e.g., watered every other day, fertilized and clipped every 2 weeks), and used for further analysis.

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3.6 Molecular Analysis of Transgene Integration and Expression in the Transformed Plants 3.6.1 PCR Analysis

3.6.2 Southern Blot Analyses

1. Plant genomic DNA is extracted as previously described using the cetyltrimethylammonium bromide (CTAB) method [14]. 2. PCR was conducted in Bio-Rad Dual 48 Well DNA Engine Thermal Cycler with the bar gene-specific primers, which amplifies a product of 202 bp [20]. 3. The PCR program is set as follows: Denaturation at 94  C for 5 min; 28 cycles of denaturation at 94  C for 30 s, annealing at 58  C for 30 s, extension at 72  C for 30 s, and the final extension at 72  C for 5 min. Commercial Taq DNA polymerase (Promega Corporation, Madison, WI) can be used in PCR reactions. 1. Southern blot analyses are performed to confirm the transgene integration into the turfgrass genome. Total DNA is extracted as previously described using the cetyltrimethylammonium bromide (CTAB) method [14]. 2. The probe DNA for the bar gene is isolated using a QIAquick gel extraction kit (QIAGEN, CA) following the instructions of the manufacturer. After digestion of the genomic DNA with BamHI according to the supplier’s instructions (New England Biolabs, Beverly, MA, USA), DNA was electrophoresed on 0.8% agarose gels, transferred onto nylon membranes (GE Healthcare Bio-Sciences Corp., Piscataway, NJ, USA), and hybridized to 32P-labeled DNA probes of the bar gene. Hybridization is carried out in the modified Church and Gilbert buffer at 65  C following the standard protocol [21]. 3. Hybridizing fragments are detected by exposure of the membrane on a phosphor screen at room temperature overnight, and scanning on a Typhoon 9400 phosphorimager.

3.6.3 Northern Blot Analyses

1. Total RNA is isolated from the leaves of transgenic and wildtype control plants using TRIzol reagent (Invitrogen). 2. RNAs are subjected to formaldehyde-containing agarose gel electrophoresis and transferred onto Hybond-N+ filters (GE Healthcare Bio-Sciences Corp.). 3. The DNA fragment coding for the bar gene is used as a probe. Hybridization and membrane wash are performed following the standard protocol [21].

4

Notes 1. Some stock solutions such as PPT used in the tissue culture media are heat sensitive. To avoid degradation of the chemical components, this kind of stock solution is filter-sterilized in

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advance and added to the media after the media is autoclaved and cooled down to 55  C in a water bath. It is recommended to use a stir bar during the whole process. 2. It is recommended to rinse the seeds with sterile water under a laminar flow hood. 3. Too many seeds (>150) placed on the medium may result in inconsistent callus initiation and difficulty in the separation of calli. 4. It is recommended to use a dissecting microscope under sterile conditions to select embryogenic calli to obtain the healthiest and the most homogeneous embryogenic cell lines for transformation. 5. This step helps the calli stay fresh and serves as a checkpoint for contamination and allows the subcultured calli some time for recovery. 6. Although the calli are subcultured every 2–3 weeks, too many rounds of subcultures will also impact the health and desirability of the calli. Eight to twelve weeks old embryogenic calli are good for transformation. 7. Several studies in wheat, maize, sugarcane, and pearl millet have indicated that utilization of minimal transgene cassette instead of the whole plasmids could increase the transformation frequency of the single gene integration [11, 16, 22–24]. 8. One of the factors that impact the reproducibility of the biolistic transformation method is the uniformity of the DNA– gold suspension, which is influenced by the coating procedures. It is recently reported that the single transgene insertion can be achieved in higher frequencies by using nanogram quantities of DNA [23]. 9. When selecting the potentially transformed colonies, dead colonies should be discarded in time. Each independently transformed colonies should be kept in the distance to differentiate independent transformation events. Different potentially transformed colonies may have different growth speed. The timing of transferring them to the R1 or R2 medium is judged not only by incubation time but also by the size and health of the colonies. 10. For turfgrass callus subculturing, 2 weeks is the ideal incubation time. If the callus is placed on the same medium for over 4 weeks, the callus will be obviously stressed and undesirable for transformation anymore.

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Acknowledgments This project was partially supported by the Biotechnology Risk Assessment Grants (BRAG) Program Competitive Grant no. 2019-33522-30102 from the USDA National Institute of Food, Agriculture (NIFA) and the United States Golf Association, Inc. (USGA) to H.L. References 1. Sanford JC (1988) The biolistic process. Trends Biotechnol 6:299–302 2. Fribourg HA, Hannaway DB, West CP (2009) Tall fescue for the twenty-first century. American Society of Agronomy, Agronomy Monograph No. 53, Madison 3. Chen L, Auh C-K, Dowling P, Bell J, Lehmann D, Wang Z-Y (2004) Transgenic down-regulation of caffeic acid O-methyltransferase (COMT) led to improved digestibility in tall fescue (Festuca arundinacea). Funct Plant Biol 31:235–245 4. Cho M-J, Yano H, Okamoto D, Kim HK, Jung HR, Newcomb K, Le VK, Yoo HS, Langham R, Buchanan BB, Lemaux PG (2004) Stable transformation of rice (Oryza sativa L.) via microprojectile bombardment of highly regenerative, green tissues derived from mature seed. Plant Cell Rep 22:483–489 5. Hu Y, Jia W, Wang J, Zhang Y, Yang L, Lin Z (2005) Transgenic tall fescue containing the Agrobacterium tumefaciens ipt gene shows enhanced cold tolerance. Plant Cell Rep 23:705–709 6. Wang K, Frame B (2009) Biolistic gun-mediated maize genetic transformation. In: Scott MP (ed) Transgenic maize. Springer, New York, pp 29–45 7. Wang Y, Kausch AP, Chandlee JM, Luo H, Ruemmele BA, Browning M, Jackson N, Goldsmith MR (2003) Co-transfer and expression of chitinase, glucanase, and bar genes in creeping bentgrass for conferring fungal disease resistance. Plant Sci 165:497–506 8. Wang Z-Y, Ge Y (2006) Recent advances in genetic transformation of forage and turf grasses. In Vitro Cell Dev Biol Plant 42:1–18 9. Zhang K, Liu J, Zhang Y, Yang Z, Gao C (2015) Biolistic genetic transformation of a wide range of Chinese elite wheat (Triticum aestivum L.) varieties. J Genet Genomics 42:39–42 10. O’Kennedy MM, Stark HC, Dube N (2011) Biolistic-mediated transformation protocols for maize and pearl millet using pre-cultured

immature zygotic embryos and embryogenic tissue. In: Thorpe TA, Yeung EC (eds) Plant Embryo Culture. Springer, New York, pp 343–354 11. Jackson MA, Anderson DJ, Birch RG (2013) Comparison of agrobacterium and particle bombardment using whole plasmid or minimal cassette for production of high-expressing, low-copy transgenic plants. Transgenic Res 22:143–151 12. Hamada H, Linghu Q, Nagira Y, Miki R, Taoka N, Imai R (2017) An in planta biolistic method for stable wheat transformation. Sci Rep 7:11443 13. Li Z, Baldwin CM, Hu Q, Liu H, Luo H (2010) Heterologous expression of Arabidopsis H(+)-pyrophosphatase enhances salt tolerance in transgenic creeping bentgrass (Agrostis stolonifera L.). Plant Cell Environ 33:272–289 14. Luo H, Kausch AP, Hu Q, Nelson K, Wipff JK, Fricker CCR, Owen TP, Moreno MA, Lee J-Y, Hodges TK (2005) Controlling transgene escape in GM creeping bentgrass. Mol Breed 16:185–188 15. Zhou M, Li D, Li Z, Hu Q, Yang C, Zhu L, Luo H (2013) Constitutive expression of a miR319 gene alters plant development and enhances salt and drought tolerance in transgenic creeping bentgrass (Agrostis stolonifera L.). Plant Physiol 161:1375–1391 16. Zhou M, Hu Q, Li Z, Li D, Chen CF, Luo H (2011) Expression of a novel antimicrobial peptide Penaeidin4-1 in creeping bentgrass (Agrostis stolonifera L.) enhances plant fungal disease resistance. PLoS One 6:e24677 17. Hu Q, Kononowicz-Hodges H, NelsonVasilchik K, Viola D, Zeng P, Liu H, Kausch AP, Chandlee JM, Hodges TK, Luo H (2008) FLP recombinase-mediated site-specific recombination in rice. Plant Biotechnol J 6:176–188 18. Murashige T, Skoog F (1962) A revised medium for rapid growth and bio assays with tobacco tissue cultures. Physiol Plant 15:473–497

Turfgrass Biolistic Transformation 19. Chip Longo CL, Hu Q, Nelson K, Viola D, Hague J, Chandlee JM, Luo H, Kausch AP (2006) Turfgrasses. In: Wan K (ed) Agrobacterium Protocols. Humana Press Inc., Totowa, NJ, pp 83–95 20. Thompson CJ, Movva NR, Tizard R, Crameri R, Davies JE, Lauwereys M, Botterman J (1987) Characterization of the herbicide-resistance gene bar from Streptomyces hygroscopicus. EMBO J 6:2519–2523 21. Maniatis T, Fritsch EF, Sambrook J (1982) Molecular cloning: a laboratory manual. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY 22. Yao Q, Cong L, Chang JL, Li KX, Yang GX, He GY (2006) Low copy number gene transfer

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and stable expression in a commercial wheat cultivar via particle bombardment. J Exp Bot 57:3737–3746 23. Ismagul A, Yang N, Maltseva E, Iskakova G, Mazonka I, Skiba Y, Bi H, Eliby S, Jatayev S, Shavrukov Y, Borisjuk N, Langridge P (2018) A biolistic method for high-throughput production of transgenic wheat plants with single gene insertions. BMC Plant Biol 18:135 24. Lowe BA, Shiva Prakash N, Way M, Mann MT, Spencer TM, Boddupalli RS (2009) Enhanced single copy integration events in corn via particle bombardment using low quantities of DNA. Transgenic Res 18:831–840

Chapter 14 Use of Microspore-Derived Calli as Explants for Biolistic Transformation of Common Wheat Sachin Rustgi, Samneet Kashyap, Nii Ankrah, and Diter von Wettstein Abstract There are specific advantages of using microspores as explants: (1) A small number of explant donors are required to obtain the desired number of pollen embryoids for genetic transformation and (2) microspores constitute a synchronous mass of haploid cells, which are transformable by various means and convertible to doubled haploids therefore allow production of homozygous genotypes in a single generation. Additionally, it has been demonstrated in wheat and other crops that microspores can be easily induced to produce embryoids and biolistic approach to produce a large number of transformants. In view of these listed advantages, we optimized the use of microspore-derived calli for biolistic transformation of wheat. The procedure takes about 6 months to obtain the viable transformants in the spring wheat background. In the present communication, we demonstrated the use of this method to produce the reduced immunogenicity wheat genotypes. Key words Genetic transformation, Particle bombardment, Microspore embryogenesis, Wheat

1

Introduction Common wheat (Triticum aestivum L.), one of the major staple grains, is cultivated throughout the world from temperate to tropical regions and feeds about 30% of the world populations [1]. However, it prefers a temperate climate (https://wheat.org/wheat-inthe-world/). With the warming up of climate and the pollution, a stagnation in the wheat yield gain is witnessed and a northward shift in its cultivation zone is anticipated [2, 3]. Both stagnation in yield gain and a northward shift of the wheat production zone are expected to have a dramatic effect on the availability of wheat especially to the inhabitants of the resource-deprived countries. Additionally, with an increase in the world population, which is expected to touch a 9.7 billion mark by 2050, a 70% increase in wheat production is required (http://www.fao.org/docrep/016/

Diter von Wettstein was deceased at the time of publication. Sachin Rustgi and Hong Luo (eds.), Biolistic DNA Delivery in Plants: Methods and Protocols, Methods in Molecular Biology, vol. 2124, https://doi.org/10.1007/978-1-0716-0356-7_14, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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ap106e/ap106e.pdf). The yield gains from the current management practices and the breeding approach, which we have been relying on for the last 50 years, have reached to their upper limits and no further yield gains are expected until and unless we complement the current approach with new technologies. Therefore, it has been repeatedly emphasized by the experts like the Nobel laureate Dr. Norman Borlaug and the World Food Prize winner Dr. Sanjay Rajaram that the targeted 70% increase in wheat productivity can only be achieved by adapting to the recent technologies specifically the transgenic approach. Precise genetic manipulations which fall under the confines of genetic engineering, differ from the conventional breeding approaches in two ways: (1) the gene transfers are not limited by the crossability range of a species and can be possible even between prokaryotes and higher plants and (2) these changes are directed unlike the conventional crossbreeding or mutation breeding approach. Indeed, genetic changes can be now introduced in the genome of virtually any organism in a precise way using programmable nucleases such as meganuclease, zinc finger nuclease, TAL nuclease, or RNA guided nuclease [4–6]. In parallel to these developments, a number of gene delivery methods or genetic transformation methods were also investigated, which include particle bombardment or biolistic, use of bacterium with type 4 secretion system (e.g., Agrobacterium), electroporation, magnetofection, sonication, polyethylene glycol, use of plant viruses, cellpenetrating peptide (CPP), direct delivery of triplex-forming oligonucleotides (TFOs), and single or double-stranded RNA (BioDirect™) [7–11]. Despite, wheat is one of the first cereals, which was genetically transformed there is still no single widely accepted and reliable protocol for wheat genetic transformation [12, 13]. Therefore, wheat genetic transformation is not routine in most of the laboratories worldwide. However, among different genetic transformation procedures, biolistic is still the most popular method in common wheat, as Agrobacterium-based genetic transformation exhibited reduced efficiency, and other genetic transformation methods such as the use of cell-penetrating peptides, electroporation, and PEG need further optimization [7]. Furthermore, other methods such as the use of viruses for genetic transformation have so far only shown transient expression of the gene [14]. There is also a great debate on the use of different plant parts as explant in tissue culture and their use in genetic transformation [15]. In the past, different wheat tissues were used as explants the most commonly used being the scutellar calli and microspores [16, 17]. Some of the advantages of using microspores as explants are as follows: (1) each anther produces a large number of microspores, for instance, 2800 in Arabidopsis, 30,000 in Petunia, and 580–3000 in wheat. Therefore, a small number of explant donors

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are required for an experiment. (2) Microspores represent a synchronous mass of haploid uni
/binucleate cells with androgenic potential. (3) Microspores are transformable by various means, for instance, electroporation, magnetofection, cocultivation with Agrobacterium, particle bombardment, and sonication [18]. (4) Chromosome doubling can be achieved by the application of antimitotic chemicals such as colchicine or laughing gas or via spontaneous doubling. Therefore, homozygosity can be achieved in a single generation. Our group optimized the use of microspores for genetic transformation and identified the optimal electroporation conditions to transform microspores via a single electric pulse of 375 V while retaining their viability and determined conditions for cocultivation of microspores with Agrobacterium AGL-1 cells [19, 20]. The major drawback of the abovementioned methods is the low transformation efficiency relative to the biolistic method (cf. Refs 19, 20). Use of microspore-derived calli for genetic transformation was first demonstrated for barley by Peggy Lemaux and group [21], which inspired studies in wheat. Use of biolistic approach and microspores as explants for genetic transformation has thus been optimized in wheat and the procedure is reported in the present communication.

2

Materials

2.1

Plants

2.2

Stock Solutions

For the genetic transformation of microspore embryoids by particle bombardment, hard red spring wheat variety WestBred 926 was used as an explant donor. 1. The preparation of the induction medium Northwest Plant Breeding-99 (NPB-99) and regeneration medium 190-2 is as described in Brew-Appiah et al. [19]. 2. The preparation of the Modified Murashige and Skoog 5 (MMS5) regeneration/rooting media is as described in Brew-Appiah et al. [19]. 3. Luria-Bertani (LB) medium: 10 g tryptone, 5 g yeast extract, 5 g NaCl in 950 mL of deionized H2O. Adjust the pH to 8.0 with 5 N NaOH. Adjust the volume of the solution to 1 L with deionized H2O. For solid medium add Bacto agar to 15 g/L before autoclaving, sterilize by autoclaving for 20 min at 15 psi (1.05 kg/cm2) on liquid cycle. 4. Kanamycin stock solution (50 mg/mL): 500 mg kanamycin sulfate powder in 10 mL deionized H2O. Filter-sterilize antibiotic solution using a 0.22 μm syringe filter. Aliquot (200 μL) and store at
20  C.

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5. Ampicillin stock solution (100 mg/mL): Dissolve 1 g of sodium ampicillin in 10 mL deionized H2O. Filter-sterilize antibiotic solution using a 0.22 μm syringe filter. For filter sterilization, prewash the sterile filter (0.22 μm) by drawing through 50 mL of H2O. Then pass the ampicillin solution through the washed filter. Aliquot (200 μL) and store at
20  C. 6. CaCl2 (2.5 M): 11 g CaCl26H2O in 20 mL of sterilized deionized H2O. Sterilize the solution by passing it through a 0.22 μm filter. 7. Spermidine (0.1 M): 255 mg spermidine in 10 mL of sterilized deionized H2O. Store at
20  C. 8. Acetocarmine solution: 0.5 g carmine in 100 mL of 45% boiling acetic acid. Heat for another few minutes in a fume hood while stirring. Cool down the solution to room temperature, add one to two drops of a saturated ferric acetate solution and filer. Store acetocarmine solution at room temperature. 9. Colchicine stock solution (0.1%): 1 g colchicine in a liter of sterilized deionized water. Store the solution in refrigerator (take necessary precautions while handling colchicine). For working solution add five drops of dimethyl sulfoxide (DMSO) to 10 mL of the colchicine stock solution. Store at 4  C. 10. CuSO4 solution (2 mM): 500 mg CuSO4·5H2O in 1 L 0.4 M mannitol solution. Store at 4  C. 11. Mannitol solution (0.4 M): 72.85 g mannitol in 1 L of sterilized deionized H2O. Store at 4  C. 12. Maltose solution (21%): 21 g maltose in 100 mL of sterilized deionized water. Store at 4  C. 2.3 Specific Laboratory Equipment and Supplies

1. Refrigerated and regular desktop centrifuge. 2. Glass Bead Sterilizers (BS-1000). 3. Waring blender and vessel. 4. Gene gun (PDS-1000/He™ and Hepta™ Systems) (Fig. 1). 5. Inverted microscope with camera. 6. Stereomicroscope. 7. Table top horizontal airflow workstation. 8. pH meter, 9. Incubator shaker and tissue culture chamber. 10. Magnetic stirrer. 11. Vortex mixer. 12. Spectrophotometer.

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Fig. 1 Wheat genetic transformation workflow. (a) Axenically growing wheat explants in the Plant Growth Facility at the Clemson University Pee Dee Research and Education Center. (b) Primary spikes harvested and surface-sterilized with 75% ethanol for microspore isolation. (c) Primary spikes being surface-sterilized with 10% bleach solution. (d) Primary spikes under pretreatment with a solution containing 2 mM CuSO4 and 0.4 M mannitol to induce embryogenesis. (e) 20- to 21-day-old microspore embryoids in induction medium. (f) Gene gun

13. Filter paper disks. 14. Parafilm. 15. Nylon mesh. 16. Magenta boxes, sterile small funnels, and Erlenmeyer flasks. 17. Sterile petri dishes with lids. 18. Sterile filter-stopped tips. 19. Standard pipettes. 20. Sterile screw-cap polypropylene centrifuge tubes (15 mL).

3

Methods

3.1 Growth Conditions and Selection of Spikes

1. WestBred 926 plants are cultivated in 20  25 cm pots using Metro-Mix® 360 potting mixture (SunGro Horticulture, Anderson, SC, USA) in a glasshouse maintained at the day temperature of 20–23  C and night temperature of 14–16  C and a photoperiod of 18 h. 2. Fertilizer is applied once a week with nutrient water containing 200–250 ppm of water-soluble fertilizer [nitrogen (N), phosphorus (P2O5) and potassium (K2O)].

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3. Primary tillers at the Feeke’s developmental stage 10–10.1, showing a 3–5 cm slit in the boot are excised below the second node and placed in a clean container with distilled water. These boots are expected to have microspores at the mid to lateuninucleate stage of development (specifically in the florets located in the central part of the immature spike) (Fig. 1a, b). 4. Stage of microspore development is determined by microscopic examination of crushed anthers mounted with acetocarmine or water (use phase contrast when water is used for mounting). At uninucleate stage, the microspores are round in shape with a nucleus located opposite the germ pore (aperture). 5. Tillers can be stored by dipping them up to the second node (from the top) in the water at 4  C for a week without a significant loss of microspore viability and cover spikes with a plastic bag to minimize moisture loss. 3.2

Pretreatment

1. The boots are pretreated with 2 mM CuSO4 solution prepared in 0.4 M mannitol for 10–14 days at 4  C. This pretreatment increases the possibility of obtaining green seedlings. 2. After pretreatment the spikes were sterilized for 10 min with 10% commercial bleach solution (active ingredient 6.15% sodium hypochlorite) (Fig. 1c, d).

3.3 Isolation of Microspores

1. Carefully cut the florets from spikelets with a pair of sterile scissors and transfer sterilized florets into an autoclaved Waring blender cup containing 50 mL of 0.4 M mannitol solution. For this step use a chilled blender cup stored in a refrigerator after autoclaving. 2. Blend spikes for 5 s at 2200 rpm in a Waring blender, pass the resulting slurry through an autoclaved 100-μm mesh to reduce microspore debris. 3. Rinse microspore debris from mesh into blender cup with 50 mL of 0.4 M mannitol and blend for another 5 s at 2200 rpm in a Waring blender (see Note 1). Filter with 100-μm mesh as in step 2. Rinse the mesh cup three times with 5 mL of 0.4 M mannitol solution each time. 4. Pour filtrate onto an autoclaved 50-μm mesh filter to trap viable microspores. Rinse the mash with 10 mL of 0.4 M mannitol solution. 5. Invert the mesh cup and rinse off microspores that were trapped on the mesh with 15 mL of 0.4 M mannitol into a beaker. 6. Transfer the resulting microspore suspension to a 15 mL tube and centrifuge at 900 rpm (149  g) for 5 min.

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7. Pour off supernatant and add 10 mL of 0.4 M mannitol and centrifuge at 900 rpm for 5 min. 8. Pour off supernatant and resuspend the microspores in 2 mL of 0.4 M mannitol solution and layer it over 10 mL of 21% maltose solution. 9. Density gradient centrifugation at 800 rpm (118  g) for 3 min (see Note 2) followed by transfer of 2–3 mL of the upper part of the band to a new 15 mL tube and add 10 mL of mannitol. Centrifuge at 1000 rpm (184  g) for 5 min, pour off supernatant and repeat this process two more times. Finally suspend the pellet in 2–5 mL of the NPB-99 medium. 3.4 Preparation of Plant Material for Microprojectile Bombardment

1. The method for obtaining microspores is as described in Subheading 3.3. 2. Transfer microspores to a 60  15 mm petri dish at a minimum density of 1  104/mL and a maximum density of 1  107/ mL, as estimated with the hemocytometer (see Note 3). 3. Add mature ovaries of same or different wheat variety to the culture medium. Use 6–9 ovaries per dish (i.e., 3 ovaries per mL of culture medium) (see Note 4) [22]. Cover dishes and seal with Parafilm. 4. Incubate petri dish at 28  C in the dark for embryoid development. After 20–21 days of incubation, transfer petri dish to an incubator maintained at 28  C and equipped with a shaker (Fig. 1e) (see Note 5). 5. Transfer 35- to 40-day-old microspore embryoids to the regeneration media (190-2) in 60  15 mm petri dish and arrange them in the center of the petri plate prior to the bombardment with DNA-coated gold particles (about 20 embryos per 100 mm petri dish or a minimum of about 100 embryoids per petri dish).

3.5 Preparation of Gold Particles Stock Solution for Biolistic Transformation of Microspore-Derived Calli

1. Weigh 60 mg of 0.6 μm gold particles in 1.5 mL Eppendorf tube. Add 1 mL of 100% ethanol to it and vortex the ingredients for 3 min. Centrifuge the mixture at 16,873  g for 2–5 min using a benchtop microcentrifuge and discard the supernatant. 2. Add 1 mL of 70% ethanol to the pellet and mix ingredients by vortexing for 2 min. Incubate tube for 15 min at room temperature. Mix contents at least three times during this period. After incubation centrifuge tube at 16,873  g for 3 min and discard the supernatant.

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3. Add 1 mL of filter sterilized deionized H2O to the tube, vortex the ingredients for 10 s and let it stand for 1 min. Centrifuge tube at 16,873  g for 2 min and discard the supernatant. 4. Repeat steps 3 for two times. 5. Add 1 mL 50% glycerol to the Eppendorf tube and store it at
20  C for later use. The gold particle stock solution can be prepared 1 day in advance and can be stored at
20  C for at least 4 months. 3.6 Preparation of Gold Particles for Bombardment

1. Vortex the previously stored gold particle solution until fully suspended. Pipette out 50 μL of the gold suspension into a 1.5 mL microcentrifuge tube. 2. Add following ingredient to the gold suspension in the stepwise fashion, and vortex for 5 s after each addition: 18 μL of plasmid DNA adjusted at a concentration of 1 μg/μL, 50 μL of freshly prepared filter-sterilized 2.5 M CaCl2, and 20 μL of 0.1 M spermidine free-base. After addition of all ingredients, vortex the mixture for 30–40 min at 4  C. Subsequently, add 200 μL of absolute ethanol to the mixture, vortex the suspension for 5 s and centrifuge it at 3000 rpm for 30 s. Remove supernatant, and repeat the ethanol wash four times. 3. After final centrifugation step, resuspend the pellet in 30 μL of 100% ethanol, keep the mixture on ice until used.

3.7 Coating the Macrocarrier

1. Submerge macrocarriers in 100% ethanol in a petri dish. Use forceps to remove macrocarriers from ethanol and dry them on Kimwipes. 2. Load macrocarrier into the macrocarrier holder using the seeding tool. 3. Vortex the gold suspension (microcarriers) and quickly transfer 10 μL of it to the center of the macrocarrier and let it dry for 10 min.

3.8 Microprojectile Bombardment

1. For microprojectile bombardment the following steps are carried out in a serial order. 2. Turn on the vacuum pump. 3. Turn on the Helium cylinder valve slowly to 14,000 kpas, and set the regulator to 1500 psi. 4. Turn on power to the chamber. 5. Place rupture disk into center of screw-on cylinder, and tighten the screw cylinder with torque tool. 6. Place stooping screen and macrocarrier holder and screw the lid.

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7. Place tissue on shelf (second from the bottom) and close the door. 8. Turn on vacuum switch. When the vacuum gauge reads 27, it is ready to fire. 9. Turn vacuum switch to “hold,” and press “fire” switch. 10. After firing turn the vacuum switch to vent. 11. Bombarded embryoids plated on regeneration media (190-2) are kept in the dark for 3 days at 28  C for root development and then exposed to 16 h light for shoot growth. 3.9

Regeneration

1. In a laminar hood, transfer green germinated plantlets to test tube or magenta box containing the rooting medium (MMS5). 2. Allow the plantlets to grow until the apex reaches the lid of the magenta box or test tube. 3. Transfer the plants to pots containing the Cornell mix and place the root trainer in a growth cabinet or glasshouse (see Note 6). 4. Test the T0 plants for transgenesis using PCR, qPCR, gel-blot analysis and DNA sequencing.

3.10 Colchicine Treatment

1. Plants at tillering stage are appropriate for colchicine treatment. Remove plant from the pot and thoroughly wash to eliminate soil particles. Trim roots down to 2–3 in. and remove one third of the shoot to expose the lateral bud. 2. Subsequently submerge the roots and crown in the colchicine working solution for 7 h (bubble solution with air during treatment). After colchicine treatment thoroughly wash the plant under running water and replant in soil.

3.11 Determination of Ploidy by Chromosome Analysis

1. Germinate T1 seeds of the colchicine treated (T0) plants at room temperature in germination boxes. Excise root tips when ~2–3 inches in size. Transfer root tips to 1.5 mL Eppendorf tubes containing chilled water followed by fixation at 0  C in an ice bucket containing ice water slurry. Pretreat root tips for 20–22 h. After pretreatment replace water with 3:1 solution of absolute ethanol and acetic acid. Keep the tubes for 1 week at room temperature and then store at 4  C until used. 2. Stain root tips in acetocarmine solution for 1 h, boil for a minute and then squash in 45% acetic acid. Put cover slip and gently tap the slide and heat gently on the flame of a spirit lamp. Press the cover slip evenly with thumb and observe under 10, 40 and 63/100 (use immersion oil) optical lenses. Count 42 chromosomes to confirm a euhexaploid wheat cell.

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Development of Wheat Genotypes with Reduced Immunogenicity Based on the research it is evident that development of a general dietary therapy for gluten-induced disorders is possible either by elimination or detoxification of the cause of these disorders [23– 25]. This hypothesis was tested by tissue-specific suppression of the wheat genes encoding a DNA glycosylase, DEMETER, and a Fe–S cluster biogenesis enzyme, Dre2 [26–28]. These genes collectively control transcriptional activation of about 100 different prolamins, except high-molecular-weight glutenins, which also are nonessential for baking [27, 28]. In order to achieve cosuppression of the master regulators of prolamin accumulation in wheat grains, the following two constructs were developed (1) a DEMETER TALErepressor-based donor construct and (2) a Dre2 gene-specific CRISPR/Cas9 construct. The donor construct carries a TALE repressor that targets a 17-nucleotide sequence, in the promoter region of the wheat DEMETER homoeologs, and the CRISPR/ Cas9 construct targets a 22-bp site in the wheat Dre2 homoeologs. In the donor construct, the DEMETER TALE repressor was cloned under the control of the maize endosperm-specific promoter of the EBE2 gene and a nopaline synthase (nos) terminator, whereas in the nuclease construct, the single guide RNA module was cloned under the control of the Rice snoRNA U3 promoter and the gene encoding Cas9 nuclease was cloned under the control of the P1 promoter.

4.1 Construction of dTALE Repressor (Donor) and CRISPR/ Cas9 Nuclease Constructs

In order to achieve endosperm specific silencing of the three homoeologous wheat DEMETER (DME) genes, a unique TALE repressor construct flanked on either side by the Dre2 sequences (the second target gene) was developed. A 16.5 repeat TALE array specifically designed to target a 17-nucleotide sequence, “TGCAGCTGGAGACTGGG” in the promoter region of the DME homoeologs was assembled following a stepwise procedure described in Cermak et al. [29]. The blueprint of the complete construct and the intermediate steps involved in its assembly are shown in Fig. 2. In this construct the DME TALE repressor is cloned under the control of an endosperm specific promoter of the maize EBE-2 gene and a nos terminator. As mentioned, the expression cassette of DME TALE repressor (pEBE2::DME-TALE-SRDX::Nos) was flanked on either side by the Dre2 specific sequences: Dre2-Left (852 bp) and Dre2-Right (708 bp). The purpose of introducing these sequences is to induce homologous recombination between the donor or DME TALE repressor construct and the target gene sequence. Amplification profiles of different DNA fragments used in the assembly of the

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DME TALE repressor construct pSL7-3 are shown in Fig. 3a. Intermediate plasmids obtained during the assembly process and the final plasmid, were validated using PCR amplification and restriction digestion followed by DNA sequencing (Clemson University Genomics Institute). Results of the PCR analyses and restriction digestions are shown in Fig. 3b. A second (nuclease) construct specifically targeting a conserved region of 20 nucleotides in the wheat Dre2 homoeologs was developed. The purpose of this construct is to introduce double stranded breaks (DSBs) in the wheat Dre2 homoeologs, which increases the possibility of homologous recombination-mediated DNA repair at this site in the genome. In order to develop a Dre2 specific CRISPR Cas9 construct, a plasmid dubbed pRGEB31 was procured from Addgene, a public plasmid repository and modified following the steps shown in Fig. 4. The new plasmid was confirmed to carry the guide RNA sequence of interest via restriction digestion followed by agarose gel electrophoresis and DNA sequencing. The validated plasmid was used to retransform E. coli strain DH5α to obtain the desired plasmid in large quantity. Then the purified plasmid was used to genetically transform hard red spring wheat cultivar WB926.

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Fig. 3 Restriction digestion and PCR amplification profiles of (a) DNA fragments used in the assembly of the DME TALE repressor construct, pSL7-3: (A) PCR amplification product of the EBE-2 promoter amplified from maize genomic DNA using specific primers [32]; (b) lane 2—PCR amplification product of Nos terminator amplified from plasmid 728 using specific primers (Wen et al. [26]), lane 3 and 4—PCR amplification products of the C-terminal and N-terminal ends of the TAL effector AvrHah1 of Xanthomonas gardneri, amplified from plasmid TAL2 using specific primers (Cermak et al. [29]). (c) PCR amplification products of SOE (splicing by overlap extension) reaction, lane 2—EBE2 and N-terminal end TALE fragment, and lane 3—C-terminal end TALE fragment and Nos. (d) Product of restriction digestion of pTAL2_17R13925 with BamHI. A 3.5 kb band consisting of 16.5 TALE repeats and parts of N- and C-terminal ends of TALE is visible. (e) PCR amplification products of the Dre2 gene. The left (Lane 2) and right (Lane 3) Dre2 fragments visible in the picture flanking the Cas9 endonuclease target site in the Dre2 genes. (B) Intermediate and final plasmids. (a) Lanes 2 and 3 show digestion products of pLig_Dre2-1 with KpnI/PacI and XbaI/PacI, respectively and Lanes 5 and 6 show products of the PCR amplification using Dre2-right and Dre2-left fragments specific primers. (b) Agarose gel showing products of restriction digestion of plasmid Lig2-7 with AatII (Lane 2) and PacI (Lane 3), and plasmid γ.TaDMC1as with PacI (lane 4). (c) Product of restriction digestion of an intermediate plasmid pSL14-1 with BamHI. (d) Products of restriction digestion of the final construct, pSL7-3 with PacI (Lane 2) and BamHI (Lane 3)

4.2 Identification of Candidate Transformants with Adduct Lesions in the Wheat Dre2 Gene Homoeologs

Out of 120 plants transferred to the greenhouse, 37 plants survived the transition (due to aphid invasion). Subsequently, DNA was extracted from the lyophilized leaf samples of these 37 putative wheat transformants. After extraction the DNA quantities were adjusted to 50 ng/μL using a spectrophotometer. Following this step DNA was used for PCR amplification with the gene-specific primers, flanking the target site (50 -ATCGAGCGCCAGCTACTCATGG-30 ) in the wheat Dre2 gene (Fig. 4a). The two overlapping

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Fig. 4 Diagrammatic representation of a stepwise modular cloning scheme followed to assemble the Dre2 specific CRISPR Cas9 expression cassette in the pUC19 backbone. (A) Two mother plasmids, pRGRB31 and pUC19 that respectively contribute to the guide RNA module plus Cas9 expression cassette and the plasmid

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sets of PCR primers (1) KpnI-Dre2-Left-F + Dre2-3-R and (2) Dre2-3-F + XbaI-Dre2-Rig-R (Fig. 5a) were used to amplify the genomic DNA in the two consecutive rounds of PCR. Product (1/10th dilution) of the first PCR reaction was used as template in the second round, and the product of second PCR reaction (1/10th dilution) was used as template in the third PCR set using a target specific primer (gRNA-F) and a template specific primer (XbaI-Dre2-Rig-R). Genotypes that failed to amplify in the latter reaction were tested with another set of primers: Dre2-3-F and gRNA-R to confirm lack of product in the former reaction. The rationale behind amplifying the wheat transformants with overlapping sets of primers is to increase specificity of amplification and that with the target and template specific primers is to identify mutation(s) induced by the Cas9 enzyme. Results of the PCR reaction suggested genotypes 9, 21, 24, 27 and 38 to carry mutations at the target site, as these lines failed to amplify products in both primer combinations mentioned above (Fig. 5b). Gliadins and glutenins were extracted from the grains collected from individual spikes and analyzed using polyacrylamide gel electrophoresis followed by densitometry. The preliminary analysis of seed storage proteins from candidate transformants suggested that plant number 24 and 38 are significantly reduced in their respective gliadin contents (Fig. 6).

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Fig. 6 Sodium dodecyl sulfate polyacrylamide gel electrophoresis and densitometric analysis of the gliadin samples derives from putative wheat transformants. First lane in the gel is the prestained protein ladder (Thermo Fisher cat.# 26612); WB926 ¼ untransformed control

5

Notes 1. Extended period of blending may cause damage to the microspores. 2. Density centrifugation allows separation of the nonembryogenic microspores from the embryogenic microspores; the latter accumulates at the interphase between the maltose and the mannitol solutions. 3. Adequate microspore density in microspore cultures reduces the competition for nutrients, oxygen, and space for cell divisions, which improves both the number and quality of embryoids [29]. 4. Use of nursing ovaries to condition medium increase the frequency of microspore embryogenesis by providing essential substance(s) “nurse factors” for embryogenesis [30]. 5. To ensure an adequate supply of “nurse factors,” aged ovaries in coculture may be replaced with fresh ones after 28 days [30]. 6. In vitro regenerated plants needs acclimation before transfer to soil (in the growth cabinet or the greenhouse) as sudden transfer of the plantlet from the highly humid and heterotrophic (low intensity light and medium containing sugars and nutrients) tissue culture conditions to soil under high intensity light causes severe stress leading to plant death. Therefore, when transferred to the soil plants should be kept under low light intensity and temperature, and covered with a transparent plastic dome to maintain high humidity. Direct transfer to the broad-spectrum sunlight and high temperature might cause charring of leaves and wilting of plantlets, leading to premature plant death [31].

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Acknowledgments This project is supported by the funding from the State of South Carolina grant SC-1700482, the Life Sciences Discovery Fund Grant 3143956-01 and the Clemson Faculty Succeeds Grant. References 1. Eversole K, Feuillet C, Mayer KFX, Rogers J (2014) Slicing the wheat genome. Science 345:285–287 2. King M, Altdorff D, Li P, Galagedara L, Holden J, Unc A (2018) Northward shift of the agricultural climate zone under 21stcentury global climate change. Sci Rep 8:7904 3. Olmstead AL, Rhode PW (2011) Adapting North American wheat production to climatic challenges, 1839–2009. Proc Natl Acad Sci U S A 108:480–485 4. Weeks DP, Yang B (2017) Genome editing in plants. Prog Mol Biol Transl Sci 149:1–249 5. The National Academies of Sciences, Engineering and Medicine (2016) Genetically engineered crops: Experiences and prospects. https://www.nap.edu/catalog/23395/geneti cally-engineered-cropsexperiences-andprospects 6. Voytas DF, Gao C (2014) Precision genome engineering and agriculture: opportunities and regulatory challenges. PLoS Biol 12:e1001877 7. Cunningham FJ, Goh NS, Demirer GS, Matos JL, Landry MP (2018) Nanoparticle-mediated delivery towards advancing plant genetic engineering. Trends Biotechnol 36:882–897 8. Songstad DD, Petolino JF, Voytas DF, Reichert NA (2017) Genome editing of plants. Crit Rev Plant Sci 36:1–23 9. Sauer NJ, Mozoruk J, Miller RB, Warburg ZJ, Walker KA, Beetham PR, Scho¨pke CR, Gocal GF (2016) Oligonucleotide-directed mutagenesis for precision gene editing. Plant Biotechnol J 14:496–502 10. Cardi T, Stewart CN Jr (2016) Progress of targeted genome modification approaches in higher plants. Plant Cell Rep 35:1401–1416 11. Bilichak A, Luu J, Eudes F (2015) Intracellular delivery of fluorescent protein into viable wheat microspores using cationic peptides. Front Plant Sci 6:666 12. Altpeter F, Springer NM, Bartley LE, Blechl AE, Brutnell TP, Citovsky V, Conrad LJ, Gelvin SB, Jackson DP, Kausch AP, Lemaux PG, Medford JI, Orozco-Ca´rdenas ML, Tricoli DM, Van Eck J, Voytas DF, Walbot V, Wang K, Zhang ZJ, Stewart CN Jr (2016)

Advancing crop transformation in the era of genome editing. Plant Cell 28:1510–1520 13. Bhalla PL, Singh MB (2017) Wheat biotechnology: methods and protocols. Humana Press, New York, NY 14. Zaidi SS, Mansoor S (2017) Viral vectors for plant genome engineering. Front Plant Sci 8:539 15. Chakraborty N, Chakraborty P, Sen M, Bandopadhyay R (2019) Choice of explant for plant genetic transformation. In: Rustgi S, Lup H (eds) Biolistic DNA delivery in plants. Springer, New York. (Chapter 5 this volume) 16. Chauhan H, Khurana P (2011) Use of doubled haploid technology for development of stable drought tolerant bread wheat (Triticum aestivum L.) transgenics. Plant Biotechnol J 9:408–417 17. Patnaik D, Khurana P (2001) Wheat biotechnology: a mini review. Electron J Biotechnol 4 (2):7–8. www.Ejb.Org/Content/Vol4/ Issue2/Full/4/ 18. Resch T, Touraev A (2011) Pollen transformation technology. In: Stewart NC, Touraev A, Citovsky V, Tzfira I (eds) Plant transformation technologies. Wiley-Blackwell, Chichester, pp 83–91 19. Brew-Appiah RAT, Ankrah N, Liu W, Konzak CF, von Wettstein D, Rustgi S (2013) Generation of doubled haploid transgenic wheat lines by microspore transformation. PLoS One 8: e80155 20. Rustgi S, Ankrah N, Brew-Appiah RAT, Sun Y, Liu W, von Wettstein D (2017) Doubled haploid transgenic wheat lines by microspore transformation. In: Bhalla PL, Singh MB (eds) Wheat biotechnology: methods and protocols. Humana Press, New York, NY, pp 213–234 21. Wan Y, Lemaux PC (1994) Generation of large numbers of independently transformed fertile barley plants. Plant Physiol 104:37–48 22. Zheng MY, Weng Y, Liu W, Konzak CF (2002) The effect of ovary-conditioned medium on microspore embryogenesis in common wheat (Triticum aestivum L.). Plant Cell Rep 20:802–807

Wheat Microspore-Derived Calli as Explants 23. Osorio C, Wen N, Mejias JH, Liu B, Reinbothe S, von Wettstein D, Rustgi S (2019) Development of wheat genotypes expressing a glutamine-specific endoprotease from barley and a prolyl endopeptidase from Flavobacterium meningosepticum or Pyrococcus furiosus as a potential remedy to celiac disease. Funct Integr Genomics 19:123–136 24. Rustgi S, Wen N, Osorio C, Brew-Appiah RAT, Wen S, Gemini R, Mejias JH, Ankrah N, Moehs CP, von Wettstein D (2014) Natural dietary therapies for the ‘gluten syndrome’. The Royal Danish Academy of Sciences and Letters, Copenhagen, Denmark 25. Osorio C, Wen N, Gemini R, Zemetra R, von Wettstein D, Rustgi S (2012) Targeted modification of wheat grain protein to reduce the content of celiac causing epitopes. Funct Integr Genomics 12:417–438 26. Wen S, Wen N, Pang J, Langen G, BrewAppiah RAT, Mejias JH, Osorio C, Yang MM, Gemini R, Moehs CP, Zemetra RS, Kogel KH, Liu B, Wang X, von Wettstein D, Rustgi S (2012) Structural genes of wheat and barley 5-methylcytosine DNA glycosylases and their potential applications for human health. Proc Natl Acad Sci U S A 109:20543–20548 27. Rustgi S, Shewry P, Brouns F (2020) Health hazards associated with wheat and gluten consumption in susceptible individuals and status of research on dietary therapies. In: Igrejas G, Ikeda TM, Guzma´n C (ed) Wheat quality for

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improving processing and human health. Springer, Cham, pp 471–515 28. Rustgi S, Shewry P, Brouns F, Deleu L, Delcour JA (2019) Wheat seed proteins—factors influencing their content, composition, and technological properties, and strategies to reduce adverse reactions. Comp Rev Food Sci Food Safety 18:1751–1769 29. Cermak T, Doyle EL, Christian M, Wang L, Zhang Y, Schmidt C, Baller JA, Somia NV, Bogdanove AJ, Voytas DF (2011) Efficient design and assembly of custom TALEN and other TAL effector-based constructs for DNA targeting. Nucleic Acids Res 39:e82 30. Zheng MY (2003) Microspore culture in wheat (Triticum aestivum)—doubled haploid production via induced embryogenesis. Plant Cell Tissue Organ Cult 73:213–230 31. Chandra S, Bandopadhyay R, Kumar V, Chandra R (2010) Acclimatization of tissue cultured plantlets: from laboratory to land. Biotechnol Lett 32:1199–1205 32. Magnard J, Lehouque G, Massonneau A, Frangne N, Heckel T, Gutierrez-Marcos J, Perez P, Dumas C, Rogowsky P (2003) ZmEBE genes show a novel, continuous expression pattern in the central cell before fertilization and in specific domains of the resulting endosperm after fertilization. Plant Mol Biol 53:821–836

Chapter 15 Plant Transformation Techniques: Agrobacteriumand Microparticle-Mediated Gene Transfer in Cereal Plants Jafargholi Imani and Karl-Heinz Kogel Abstract Biotechnological methods for targeted gene transfers into plants are key for successful breeding in the twenty-first century and thus essential for the survival of humanity. Two decades ago, genetic transformation of crop plants was not routine, and it was all but impossible with important cereals such as barley and wheat. The recent focus on crop plant genomics—yet based on the Arabidopsis toolbox—boosted the research for more efficient plant transformation protocols, thereby considerably widened the number of genetically tractable crops. Moreover, modern genome editing methods such as the CRISPR/Cas technique are game changers in plant breeding, though heavily dependent on technical optimization of plant transformation. Basically, there are two successful ways of introducing DNA into plant cells: one is making use of a living DNA vector, namely, microbes such as the soil bacterium Agrobacterium tumefaciens that infects plants and naturally transfers and subsequently integrates DNA into the plant genome. The other method uses a direct physical transfer of DNA by means of microinjection, microprojectile bombardment, or polymers such as polyethylene glycol. Both ways subsequently require sophisticated strategies for selecting and multiplying the transformed cells under tissue culture conditions to develop into a fully functional plant with the new desirable characteristics. Here we discuss practical and theoretical aspects of cereal crop plant transformation by Agrobacterium-mediated transformation and microparticle bombardment. Using immature embryos as explants, the efficiency of cereal transformation is compelling, reaching today up to 80% transformation efficiency. Key words Agrobacterium, Barley, Gene transfer, Microparticle bombardment, Wheat

1

Introduction In 1974, Marc Van Montagu and Jeff Schell at Ghent University showed that Agrobacterium tumefaciens uses plasmids, known as the tumor-inducing (Ti) plasmids, to transfer its genetic material into a plant cell. About 10 years later, three groups published research papers demonstrating that they could stably introduce and express bacterial genes into a plant genome using A. tumefaciens as a vector [1–3]. It was not only that A. tumefaciens could insert bacterial genes reliably into tobacco plants, but that the inserted genes were inherited and expressed by

Sachin Rustgi and Hong Luo (eds.), Biolistic DNA Delivery in Plants: Methods and Protocols, Methods in Molecular Biology, vol. 2124, https://doi.org/10.1007/978-1-0716-0356-7_15, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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subsequent generations of the plants. Interestingly, first attempts into plant transformation in the late 1970s were focused on using protoplasts—plant cells with their rigid cellulose cell walls removed. The discovery that A. tumefaciens could transfer its DNA into plants, plant scientists altered the focus of the transformation strategies. Today, plant transformation by A. tumefaciens is a routine method that underpins many of the techniques used in plant molecular biology. One of the first practical applications, which resulted in commercialization, was the use of A. tumefaciens to introduce to tomato (Solanum lycopersicum) plant cells “antisense” versions of a gene being expressed during tomato ripening. The antisense approach is one form of “reverse genetics.” Unlike forward genetics, where researchers seek to understand the genetic changes responsible for an observed mutation, reverse genetics begins with a known gene and allows researchers to identify the function of that gene. For instance, when a plant is transformed, the T-DNA is inserted essentially at random into the genome. If it is inserted anywhere within an existing gene, it can knock out (KO) that gene and create what is known as a loss of function mutant. The randomness can be exploited by transforming plant cells with a small piece of T-DNA to create large libraries of plants, which each carry a single gene KO. This library can be screened looking for mutations of interest and, as the sequence of the T-DNA is known, we can identify and clone the gene into which it has been inserted [4, 5]. 1.1 State-of-the-Art Transformation Techniques in Cereals

Genetic transformation mediated by A. tumefaciens has become the gold standard for plant genetic engineering. A. tumefaciens naturally infects wound sites in the dicotyledonous plants, causing the pathogenic development of crown galls. A large Ti plasmid of >200 kb in size plays a key role in gall induction (cf. Ref. 6 and Chapter 3 in this volume for more details). The biotechnological adaptation of Agrobacterium-mediated gene transfer to monocotyledonous species, such as rice [7], maize [8], barley [9], wheat [10, 11], and rye [12], as well as to fungi [13], yeasts [14], and protists, such as algae [15], and animal cells [16] makes it a universal tool both for functional genetic studies and for problem-solving applications in agriculture. Important advantages of Agrobacterium-mediated transformation techniques are a high frequency of gene transfer, proper integration of the foreign gene into the host genome, and low copy number of the inserted transgene. Natural infections of plant hosts by A. tumefaciens follows a default program: (1) perceiving plants’ wound molecules such as the phenolic acetosyringone (3,5-Dimethoxy-4-hydroxyacetophenon), (2) colonizing the plants’ extracellular space, (3) Activating the bacterial virulence genes, (4) assembling the T-DNA transfer complex, and (5) integrating the T-DNA complex into the plant genome. Modern transformation protocols follow this natural strategy of the

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Fig. 1 Vector maps for plant cell transformation. (a) Map of the super helper plasmid (pBS1) with a tetracycline-resistance (TC) gene. Abbreviations: ORI, origin of replication COS, cos site of phage virC, virB, virC, virG genes from A. tumefaciens strain A281pTiBo542. pSB1 introduced to A. tumefaciens [24] by triparental mating [25]. (b) Map of pLH6000 35s:GFP binary vector. (c) Map of pWBVec10aUbi-int-GUSbinary vector. Abbreviations: pUbi-int: Maize (Zea mays) Ubiquitin 1 promoter containing the first intron in front of the start codon. GFP: Sequences coding Green fluorescence protein as a reporter gene. GUS Sequences encoding β-glucuronidase as a reporter protein, Nos-T Terminator from nopaline synthase gene, p35S Promoter driven by the Cauliflower mosaic virus, Hpt hygromycin phosphotransferase (hpt) gene as a selectable marker gene, BR right border sequences, BL left Border sequences, Sm/Sp or SpecR Bacterial antibiotic resistance genes streptomycin/spectinomycin

bacterium to integrate transgenes into the plant genome. Successful protocols are specifically adapted for the plant species, and within a species for a cultivar. The optimization of transformation protocols requires optimization of many working steps, including producing competent cells from tissues and regenerating cells to a healthy plant after transformation [17]. Moreover, Agrobacteriummediated transformation of monocotyledonous plants requires the use of “superbinary” vector systems, that is, binary vectors carrying a DNA fragment from the A. tumefaciens virulence region (Fig. 1). Using highly virulent strains of A. tumefaciens and improved vectors boosted the transformation efficiencies in cereals. A critical criterion for achieving explant development and high transformation efficiency is the light quality, as it influences synchronized development of the recipient plants’ spikes. Especially, red light affects phytochrome reversibility and is most important for flowering [18, 19] and the development of caryopses [20]. Caryopsis explant tissue or embryogenic calli derived from such tissues serve as starting material for transformation of monocotyledonous plants [21], while in contrast leaves, stems, hypocotyls, or petioles serve as starting material for transformation of dicotyledonous plants [22]. Here we describe state of the art in cereal transformation using barley and wheat as examples. Despite considerable progress in the transformation of cereals that methods remains rather timeconsuming and cost-intensive.

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Agrobacterium-Mediated Transformation For barley genetic transformation, we describe here an Agrobacterium-mediated method. The purpose was to compare two genetic transformation methods used in two related cereals.

2.1

Materials

1. Na-hypochlorite solution (20%). 2. Tween 20. 3. Ethanol (70% v/v). 4. Deionized H2O (pH 3). 5. Petri dishes (9 cm in diameter). 6. Barley spikes (~2 weeks after anthesis). 7. Magnetic stirrer. 8. Barley callus induction medium (BCIM). 9. Barley shoot induction medium (BSIM). 10. Barley root induction medium (BRIM). 11. YEB (Yeast Extract Beef)-medium for A. tumefaciens culture. 12. Binocular microscope. 13. Sterilized preparation lancet. 14. Sterilized forceps. 15. 3:1 mixture of vermiculite (diameter 0–3 mm). 16. Small pots (diameter 10 cm). 17. Binary vector for expression of β-Glucuronidase gene (GUS). 18. Binary vector for expression of Green fluorescent protein gene (GFP). 19. Gene Pulser apparatus (Bio-Rad). 20. Incubator. 21. Plant growth chamber.

2.2 Plant Material and Growth Conditions

Spring barley (Hordeum vulgare L.) cultivar Golden Promise is routinely used for barley transformation. Pots of 3 L volume are filled with soil type LD80 (Fruhstorfer, Germany) (see Note 1). Three barley seeds are sown in each pot and cultivated in an environmentally controlled plant growth cabinet at 18
C/14
C (day/night cycle) with 60% relative humidity (rH) and a photoperiod of 16 h (240 μmol m2 s1 photon flux density) (see Note 2). Floral transitions and reproductive development in cereals has a major effect on yield. Different light spectra have various effects on plants. Some spectra stimulate vegetative growth, while others increase the yield in flowers and fruits. Monostori et al. [23] showed that flowering time was delayed, when a “Blue regimen”

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1,0 0,9

Relative Intensity

0,8 0,7 0,6 0,5 0,4 0,3 0,2 0,1 0,0 380

420

460

500

540

580 620 Wavelength

660

700

740

780

Fig. 2 Comparison of light spectra of LED (VALOYA wide-spectra NS1, Finland; blue) and Osram HQI-B-T (plus light bulb; red) in a plant growth chamber. The red-light intensity corresponding to the wavelength of 570 and 610 nm is higher in the normal light Osram HQI-B-T + light bulb than by LED. The red light is most important for flowering and fruiting regulation. Far-red (730 nm) seems to be very beneficial (red curve)

was applied (Fig. 2). In spite of the fact that the “Blue regimen” stimulated stomatal opening, the CO2 assimilation capacity of flag leaves remained low. Furthermore, LED produces discontinuous light spectra, thus missing color components, which is why not all colors can be reproduced correctly. In contrast, spectra of sun and light bulb are continuous and all colors of visible light are reproduced. This phenomenon could be the reason for malformation of barley kernel development under light-emitting diode (LED) regimes (Fig. 3a). Thus, the light spectrum under which recipient plants are grown is crucial for flowering stimulation and synchronized formation of caryopses as starting material for transformation. To increase the IR intensity, four light bulbs were additionally installed per square meter in the plant growth chamber. For reliable automated watering, we used the Drip Irrigation Fully Automatic Watering System (Galcon, Germany) with the following program (see Note 3): 0–2-week-old plants: every 2 days 100 mL. 2–4-week-old plants: daily 130 mL. 4–6-week-old plants: daily 260 mL. 6–8-week-old plants: daily 200 mL. 8–10-week-old plants: daily 100 mL. 2.3 Preparation of A. tumefaciens Electrocompetent Cells

Before the pulse, bacterial cells are grown in YEB medium (yeast extract 1 g/L; beef extract 5 g/L; sucrose 5 g/L; casein hydrolysate 5 g/L; MgCl2 2 mM) till a density of OD600 ¼ 0.7–1.0. The

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Fig. 3 Barley spikes grown under different light spectra. (a) Kernel development under Osram HQI-B-T lamp (left) and under LED (VALOYA wide-spectra NS1, Finland) (right). (b) Immature scutella tissues of barley cultivar Golden Promise isolated from A (left) kernel. The tissues were isolated from the caryopses and the embryos were removed (arrows). (c) Confocal laser scanning microscopy of ZmUbi1:GFP barley scutella tissue 2 days after transformation. (d) GFP expressing stably transformed barley plant derived from transformed scutella tissue

bacteria are pelleted by centrifugation (3113 RCF, 10 min at 4
C), and the pellet is washed (1) with 10% sterilized glycerol (v/v) in the same volume as the above suspension and repelleted by centrifugation, followed by washing the pellet with 1/2 of the above volume of bacteria suspension and pelleting by centrifugation.

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Finally, the pellet is resuspended in 1 mL of 10% glycerol and kept at 85
C. For vector introduction, 10–100 ng purified binary vector pLH6000 Ubi-int:GFP (GFP as reporter gene; see Fig. 1b) is introduced into 50 μL electrocompetent A. tumefaciens strain AGL1 [23] cells by electroporation using the preexisting protocol on Gene Pulser Xcell™ unit (Bio-Rad, Germany). Bacterial cells are cultured for 1 h at 28
C without antibiotics to allow expression of the antibiotic resistance gene. Subsequently, A. tumefaciens containing the vector is suspended in 20 mL liquid YEB medium and cultured overnight under 28
C and gentle shaking with an orbital shaker at 150 rpm. It is important to assure a temperature of 28
C in order to avoid release of the disarmed Ti plasmid from the bacteria by heat stress. The following antibiotics are added to the overnight culture: rifampicin (25 mg/L) for selection of the A. tumefaciens strain; carbenicillin (25 mg/L) for selection of the disarmed Ti plasmid (helper plasmid); and spectinomycin (50 mg/L) for selection of the binary vector pLH6000 Ubi-int:GFP. 2.4 Surface Sterilization of Immature Barley Seed

About 200 kernels of a few spikes (10–14 days after anthesis) are collected in a flask, incubated in 70% ethanol for 3 min and 20% Na-hypochlorite (with one drop of tween 20) under stirring for 20 min. Subsequently, the seeds are rinsed 1 with deionized H2O (adjusted with 1 N HCl to pH 3), followed 3 with sterile deionized water, and finally transferred to petri dishes (see Note 4).

2.5 Isolation of Immature Embryos and Inoculation with A. tumefaciens

Aseptic scutella tissue is excised from immature zygotic embryos (1.5–2.5 mm) using forceps and a preparation lancet and immediately placed onto callus induction medium (BCID, Table 1) with the scutellum upside down (Fig. 3b). About 25–30 scutella are collected and 200 μL A. tumefaciens overnight culture is dropwise added to embryos. Embryos are turned upside and cultivated at 24
C for 40–60 min in the dark. Subsequently, ca. 10–12 embryos are transferred to the BCIM plate (Table 1) and cocultivate at 24
C in the dark. Two days later, the embryos are placed onto BCID medium with antibiotics ticarcillin (150 mg/L) and hygromycin B (50 mg/L).

2.6 Generation of Plants from Calli

After infection, callus development is induced on barley shoot induction medium (BSIM, Table 1) supplemented with containing 50 mg/L hygromycin B as a selective agent for the transformed plant cells. Established calli are sub-cultured on regeneration medium (BRIM, Table 1) supplemented with 25 mg/L hygromycin B until rooted plantlets can be transferred to soil. The antibiotic ticarcillin (150 mg/L) is applied in all culture nutrient media as long as tests for the presence of bacteria are negative.

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Table 1 Nutrient media compositions 1L

BCIM

BSIM

BRIM

WCIM

WOM

WSIM

WRIM

MS-stock (Duchefa M0221)

4.3 g

–

2.15 g

4.3 g

4.3 g

4.3 g

4.3 g

MS-stock (Duchefa M0238) NH4NO3 free)

–

2.7 mg –

–

–

–

–

CuSO4·5H2O

1.2 mg 1.2 mg 0.6 mg –

–

–

–

NH4NO3

–

165 mg –

–

–

–

–

Mannitol

–

–

–

–

75 g

Maltose

30 g

62 g

15 g

–

–

–

–

Sucrose

–

–

–

40 g

40 g

20 g

20 g

Thiamine–HCl

1 mg

0.4 mg 0.5 mg –

–

Myoinositol

250 mg 100 mg 125 mg –

–

Glutamine

–

150 mg –

Casein hydrolysate

1g

–

L-Proline

690 mg –

345 mg –

–

Dicamba

2.5 mg –

–

–

–

–

–

BAP (benzylamine purine)

–

1 mg

–

–

–

1 mg

–

2,4-D

–

–

–

2 mg

2 mg

–

–

Phytoagar

5g

5g

5g

–

–

–

–

pH

5.9

5.6

5.6

5.8

5.8

5.8

5.8

Gelrite

–

–

–

4g

4g

4g

4g

–

– –

–

500 mg 500 mg –

–

500 mg 100 mg 100 mg 100 mg 100 mg –

BCIM Barley callus induction medium, BSIM Barley shoot induction medium, BRIM Barley root induction medium, WCIM Wheat callus induction medium, WOM Wheat osmotic medium, WSIM Wheat shoot induction medium, WRIM Wheat root induction medium

2.7 Determination of GFP in Transformed Tissue

3

GFP expression is detected in stably transformed plant material with a standard fluorescence microscope (excitation: laser line 488 nm; emission: 500–540 nm).

Biolistic Transformation For wheat transformation, we describe here a biolistic method, which can be used as an alternative to Agrobacterium-mediated transformation.

3.1

Materials

1. Wheat spikes for isolation of immature embryos. 2. Ethanol (70%; v/v) and Na-hypochloride (20%; v/v) for surface sterilization.

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3. H2O (adjusted with 1 N HCl to pH 3). 4. Gold particles (0.6–1 μm). 5. Sonicator. 6. Ethanol (100%) for precipitation of DNA to gold particles. 7. Sterilized macrocarrier holders, macrocarriers, screens, and rupture disks.

stopping

8. Wheat callus induction medium (WCIM). 9. Wheat osmotic medium (WOM). 10. Wheat shoot induction medium (WSIM). 11. Wheat root induction medium (WRIM). 12. Gen Gun machine. 13. Helium gas. 14. X-Gluc (5-bromo-4-chloro-3-indolyl-β-D-glucuronic acid, cyclohexyl ammonium salt) for histochemical GUS detection. 3.2 Collection and Sterilization of Wheat Caryopses

Wheat (Triticum aestivum L. var. Bobwhite) plants are grown in an environmentally controlled plant growth cabinet with the same culture condition described above for barley. The first step in the transformation of wheat is the isolation of immature embryos. Spikes are collected from plants grown in a tightly controlled growth cabinet. Embryos at the correct stage usually are found 12–14 days post anthesis. The caryopses are removed and sterilized by rinsing in 70% ethanol for 5 min and soaking in 20% Na-hypochlorite for 20 min under gentle shaking. Subsequently, the caryopses are rinsed 4 with sterile water.

3.3 Isolation of Immature Wheat Embryos

Embryos are isolated with the help of a microscope under a laminar flow hood; unlike barley, it is not necessary to remove the embryo axis. Embryos of 0.5–1.5 mm in size are generally most responsive. Thirty immature embryos are placed on the wheat callus induction (WCIM) medium without selective agents for 5 days. The callus tissue is placed on osmotic media (WOM, Table 1) in a circle on petri plates that corresponds to the area that will be bombarded with DNA coated gold particles and is kept there for 20 h following the shooting. The callus tissue is then placed on a WCIM containing hygromycin B (50 mg/L) for 3 weeks. Established calli are sub-cultured on shoot induction medium (WSIM, Table 1) and 4 weeks later on regeneration medium (WRIM, Table 1) supplemented with 25 mg/L hygromycin B until rooted plantlets can be transferred to soil.

3.4 Coating of Gold Particles with DNA (for Six Shots)

Twenty mg of gold particles (0.6–1.0 μm in diameter), in 1 mL 100% ethanol are mixed in a 1.5 mL Eppendorf tube, sonicated for 2 min, and pulse-spun in a microfuge for 3 s. The ethanol wash is repeated twice. Subsequently, 1 mL sterile water is added to the

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pellet, sonicated for 2 min, and pulse spun in a microfuge for 3 s. This step is repeated twice. Finally, gold particles in 1 mL sterile water are split into 50 μL aliquots and 5 μL of vector pWB10a Ubi: GUS (1 μg/μL; Fig. 1c) is added, followed by vortexing. Then 50 μL of 2.5 M CaCl2 and 20 μL 0.1 M spermidine is added followed by vortexing. The DNA-coated particles are then pelleted by centrifugation. The particles are washed in 150 μL of 100% ethanol, and the pellet is resuspended in 85 μL of 100% ethanol and kept on ice. 3.5 Delivery of DNA-Coated Gold Particles

The following settings are recommended for this procedure (Fig. 4): (1) sterilize the gun’s chamber and component parts with 70% (v/v) ethanol; (2) sterilize macrocarrier holders, macrocarriers, stopping screens and rupture disks by dipping in 100% ethanol and allow to evaporate completely; (3) briefly vortex the coated gold particles, take 5 μL, place them centrally onto the macrocarrier membrane and allow drying; (4) place a stopping screen into the fixed nest. Invert the macrocarrier holder containing macrocarrier plus gold particles/DNA and place over the stopping screen in the nest and maintain its position using the retaining ring; (5) mount the fixed nest assembly onto the second shelf from the top to give a gap of 2.5 cm; (6) place a sample on the target stage (target distance 5.5 cm); and (6) draw a vacuum of 27–28 in. Hg and fire the gun.

Fig. 4 Transformation scheme for particle bombardment

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3.6 Histochemical Assays for GUS Detection

291

The E. coli ß glucuronidase (GUS) is one of the most popular reporter proteins currently used. The protein has a molecular weight of 68,200 Da and is very stable. It may be assayed at any physiological pH, with an optimum between 5.2 and 8.0. The GUS gene can be used in a gene fusion; this means that the GUS coding sequence is under the direction of the controlling sequence of a gene of interest. The best substrate currently available for histochemical localization of ß-glucuronidase activity in tissues and cells is 5-bromo-4chloro-3-indolyl glucuronide (X-Gluc). The substrate works very well, giving a blue precipitate at the site of enzyme activity. The indoxyl derivative produced must undergo an oxidative dimerization to form the insoluble and highly colored indigo dye. This dimerization is stimulated by atmospheric oxygen and can be enhanced by using an oxidation catalyst such as a K+ ferricyanide– ferrocyanide mixture. Whole tissues, callus, suspension culture cells, protoplasts, whole plants, or plant organs can be stained, but the survival of the stained cells is not certain. After staining, clearing the tissue with 70% ethanol enhances the contrast in many cases. The method of GUS staining is elaborated below. 1. Transfer the bombarded tissue in the histochemical reagent (X-Gluc, Table 2) in a microfuge tube and incubate overnight at 37
C.

Table 2 Composition of X-Gluc solution 100 mL X-GlcA (Duchefa, Holland)

100 mg

N,N-Dimethyl-formamide (Sigma)

15–20 drops

0.1 M phosphate buffer (pH 7.0)

98 mL

5 mM potassium ferricyanide (Sigma)

1 mL

5 mM potassium ferrocyanide (Sigma)

1 mL

Triton X-100

100 μL

Filter sterilized and stored at 20
C Preparation of phosphate buffer (0.1 M) for X-Gluc solution 0.2 M KH2PO4

87 mL

0.2 M K2HPO4

122 mL

Deionized water

209 mL

Total

418 mL

Adjust pH of 7.0

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Fig. 5 GUS activities in wheat immature embryos. Immature embryo was bombarded with pWB10a-Ubi:GUS (Fig. 1c) constructs. The blue spots indicate the transformation efficiency

2. After staining, rinse the sample in 70% ethanol (v/v) for at least 5 min. 3. Examine for GUS stain under the microscope (Fig. 5).

4

Notes 1. This standardized soil mixture contains high-quality volcanic clay and peat (70% peat and 30% clay). Nutrients are readily absorbed into the clay. The buffering effect of volcanic clay protects against damage from fluctuations in the pH-range 6 and from the nutrient supply. Volcanic clay is stable as a topsoil, constantly delivers trace elements, and gives the cultivation substrate the necessary richness. The Type LD80 soil contains fertilizer (4 kg/m2). Therefore, there is no need for any additional fertilizer during the whole vegetation period. 2. Cultivation at low temperature causes a synchronous growth and reduction of disease. The use of fungicides may adversely affect cell growth or agro-transformation. 3. Water deficiencies and lack of oxygen in the tissue can occur especially in the case of waterlogging in the soil. In particular, barley is very sensitive to waterlogging. To prevent waterlogging in the soil, the plants should be watered with defined amounts of water during growth. 4. The seed surface sterilization agent Na-hypochlorite has a pH value >11. Therefore, it is recommended to normalize the pH value by 1 washing with water (pH 3).

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Acknowledgments We thank E. Swidtschenko and C. Dechert for excellent technical assistance. References 1. Fraley RT, Rogers SG, Horsch RB, Sanders PR, Flick JS, Adams SP, Bittner ML, Brand LA, Fink CL, Fry JS, Galluppi GR, Goldberg SB, Hoffmann NL, Woo SC (1983) Expression of bacterial genes in plant cells. Proc Natl Acad Sci U S A 80:4803–4807 2. Herrera-Estrella L, Depicker A, Van Montagu M, Schell J (1983) Expression of chimaeric genes transferred into plant cells using a Ti-plasmid-derived vector. Nature 303:209–213 3. Bevan M, Flavell R, Chilton M (1983) A chimaeric antibiotic resistance gene as a selectable marker for plant cell transformation. Nature 304:184–187 4. Koncz C, Martini N, Mayerhofer R, KonczKalman KH, Redei GP, Schell J (1989) Highfrequency T-DNA-mediated gene tagging in plants. Proc Natl Acad Sci U S A 86:8467–8471 5. Feldmann KA (1991) T-DNA insertion mutagenesis in Arabidopsis: mutational spectrum. Plant J 1:71–82 6. Gordon JE, Christie PJ (2014) The Agrobacterium Ti Plasmids. Microbiol Spectr 2(6). https://doi.org/10.1128/microbiolspec. PLAS-0010-2013 7. Raineri DM, Bottino P, Gordon MP, Nester EW (1990) Agrobacterium-mediated transformation of rice (Oryza sativa). Bio-Technol 8:33–38 8. Ishida Y, Saito H, Ohta S, Hiei H, Komari T, Kumashiro T (1996) High efficiency transformation of maize (Zea mays L) mediated by Agrobacterium tumefaciens. Nat Biotechnol 14:745–750 9. Tingay S, McElroy D, Kalla R, Fieg S, Wang M, Thornton S, Brettell R (1997) Agrobacterium tumefaciens-mediated barley transformation. Plant J 11:1369–1376 10. Mooney PA, Goodwin PB, Dennis ES, Llewellyn DJ (1991) Agrobacterium tumefaciens -gene transfer into wheat tissues. Plant Cell Tissue Organ Culture 25:209–218 11. Cheng M, Fry JE, Pang S, Zhou H, Hironaka CM, Duncan DR, Conner TW, Wan Y (1997) Plant Physiol 115:971–980

12. Popelka JC, Altpeter F (2003) Agrobacterium tumefaciens-mediated genetic transformation of rye (Secale cereale L.). Mol Breed 11:203–211 13. de Groot MJ, Bundock AP, Hooykaas PJJ, Beijersbergen AGM (1998) Agrobacterium tumefaciens-mediated transformation of filamentous fungi. Nat Biotechnol 16:839–842 14. Bundock P, den Dulk-Ras A, Beijersbergen A, Hooykaas PJ (1995) Trans-kingdom T-DNA transfer from Agrobacterium tumefaciens to Saccharomyces cerevisiae. EMBO J 14:3206–3214 15. Cheng R, Ma R, Li K, Rong H, Lin X, Wang Z, Yang S, Ma Y (2012) Agrobacterium tumefaciens mediated transformation of marine microalgae Schizochytrium. Microbiol Res 167:179–186 16. Kunik T, Tzfira T, Kapulnik Y, Gafni Y, Dingwall C, Citovsky V (2001) Genetic transformation of HeLa cells by Agrobacterium. Proc Natl Acad Sci U S A 98:1871–1876 17. Keshavareddy G, Kumar ARV, Vemanna SR (2018) Methods of plant transformation—a review. Int J Curr Microbiol App Sci 7:2656–2668 18. Zhou Y, Singh BR (2002) Red light stimulates flowering and anthocyanin biosynthesis in American cranberry. Plant Growth Regul 38:165–171 19. Ugarte CC, Trupkin SA, Ghiglione H, Slafer G, Casal JJ (2010) Low red/far-red ratios delay spike and stem growth in wheat. J Exp Bot 61:3151–3162 20. Imani J, Li L, Sch€afer P, Kogel KH (2011) STARTS—a stable root transformation system for rapid functional analyses of proteins of the monocot model plant barley. Plant J 67:726–735 21. Imani J, Berting A, Nitsche S, Sch€afer S, Gerlich WH, Neumann KH (2002) The integration of a major hepatitis B virus gene into cellcycle synchronized carrot cell suspension cultures and its expression in regenerated carrot plants. Plant Cell Tissue Organ Culture 71:157–164

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22. Lazo GR, Stein PA, Ludwig RA (1991) A DNA transformation-competent Arabidopsis genomic library in Agrobacterium. Biotechnology 9:963–967 23. Monostori I, Heilmann M, Kocsy G, Rakszegi M, Ahres M, Altenbach SB, Szalai G, Pa´l M, Toldi D, Simon-Sarkadi L, Harnos N, Galiba G, Darko E´ (2018) LED lighting—modification of growth, metabolism, yield and flour composition in wheat by spectral quality and intensity. Front Plant Sci 9:605

24. Hoekema A, Hirsch PR, Hooykaas PJJ, Schilperoort RA (1983) A binary plant vector strategy based on separation of vir- and T-region of the Agrobacterium tumefaciens Ti-plasmid. Nature 303:179–180 25. Ditta G, Stanfield S, Corbin D, Helinski DR (1980) Broad host range DNA cloning system for Gram-negative bacteria: construction of a gene bank of Rhizobium meliloti. Proc Natl Acad Sci U S A 77:7347–7351

Chapter 16 Proteolistics: A Protein Delivery Method Susana Martin-Ortigosa and Kan Wang Abstract Intracellular protein delivery in plant tissues is becoming an important tool for addressing both basic and applied research questions by plant biologists, especially in the era of genome editing. The ability to deliver proteins or protein/RNA complexes into cells allows for producing gene-edited plants that are free of transgene integration in the genome. Here we describe a protocol for the delivery of a protein/gold particle mixture in plant cells through biolistics. The key for the delivery is the drying of the protein/gold suspension directly onto the gene-gun cartridge or macrocarrier. The intracellular protein delivery into plant cells is achieved through the bombardment using the Bio-Rad PDS-1000/He particle delivery device. We termed this methodology “proteolistics.” Key words Biolistics, Gene-gun, Intracellular delivery, Proteolistics, Protein delivery

1

Introduction In this chapter, we present a technology termed “proteolistics,” in which the intracellular delivery of proteins, chemicals, and biomolecule combinations can be achieved through bombardment in both plant and animal tissues [1]. This method can expand the biotechnological toolbox for the plant researchers. For instance, the intracellular delivery of a protein with modifications that cannot be easily achieved by DNA expression in the plant cell could facilitate its study. Another example is that researchers are applying protein delivery to improve genetic transformation through transposome complexes [2, 3] or to develop transient kanamycin resistance for plant selection by the delivery of neomycin phosphotransferase II protein using leaf infiltration [4]. However, the main biotechnological application of protein delivery is the generation of genetically modified plants using the DNA-free methodology. The delivery of, for example, recombinases [5–7], or CRISPR/Cas9 ribonucleoproteins [8–12] has opened new opportunities to engineer the plant genome without a need of integration and expression of a nucleases gene. The advantages of such approaches include

Sachin Rustgi and Hong Luo (eds.), Biolistic DNA Delivery in Plants: Methods and Protocols, Methods in Molecular Biology, vol. 2124, https://doi.org/10.1007/978-1-0716-0356-7_16, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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avoiding random integration of the foreign DNA, diminished off-target effects due to reducing prolonged/continuous expression of the genome editing reagents and the possibility of circumvention of genetically modified plants regulations, among others. Compared to DNA delivery, protein delivery is a transient approach. The effect of the protein will depend on its stability in the cell, which could last seconds to a few days. Another important aspect is that the delivery of DNA is universal and well-studied: protocols for its purification and delivery are available. On the contrary, protein delivery can be more challenging because each protein has its unique set of characteristics such as size, tertiary structure, stability, solubility, and isoelectric point as well as other physical and chemical parameters. These unique properties make the development of protein delivery methods more difficult because their usability and efficiency will directly depend on the nature of the protein to be delivered. Another key factor is that each protein needs an optimized purification and solubilization protocol, which could make the approach more expensive and timeconsuming. One major challenge in delivering proteins into plant cells is the presence of the plant cell walls. To circumvent this barrier, polyethylene glycol (PEG) mediated protoplast transformation has been used to deliver proteins to plant cells [11, 12]. There are a few other methods published describing the delivery of proteins to intact plant cells. Some of the protocols require self-made equipment like multi-gas plasma jets [13], specialized equipment and training like microinjection [14, 15] or engineering Agrobacterium to translocate proteins via its transport system [7]. Leaf infiltration has also been used to deliver proteins complexed with nanoparticles or fusion peptides [4, 16, 17]. Apart from these methodologies, there are two that have been more widely used. The first one is protein transduction. There are several reports that use protein transduction domains or cellpenetrating peptides to facilitate intracellular delivery of proteins in different plant tissues [18–23]. The covalent or noncovalent transduction can be done by simple incubation of protoplasts or intact plant tissues with the transduction peptides or proteins and the protein of interest or other biomolecules. There are some factors affecting the efficiency of this method, such as type of peptide used, ratio to the protein of interest, type, and age of cell or tissue and size of the cargo protein among others (reviewed in [24]). The second methodology is biolistics. For decades, the use of biolistics has been focused on the delivery of DNA to plant cells. There are some studies in which bombardment has been used to deliver other types of cargoes to plant cells, such as nanoparticles

Proteolistics: A Protein Delivery Method

297

[25–27], Agrobacterium cells [28, 29] or indicator dyes [30]. In the latter study, the dye/microprojectile mixture was done in ethanol and let to evaporate once placed onto the gene-gun cartridge. Algal and plant cells were then bombarded using the dried mixture. This was the first publication in which the drying process was used to nonspecifically coat a chemical onto a gold microprojectile [30]. This technology was first developed for the animal cell research and was termed as “Diolistics” [31]. One of the first studies of delivery of proteins through biolistics was reported by Wu and colleagues [2, 3]. To coat the protein/ DNA complexes to gold particles, they functionalized the projectiles to bind helper proteins that locked a transposome and facilitated the release once reached the intracellular space. In the work published by our group, we used gold-functionalized mesoporous silica nanoparticles loaded with proteins to bombard plant tissues [5, 6, 32]. These nanoparticles can be modified to accommodate in their pores different proteins of different nature. The protein is protected from degradation when loaded in the pore and the pores can be capped to enable controlled release of the cargo [27]. However, there are limitations to this approach. These include the size of the protein, the physical interactions between the protein and the nanoparticle and the need to use DNA and/or gold particles to improve the nanoparticle delivery efficiency [26]. To simplify protein delivery through biolistics we then decided to directly deposit the protein on gold microprojectiles by drying the suspension directly onto the macrocarriers used for bombardment. Initially, lyophilization was used to freeze-dry the suspension to avoid protein degradation. Later, we showed that drying under room temperature (22  C) was also adequate for several proteins tested to achieve the nonspecific coating on the gold microprojectiles and retain the protein activities (Fig. 1a). Because the method relies on a simple drying step to coat the microprojectiles with protein, there is no limitation on the physical or chemical properties of the protein. With this method, it is not only possible to deliver proteins, but also DNA, chemicals, and codeliveries of different biomolecules [1]. This drying-based protocol has been already used with some modifications by other groups to deliver nanoparticles and protein cargoes to plants [16] and to produce genome edited plants by delivering CRISPR/Cas9 ribonucleoproteins into different crops [8–10]. It shows the reproducibility and the potential applicability of this technique. Any scientist trained in plant transformation will be able to follow the presented protocol that will be focused on the delivery of two fluorescent proteins to two different plant tissues, as an example.

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 Freeze-dry

a

Air-dry

Bombardment

b

Red channel

TRITC-BSA – Freeze dry

Bright field

Intracellular Protein Delivery

250 µm

DsRed – Air dry

c

d

e

TRITC-BSA – Freeze dry TRITC-BSA – Freeze dry

300 µm

500 µm

50 µm

Fig. 1 (a) Proteolistics delivery scheme. The mix of gold particles and protein is distributed in the center of a macrocarrier of the gene-gun PDS-1000/He (Bio-Rad). Then, two methods can be applied to dry the suspension: Freeze-drying using liquid nitrogen and a freeze-drier or air-drying by leaving the macrocarrier

Proteolistics: A Protein Delivery Method

2 2.1

299

Materials Plant Materials

1. Onion epidermis tissue: obtained from white onion purchased from local grocery stores. 2. Maize embryos: immature embryos (1.5–2 mm long) of genotype Hi-II (greenhouse grown) are harvested 9–12 days after pollination.

2.2 Stock Solutions and Media

1. TS [Tris(hydroxymethyl)aminomethane-sodium chloride] buffer: 15 mM Tris–HCl (pH 8.0), 250 mM NaCl. 2. Maize ear disinfection solution: 2.6% sodium hypochlorite with a drop of Tween 20. 3. N6 vitamin stock (1000): 2 g/L glycine, 1 g/L thiamine, 0.5 g/L Pyridoxine, and 0.5 g/L nicotinic acid are dissolved in ddH2O. Filter-sterilize the solution using 0.2 μM filters and store at 4  C. 4. 2,4-D stock (1 mg/mL): Dissolve 250 mg 2,4-dichlorophenoxy acetic acid (2,4-D) in 10 mL 1 N KOH (see Note 1). When dissolved, bring the final volume to 250 mL with ddH2O. Store at 4  C. 5. Silver nitrate stock (50 mM): Dissolve 850 mg of silver nitrate in 100 mL ddH2O. Filter-sterilize the solution using 0.2 μM filters and store at 4  C (see Note 2). 6. Agar medium for onion epidermis tissue bombardment [0.5 mM MES (2-(N-morpholino)ethanesulfonic acid]: In 900 mL deionized water (ddH2O), add 98 mg MES, adjust pH to 5.7 using 1 N KOH, bring final volume to 1 L, add 15 g BD Bacto agar and autoclave at 121  C for 20 m. After cooling down to about 55  C, the medium is dispensed into 15  100 mm petri dishes (see Note 3). 7. Osmotic medium for maize immature embryo bombardment (as previously described with some modifications [33]): In 900 mL ddH2O, add 4 g N6 micro and macro elements (Duchefa), 2 mL 2,4-D stock solution, 2.8 g L-proline, 100 mg casein hydrolysate, 20 g sucrose, 36.4 g sorbitol, and 36.4 g mannitol. Adjust pH to 5.8 using 1 N KOH and bring

ä Fig. 1 (continued) assembly for at least 2 h in a flow bench. Once the suspension is dried, the plant tissue of interest is bombarded. After bombardment, intracellular protein delivery is achieved. (b) The freeze-dried mixture of gold particles and TRITC-BSA protein on a macrocarrier right before bombardment. (c) DsRed protein delivery in onion epidermis cells after bombardment with an air-dried suspension of the protein and gold particles. (d) Scutellum of an immature maize embryo right after bombardment with a freeze-dried mixture of TRITC-BSA protein and gold particles. (e) Close-up of the maize scutellum of (d). Arrows point to areas where more gold particles were localized, which correspond with more intracellular protein delivery

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the final volume to 1 L with ddH2O. Add 2.5 g Gelrite and autoclave. After cooling down to about 55  C, 1 mL of the 1000 N6 vitamin stock solution and 1 mL silver nitrate stock solution is added to the medium for a final concentration of 1 and 50 μM, respectively. The medium is dispensed into to 15  100 mm petri dishes (see Note 3). 2.3 Proteins (See Note 4)

1. Discosoma sp. red fluorescent protein (DsRed, 27.6 kDa) was purchased from BioVision as a lyophilized powder and was reconstituted using autoclaved ddH2O to 100 ng/μL. Aliquots of the solution were stored at 20  C. 2. Bovine serum albumin (66.4 kDa) labeled with tetramethylrhodamine-isothiocyanate (TRITC-BSA) was purchased from Invitrogen as a lyophilized powder. It was reconstituted using TS buffer to 25 μg/μL. Aliquots of the solution were stored at 20  C.

2.4 Equipment and Related Supplies

1. Gene-gun: PDS-1000/He (Bio-Rad). All the gene-gun disposable materials for bombardment such gold microprojectiles (0.6 μm), rupture disks (650 and 1100 psi), macrocarriers, macrocarrier holders, and stopping screens are purchased from Bio-Rad (see Note 5). 2. Freeze-drier: Labconco Freezone 2.5. 3. Microscope and image capturing software: Axiostar plus from Zeiss and ProgRes Capture Pro 2.6. The microscope had a red channel filter (excitation 545 nm, emission 620 nm and beam splitter 455 nm) from Chroma Technology. 4. Horizontal flow bench. 5. Ultrasonic bath. 6. Bench microcentrifuge. 7. Vortex Genie 2.

2.5

Other Supplies

1. 100% absolute ethanol, stored in

20  C.

2. Drierite desiccant. 3. Liquid nitrogen. 4. Autoclaved ddH2O.

3

Methods

3.1 Gold Particle Preparation

The following protocol described below is based on [33] with modifications: 1. Weight 15 mg 0.6 μm gold particles (see Note 6) in a 1.5 mL microcentrifuge tube and add 500 μL freezing cold 100% ethanol. Ultrasonicate the tube in a water bath for 15 s.

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2. Centrifuge 60 s at 805  g and remove the ethanol supernatant using a pipette. 3. Add 1 mL sterile ddH2O and finger vortex the tube. 4. Repeat step 2. 5. Resuspend the washed gold particles with 500 μL sterile ddH2O (see Note 4), finger vortex and ultrasonicate the tube in a water bath for 15 s. 6. Place the tube in a vortex at medium speed (setting 3) to keep the gold particles in suspension. 7. Divide the suspension into 50 μL aliquots (30 μg/μL) and store the tubes at 20  C until use. 3.2 Protein–Gold Particle Suspension Preparation

The amounts described are per shot. 1. Assemble a macrocarrier-set according to the Bio-Rad PDS-1000/He biolistic device manual. 2. Defrost a gold aliquot (30 μg/μL, 50 μL/tube). Mix the content well by vortexing the tube on Vortex Genie 2 (setting 3) for 20 s. Take 2 μL of the gold suspension and place it into a fresh minicentrifuge tube. 3. Add 12 μL of the DsRed protein solution (100 ng/μL) or 10 μL of the TRITC-BSA protein solution (25 μg/μL; see Notes 7 and 8). Mix well by pipetting. 4. In the center of the macrocarrier, distribute the suspension as a thin liquid layer: 14 or 12 μL of final volume for DsRed or TRITC-BSA protein, respectively (Fig. 1a; see Note 9).

3.3 Drying the Suspension on the Macrocarrier

We describe here two methods for drying the protein/gold suspension (see Note 10).

3.3.1 Air-Drying

1. Leave the macrocarrier-set containing the liquid DsRed protein/gold particle suspension to dry on bench top of a laminar flow bench at room temperature (22  C) for at least 2 h (see Note 11).

3.3.2 Freeze-Drying

1. With the help of forceps, immerse the macrocarrier-set loaded with the TRITC-BSA protein/gold particle suspension in liquid nitrogen for 5–10 m. 2. Place the frozen loaded macrocarrier-set in the freeze-drier immediately. 3. Freeze-dry for 1 h (see Note 12).

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3.3.3 Storage of Macrocarrier-Set Until Bombardment and Post-Drying Protein Check

3.4 Plant Target Tissue Preparation 3.4.1 Preparation of Onion Epidermis Tissue

3.4.2 Preparation of Maize Immature Embryos

1. Keep the dried macrocarrier-set in a sterile petri dish containing Drierite desiccant at room temperature (22  C) until bombardment (see Note 13). 2. Check under the microscope the correct drying and fluorescence of the protein/gold mixture. This step is optional (Fig. 1b; see Note 14). 1. Using scalpel blade and forceps remove a rectangular piece (approximately 2.5  3 cm) of epidermis from the scale leaves and place it in the center of a petri dish containing agar medium (see Note 15). The peeled side is placed upward. 2. The plates are closed and kept at room temperature (22  C) until bombardment (see Note 16). 1. Immature ears of Hi-II maize (9–12 days after pollination) are dehusked and disinfected using maize ear disinfection solution for 25 m (see Note 17). 2. In a laminar flow bench, rinse the ears thoroughly using sterile ddH2O three times. 3. To dissect immature embryos, remove the kernel crowns by cutting off the top 1–2 mm of the kernel with a sharp scalpel blade. 4. Excise the embryos by inserting a narrow spatula between the endosperm and pericarp to remove the endosperm out of the seed coat. Collect the embryos (~1.5 mm in size) and place them on a petri dish containing osmotic medium. About 20–30 embryos are arranged on the center of plate with the scutellum side (or round surface side) up. 5. Embryo osmotic treatment duration before bombardment ranges between 2 and 4 h [33].

3.5 Bombardment Using PDS-1000/He Biolistic Device (See Note 18)

1. Operation of the PDS-1000/He gene-gun is done according to the manufacturer instructions. For experiments described here, we use 6 mm gap distance between the helium nozzle and the launch assembly, and 6 cm distance from the stopping screen to the target tissue. The tissues are bombarded once. 2. For onion epidermis tissue bombardment, we use 1100 psi rupture disks. 3. For maize immature embryos bombardment, we use 650 psi rupture disks.

3.6 Monitoring Intracellular Delivery

1. Successful delivery of fluorescent proteins into the onion epidermis tissues and maize embryos (see Note 19) can be visualized immediately or within 1–2 h after bombardment (see Note 20).

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2. Intracellular fluorescent protein delivery can be observed in onion epidermis (Fig. 1c) or maize embryo scutellum cells (Fig. 1d, e; see Note 21).

4

Notes 1. 2,4-D should be weighed in a fume hood to prevent accidental inhalation. 2. The bottle containing the silver nitrate stock solution should be wrapped in foil paper to avoid light exposure. It can be stored at 4  C for up to 1 year. 3. Media are poured at a volume of about 40 plates per liter. All plates are dried thoroughly in the laminar flow bench. They can be stored at room temperature (22  C) in the dark for up to 1 month. 4. It is common to use glycerol to store proteins in liquid form or even the stock of gold particles. Glycerol interferes with the drying process; therefore, its use should be avoided. Additionally, if some buffer components that were used to purify or solubilize the protein of interest are known to be toxic to cells, they should be removed through dialysis beforehand. Otherwise, these toxic compounds will be delivered along with the protein into cells and cause potential inadvertent damage to the cell. 5. Plastic materials such as rupture disks and macrocarriers are sterilized by soaking the disks in 70% ethanol for 15 m and leaving them to dry in a flow bench. They can be stored in sterile petri dishes until use. Metal materials such as macrocarrier holders and stopping screens are autoclaved. In this chapter, a macrocarrier set refers to a plastic macrocarrier placed in the metallic macrocarrier holder. 6. Different sizes and sources of gold particle and other materials such as tungsten particle can be used as microprojectiles. Moreover, the dried protein alone, with no microprojectiles can also be delivered, even though the efficiency was much lower [1]. 7. The protein-to-gold particle ratio is one of the key parameters for the success of this technique. We suggest starting with a constant gold particle amount (60 μg in 2 μL sterile ddH2O) and varying the protein amounts. For the proteins tested so far, the ranges vary from 1 to 275 μg of protein per shot, depending on the availability and function of the protein. Using a high amount of protein is suggested for fluorescent proteins. Since the detection is done visually, the more protein used, the easier for visual detection. However, when high amounts of protein are used, cell toxicity needs to be considered. We also have observed that using high amounts of protein can result in a

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sticky layer over the macrocarrier after drying. This sticky mixture does not result in higher frequencies of protein delivery. 8. In this step of the protocol is where a mix of proteins and/or chemicals and/or other biomolecules can be done along with the gold particles for codelivery [1]. For example, to codeliver protein and DNA, per shot, 150 μg of TRITC-BSA protein was mixed with 1 μg of GFP (Green Fluorescent Protein) expressing plasmid DNA pLMNC95 [34] and 60 μg of gold particles. 9. An important aspect is the final volume of the protein/gold mixture. Ideally, this should not exceed 20 μL in total for a good distribution on the macrocarrier and fast drying. Depending on the protein solution we have observed that the protein/gold suspension could have a high surface tension when placed in the plastic macrocarrier. To favor the distribution and drying process, a “dot” pattern is suggested, where the suspension is placed scattered through the center of the macrocarrier in 1–3 μL droplets [1]. 10. Freeze-drying the macrocarrier-set loaded with the protein/ gold mixture can help to keep the protein integrity and it is the method of choice in the case the protein degrades easily. We have delivered the same protein that was either air-drying or freeze-drying to the same type of tissue and the results were comparable. 11. The drying time will depend on the volume of the protein/ gold suspension and the temperature and humidity of the environment. We tested drying times from 1 to 16 h. The proteins were dried and were delivered successfully. To speed up drying the macrocarrier-set containing the protein/gold suspension can be placed in an opened sterile petri dish containing a desiccant like Drierite. 12. The time necessary to dry the protein/gold suspension will depend on the type of freeze-dryer available. For the Labconco Freezone 2.5, the freeze-drying process was done at 45  C for 1 h. We recommend adjusting the freeze-drying time to the freeze-dryer equipment available. 13. We recommend using the dried macrocarriers for bombardment immediately after drying or within 1–2 h after drying is completed. 14. To ensure that the protein is not affected by the coating process it is recommended to do a protein integrity test where the protein of interest undergoes the mixing with the microprojectiles and the drying process on the macrocarrier-set. This protein is then rehydrated using water or saline solution and the integrity of the protein is tested by measuring its activity, detecting its fluorescence or detecting it by western blot, for example.

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15. We have observed that the onion epidermis tissues coming from different scale leaves show significant differences in cell size and bombardment response. The outer scales have bigger cells and are harder to bombard. On the contrary, the inner scale leaves have smaller cells and the tissue is softer. When comparing protein delivery conditions in different onion epidermis tissues, we recommend using epidermis tissues coming from the same scale leaves. 16. We normally prepare first the macrocarrier-set containing the protein/gold mixture and leave it to dry, either by air-drying or freeze-drying. Since drying takes 1–2 h, this time is employed to prepare the onion epidermis tissues. The tissues can be cultured in this media for several hours, but normally they are bombarded after the drying of the protein/gold mixture is finished. 17. To facilitate the disinfection and embryo dissection, one can cut off the top 1 cm of the ear and insert a handle, this can be a pair of forceps, into the top end of the ear. The ears with the handle will be placed in a large container (e.g., 2 L beaker) for disinfection. 18. For each type of tissue to be bombarded a preliminary experiment should be done where bombardment conditions (mainly number of shots, distance to the target, and pressure) are optimized using standard DNA delivery. However, the dried mixture of gold particles and protein normally is more compact than a standard DNA coating [1]. We have observed an increase in cell damage after protein delivery. If this happens, reducing the pressure, increasing the target distance, or decreasing the amount of protein/gold mixture should be considered. 19. In this protocol, the onion epidermis tissues are nonsterile. Contaminations of the tissue can be expected as early as 1–2 days after bombardment. Therefore, the experiments conducted in this tissue should be transient. In the case of maize immature embryos, the tissues and the bombardment conditions are sterile. Normally our experiments finished after protein delivery observation, but these embryos could be subcultured to a nonosmotic media for continuation of callus formation and regeneration as previously described [33]. 20. Monitoring the delivery of fluorescent proteins should be done right after bombardment and for a period of 1–2 h. If the tissues are observed right after bombardment the dispersion of the protein throughout the cytoplasm can be visualized in real time. Depending on the protein, the fluorescence can be observed for 30 m (GFP) or up to 1–2 days after bombardment (TRITC-BSA). When delivering enzymes,

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the activity should be measured immediately after bombardment (β-glucuronidase, for example). For other enzymes (RNase A or trypsin) we measured the activity 1 day after bombardment [1]. 21. We have observed that the success of protein delivery can be more variable than that of DNA delivery. In protein delivery optimization experiments, where different parameters have been tested using fluorescent markers, we found that the comparison could only be done qualitatively. This is because the detection and counting of the fluorescent cells was achieved visually under a microscope.

Acknowledgments This work was partially supported by the US Department of Agriculture National Institute of Food and Agriculture (Hatch project no. IOW05162), and State of Iowa funds. References 1. Martin-Ortigosa S, Wang K (2014) Proteolistics: a biolistic method for intracellular delivery of proteins. Transgenic Res 23:743–756 2. Wu J, Du H, Liao X, Zhao Y, Li L, Yang L (2011) Tn5 transposase-assisted transformation of indica rice. Plant J 68:186–200 3. Wu J, Du H, Liao X, Zhao Y, Li L, Yang L (2011) An improved particle bombardment for the generation of transgenic plants by direct immobilization of relleasable Tn5 transposases onto gold particles. Plant Mol Biol 77:117–127 4. Numata K, Horii Y, Motoda Y, Hirai N, Nishitani C, Watanabe S, Kigawa T, Kodama Y (2016) Direct introduction of neomycin phosphotransferase II protein into apple leaves to confer kanamycin resistance. Plant Biotechnol 33:403–407 5. Martin-Ortigosa S, Trewyn BG, Wang K (2017) Nanoparticle-mediated recombinase delivery into maize. In: Eroshenko N (ed) Site-specific recombinases. Methods in molecular biology, vol 1642. Humana Press, New York 6. Martin-Ortigosa S, Peterson DJ, Valenstein JS, Lin VS-Y, Trewyn BG, Lyznik LA, Wang K (2014) Mesoporous silica nanoparticlemediated intracellular cre protein delivery for maize genome editing via loxP site excision. Plant Physiol 164:537–547

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Proteolistics: A Protein Delivery Method 13. Yanagawa Y, Kawano H, Kobayashi T, Miyahara H, Okino A, Mitsuhara I (2017) Direct protein introduction into plant cells using a multi-gas plasma jet. PLoS One 12: e0171942 14. Staiger CJ, Yuan M, Valenta R, Shaw PJ, Warn RM, Lloyd CW (1994) Microinjected profilin affects cytoplasmic streaming in plant cells by rapidly depolymerizing actin microfilaments. Curr Biol 4:215–219 ´ balos JM, Doonan 15. Wymer CL, Ferna´ndez-A JH (2001) Microinjection reveals cell-to-cell movement of green fluorescent protein in cells of maize coleoptiles. Planta 212:692–695 16. Makhotenko AV, Snigir EA, Kalinina NO, Makarov VV, Taliansky ME (2018) Data on a delivery of biomolecules into Nicothiana benthamiana leaves using different nanoparticles. Data Brief 16:1034–1037 17. Ng KK, Motoda Y, Watanabe S, Othman AS, Kigawa T, Kodama Y, Numata K (2016) Intracellular delivery of proteins via fusion peptides in intact plants. PLoS One 11:e0154081 18. Chuah J-A, Horii Y, Numata K (2016) Peptide-derived method to transport genes and proteins across cellular and organellar barriers in plants. J Vis Exp 118:e54972 19. Ziemienowicz A, Shim YS, Matsuoka A, Eudes F, Kovalchuk I (2012) A novel method of transgene delivery into triticale plants using the Agrobacterium transferred DNA-derived nano-complex. Plant Physiol 158:1503–1513 20. Shim Y-S, Eudes F, Kovalchuk I (2012) dsDNA and protein co-delivery in triticale microspores. In Vitro Cell Dev Biol Plant 49:156–165 21. Chang M, Chou J-C, Chen C-P, Liu BR, Lee H-J (2007) Noncovalent protein transduction in plant cells by macropinocytosis. New Phytol 174:46–56 22. Chugh A, Eudes F (2008) Study of uptake of cell penetrating peptides and their cargoes in permeabilized wheat immature embryos. FEBS J 275:2403–2414 23. Lu S-W, Hu J-W, Liu BR, Lee C-Y, Li J-F, Chou J-C, Lee H-J (2010) Arginine-rich intracellular delivery peptides synchronously deliver covalently and noncovalently linked proteins into plant cells. J Agric Food Chem 58:2288–2294 24. Chugh A, Eudes F, Shim Y-S (2010) Cellpenetrating peptides: Nanocarrier for

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Chapter 17 Biolistic Delivery of Programmable Nuclease (CRISPR/Cas9) in Bread Wheat Abhishek Bhandawat, Vinita Sharma, Vikas Rishi, and Joy K. Roy Abstract The discovery of site-specific programmable nucleases has led to a major breakthrough in the area of genome editing. In the past few years, CRISPR/Cas system has been utilized for genome editing of a large number of crops including cereals like wheat, rice, maize, and barley. In terms of consumption, wheat is second only to rice as the most important crop of the world. In the present chapter, we describe biolistic delivery method of ribonucleoprotein (RNP) complexes of programmable nuclease (CRISPR/Cas9) for targeted genome editing and selection-free screening of transformants in wheat. The method not only overcomes the problem of random integration into the genome but also reduces the off-targets. Besides the step-by-step protocol, plausible challenges and ways to overcome them are also discussed. By using the described method of biolistic delivery of CRISPR/Cas9 in plant systems, genome-edited plants can be identified within 11 weeks. Key words Biolistic delivery, CRISPR/Cas9, Guide RNA, Programmable nuclease, Wheat

1

Introduction Agrobacterium and biolistic-mediated transformations are two popular reverse genetics approaches to study the function of specific gene and to introduce important agronomic traits in crops. The former approach exploits a supervirulent Agrobacterium strain to infect and deliver gene of interest along with the T-DNA into the host genome. Biolistic approach delivers exogenous nucleic acid or protein using particle bombardment. However, when it comes to transformation of recalcitrant monocots, cotransformation of multiple genes, vector-free gene integration, biolistic approach is preferred over Agrobacterium-mediated transformation [1, 2]. Another challenge associated with genetic transformation is that, it often leads to unpredictable genetic modifications, causing silencing of the target gene, interruption of native genes, plant mosaicism, and vector backbone incorporation [3, 4], thus rendering these approaches unsuitable for efficient functional validation/

Sachin Rustgi and Hong Luo (eds.), Biolistic DNA Delivery in Plants: Methods and Protocols, Methods in Molecular Biology, vol. 2124, https://doi.org/10.1007/978-1-0716-0356-7_17, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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crop improvement. Programmable nucleases overcome the above limitations by targeting specific sequence patterns for precise genome editing. Among various programmable nuclease such as transcription activator-like effector nucleases (TALENs), zinc finger nucleases (ZFNs), RNA-guided Cas9 nuclease derived from prokaryotic clustered regularly interspaced short palindromic repeats (CRISPR) has emerged as a preferred technique for crop improvement due to its simplicity, versatility and precise gene editing [5–10]. Cas9 offers broad range of guide RNA designing target sites due to simple requirement of 20 bp target sequence preceding PAM, offering noticeable advantages over ZFN and TALEN. Multiple gene editing can be performed by using multiple sgRNA in a single transformation experiment [11, 12]. CRISPR/Cas system originally belongs to the prokaryotic defense system to protect self from invading alien DNA/RNA. The sgRNA forms functional complex with Cas9 protein (Cas9 RNPs), guides the nuclease to genomic loci having 20 bp complementarity with the sgRNA, and makes double-stranded DNA break (DSB) immediately upstream 50 -NGG protospacer adjacent motif (PAM). The DNA break is repaired either by an error-prone nonhomologous end joining (NHEJ) pathway or homology directed repair (HDR) that results in nucleotide insertion/deletion (indels) or substitution [13]. Most of the mutants developed by Agrobacterium-mediated or biolistic transformation involve CRISPR/Cas9 DNA [14]. A limitation of using DNA-encoded CRISPR/Cas system for genome editing in crops is due to constitutive expression of guide-RNA and Cas9 proteins that leads to random and nonspecific genome targeting and off-target effects. These problems, to a larger extent, limit the successful adoption of this method for crop improvement. The use of CRISPR/Cas9 in vitro transcripts (IVTs) and ribonucleoproteins (RNPs) has been proved successful in preventing random integration of exogeneous DNA, and thus reducing the frequency of off-target cleavage in many species [15– 18]. This makes the implementation of Cas9 RNP an attractive proposition for successful genome editing in crops like wheat and maize [19, 20]. Wheat (Triticum spp.) is the second most consumed staple crop in the world. Meeting the rising demands of expanding human population, a rapid method to develop new varieties having improved yield, quality and resistance to biotic and abiotic stresses is the need of the hour. Incorporation of targeted genomic alterations in bread wheat (Triticum aestivum L.) has faced several challenges due to large and complex genome (~15.8 Gb), allohexaploidy (2n ¼ 6x ¼ 42, AABBDD), difficult transformation, and poor regeneration [21]. The efficiency and preciseness of Cas9 RNPs over other genome editing approaches demonstrated by homeolog-specific gene editing which reflects its ability to distinguish single nucleotide difference in hexaploid wheat [22], along

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Fig. 1 Overview of biolistic delivery of CRISPR/Cas9 ribonucleoproteins for genome editing in bread wheat. The section at the left illustrates sgRNA designing, cloning, and validation of Cas9 activity. Cas9 RNP coating on gold nanoparticles, biolistic delivery to immature embryos, regeneration and mutant screening are illustrated in the right section. A 20 bp target sequence in the immediate upstream of PAM (Protospacer Adjacent Motif) is

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with the availability of reference bread wheat genome have opened up new avenues for wheat genetic improvement. This chapter describes an efficient biolistic transformation and regeneration protocol using preassembled Cas9-sgRNA (single-guide RNA) RNP complex-mediated targeted genome editing in bread wheat embryo, addressing the existing challenges. For the sake of simplicity, the method section is divided into two parts (Fig. 1): the first covers the design of sgRNA and validation of CRISPR/Cas9 RNP activity, the second part explains biolistic delivery of Cas9 RNPs in wheat embryos, and screening of mutant plant by antibiotic selection-free mixed-pool genotyping method [19].

2

Materials

2.1 Materials for Designing and Validating CRISPR/Cas9 RNPs

1. Scaffold vector: For cloning and in vitro transcription of sgRNA, pTaU6 plasmid containing TaU6 promoter can be used [23, 24].

2.1.1 sgRNA Cloning

3. T4 DNA ligase for ligation of sgRNA DNA to scaffold vector.

2. Restriction enzymes to cut scaffold vector. 4. Competent cells: To clone sgRNA vector, E. coli DH5-α or Top-10. 5. LB medium: Dissolve 20 g of Luria Broth (10 g tryptone, 5 g yeast extract, 5 g NaCl) powder in 1 L distilled water. Heat to homogenize the media and autoclave. If unopened, the media can be stored at 4  C for up to 6 weeks. 6. Luria Agar plates for selection of transformed cells: Dissolve 35 g of Luria Agar (10 g tryptone, 5 g yeast extract, 5 g NaCl, 15 g agar) in 1 L of distilled water. Heat to form a homogeneous media and autoclave. After cooling to about 60  C, add

ä Fig. 1 (continued) selected as potential sgRNA for targeting Cas9 nuclease. The synthesized single-guide RNA oligos has flanking restriction site complementary to its scaffold vector for sticky-end cloning. The mixture of Cas9 protein and in vitro transcribed sgRNA is validated using in vitro PCR-based assay. Two bands show cleavage of Cas9-sgRNA treated target PCR product (T), while “W” is the untreated PCR product. In vivo activity of Cas9-sgRNA is assessed in transfected wheat protoplast cells. If the target sequence contains unique restriction site, PCR of target gene followed by restriction digestion (PCR/RE assay) shows two bands in untreated wild (W), while single band in treated sample (T). T7EI cleaves at the mismatch in Cas9-sgRNA treated PCR product while not the untreated wild. The validated sgRNA is selected to form Cas9 RNP complex followed by coating on gold nanoparticles. The Cas9 RNP coated gold particles are bombarded to immature wheat embryos using biolistic gene gun. Cas9 RNP delivery is verified in pooled embryo DNA followed by target gene PCR and sequencing. Transformed embryos are regenerated on selection-free media. Genome edited mutant plants are screened by PCR/RE assay: C represents wild undigested control, W is wild digested, Taa is homozygous mutant and TAa is heterozygous mutant

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suitable concentration of antibiotic based on selection marker in the vector. Swirl well to mix and pour about 30 mL to petri dishes. The plates may be stored for up to 6 weeks when stored at 4  C. 7. Gel extraction Kit (Qiagen) for DNA purification. 8. Plasmid miniprep kit (Qiagen) for high quality plasmid DNA extraction. 9. Thermocycler for PCR amplification. 10. 9-cm sterile petri dishes. 2.1.2 In Vitro Transcription of sgRNA

1. High-fidelity DNA polymerase such as Pfu/Vent, with proofreading (30 !50 exonuclease) activity. 2. High-yield RNA synthesis kit (HiScribe T7) for transcription of RNA. 3. DNase I to remove DNA contaminants.

2.1.3 Cas9 RNP Synthesis and Validation

1. Cas9 protein: Although Cas9 protein (see Note 1) is commercially available, it contains higher glycerol making it highly viscous, reducing the efficiency of delivery to immature embryo cells. Experienced users may go for expression of recombinant Cas9 protein in E. coli cells for improved delivery of Cas9 protein as described earlier [19]. 2. Solution 1 (500 mL) pH 5.7: NaCl (1 M), CaCl2·2H2O (1 M), KCl (2 M), 2-(N-Morpholino) ethanesulfonic acid (MES) (0.2 M) in nuclease-free water. Autoclaved Solution 1 can be stored at 25  C for up to a maximum of 2 months. 3. Transfection solution: Freshly prepare PEG solution by adding 4 g PEG 4000, 1 mL 0.8 M mannitol and 400 μL of 1 M CaCl2 in nuclease-free water. 4. Restriction enzymes for PCR/RE assay. 5. T7 endonuclease I (T7EI) for identification of mismatch in DNA duplex.

2.2 Material for Biolistic Delivery of Cas9 RNPs and Mutant Screening 2.2.1 Plant Material

Imbibe the wheat (Triticum aestivum) seeds and stratify at 4  C for 2–4 days (see Note 2). Transfer the seeds in batches to growth chambers under controlled conditions, maintaining 16 h light:8 h dark photoperiod using artificial lights (at PAR intensity ~700 μmol/m2 s) (see Note 3). To facilitate efficient germination, day/night temperature should be maintained 18  C/12  C for 1–2 weeks. Vernalize the seedlings for 3 weeks as mentioned later in Sect. 3.2.1. As the plant heads toward jointing and earing stages, the day/night temperature should be maintained at 28  C/18  C.

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2.2.2 Sterilization Material

1. 70% (v/v) ethanol for surface sterilization. 2. 2.5% (w/v) sodium hypochlorite (see Note 4) for disinfection. 3. Sterile water: Autoclaved distilled water or ultrapure reverse osmosis (18.2 MΩ/cm) water for washing. 4. Scalpel. 5. Whatman filter paper grade 1.

2.2.3 Particle Bombardment

1. Gold nanoparticles (0.6 μm diameter) are suitable for biolistic delivery to small wheat cells (see Note 5). 2. Absolute alcohol. 3. Sonicator.

2.2.4 Regeneration of Embryos

1. High osmotic medium (1 L): Suspend 4.33 g of Murashige and Skoog (MS) basal salt media [ammonium nitrate (1.65 g/L), boric acid (6.2 mg/L), CaCl2 anhydrous (332 mg/L), CoCl2 .6H2O (0.025 mg/L), cupric sulfate pentahydrate (0.025 mg/L), Na2EDTA.2H2O (37.26 mg/L), ferrous sulfate heptahydrate (27.8 mg/L), MgSO4 anhydrous (180.7 mg/L), MnSO4.H2O (16.9 mg/L), molybdic acid (sodium salt) dihydrate (0.25 mg/L), KI (0.83 mg/L), potassium nitrate (1.9 g/L), potassium phosphate monobasic (170 mg/L), zinc sulfate heptahydrate (8.6 mg/L)], 2,4-dichlorophenoxyacetic acid (2,4-D; 5 mg/L), 72.9 g mannitol in 900 mL of sterile water. Adjust the pH to 5.75  0.5 using KOH, add 3.2 g Phytagel and autoclave. Pour ~25 mL media in petri dishes and store at 4  C for a maximum of 2 months. 2. Recovery medium (1 L): Suspend 4.4 g of MS basal salt with vitamins [glycine (free base) (2 mg/L), myoinositol (100 mg/ L), nicotinic acid (free acid) (0.5 mg/L) pyridoxine hydrochloride (0.5 mg/L), thiamine hydrochloride (0.1 mg/L)] (see Note 6), 30 g sucrose, 2,4-Dichlorophenoxyacetic acid (2,4-D; 2 mg/L), 0.5 g N-Z-Amine A (amino acid and peptide source) and 600 μL CuSO4 (1 mg/mL) in 900 mL of sterile water. Adjust the pH to 5.75  0.5 with KOH. Make up the volume to 1 L with nuclease-free water and add 3.2 g of Phytagel. Autoclave and pour ~30 mL into sterile petri dishes. The plates can be stored at 4  C for up to a maximum of 6 weeks (see Note 7). 3. Regeneration medium (1 L): Suspend 4.4 g MS salt with vitamins, 30 g sucrose and kinetin (0.2 mg/L) in 900 mL sterile water. Adjust the pH to 5.75  0.5 with KOH and make up the volume to 1 L with nuclease-free water. Add 3.2 g of Phytagel, autoclave, pour the plates and store at 4  C for up to a maximum of 6 weeks in dark.

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4. Rooting medium (1 L): Suspend 2.2 g MS salt with vitamins, 30 g sucrose, kinetin (0.2 mg/L) in 900 mL of sterile water. Adjust the pH to 5.75  0.5 with KOH and make up the volume to 1 L of sterile water. Add 3.2 g of Phytagel and autoclave. After cooling to ~50  C, add 100 μL of 1-naphthaleneacetic acid (NAA; stock: 0.5 mg/mL). Pour ~30 mL of medium into sterile petri dishes and store at 4  C for up to a maximum of 6 weeks. 5. Jiffy pots. 2.2.5 Mutant Screening

1. Conserved primers to amplify all the three genome copies of the target gene. 2. Restriction enzymes.

3

Methods

3.1 Designing and Validation of CRISPR/Cas9 RNPs 3.1.1 Designing and Synthesis of Single-Guide RNA (sgRNA) for Cas9

For targeting the Cas9 nuclease to cleave specific genomic loci, it is prerequisite to design a highly specific sgRNA. The sgRNA consists of two parts: one forms a scaffold by several stem-loop structures for Cas9 binding and the other 20-nt sequence complementary to target sequence. The target site with unique 30 end reduces the chances of off-target cleavage [11]. Immediately downstream to the target guide RNA region, a 50 -NGG PAM (protospacer adjacent motif) site essential for most of the Cas9 binding should be present. However, some Cas9 homologs offer different PAM recognition site. A single unique restriction enzyme site within the target sgRNA sequences just upstream to the PAM recognition site would facilitate mutant detection by PCR/RE assay discussed in the later section (see Subheading 3.1.6). The existing tools for Cas9 sgRNA designing are not yet updated with recently published wheat genome sequence, therefore, sgRNA should be manually designed to target one or all the three homeologs, A, B, or D genome copies as follows: 1. First step is to scan for PAM site and select a target sequence (20 bp + 50 -NGG) within the gene of interest. Keep in mind that target sequence should not be in the intron region, if the aim is to disrupt gene function. 2. Next challenge for designing sgRNA is specificity and efficacy. Using blast search (http://blast.ncbi.nlm.nih.gov/) against the complete wheat genome, look for the unique match. Alternatively, CasOT tool (http://casot.cbi.pku.edu.cn/) can be used to identify potential off-targets. For genome-specific targeting, the designed sgRNA should have at least 3 mismatches with the other genome copies to avoid potential off-targets. Nuclease activity varies greatly with the sgRNA; hence, at least

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3 sgRNA should be designed for each gene. It is recommended to use GW2-sgRNA as positive control to check the nuclease activity for beginners. 3. Oligos synthesis: Once the sgRNA is designed, synthesize forward and reverse oligos to ligate to sgRNA scaffold vector. Oligos should contain the 20 bp with 50 and 30 restriction site (4–5 bp) overhang complementary to the digested scaffold vector. Different RNA polymerase III promoters have preference for a specific nucleotide to start transcription, so the nucleotide must be included immediately to 50 of the target sequence. 4. If the target is a model plant like Arabidopsis, rice, maize, web-based tools like “CRISPR-P” (http://www.genome. arizona.edu/crispr/) can be used for highly specific Cas9 target site detection and prediction of all possible off-targets. 3.1.2 Designing ssDNA oligos (Only for Homology Directed Repair)

If the aim of the gene editing is to substitute one or more nucleotide in the target gene sequence, then ssDNA donor template is required for homology directed repair. Design ssDNA oligos (20–90 nts long) having target site at the centre flanked by long homology arms on each side.

3.1.3 Cloning of sgRNA into Scaffold Vector

1. Dissolve the target specific forward and reverse oligos to a final concentration of 100 μM using nuclease-free water. Dilute the dissolved oligos to the final concentration of 10 μM. 2. For annealing ssDNA (target specific oligos), prepare the reaction as follows: Components

Final concentration

Annealing buffer

1

Forward oligos

4.5 μM

Reverse oligos

4.5 μM

Nuclease-free water

Up to 20 μL

3. Incubate the reaction in the thermocycler with the following conditions: 95  C for 5 min, 95–25  C (gradual decrease of 1  C per min), 70 min and hold at 10  C. The reaction can be stored at 20  C. 4. Digest the scaffold vector with the same restriction enzyme whose restriction site was incorporated in target specific sgRNA oligos as follows:

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Components

Final concentration

Enzyme activity buffer

1

Scaffold vector

2 μg

Restriction enzyme

4U

Nuclease-free water

Up to 50 μL

5. Incubate the reaction overnight (12–16 h) at 37  C. 6. Separate the digested vector by electrophoresis (1% agarose gel in TAE buffer). Here you will get a single DNA band at the corresponding molecular weight of vector. 7. Excise the DNA band from the gel and purify the DNA using gel extraction kit as per the manufacturer’s instructions. 8. Ligate the annealed target specific oligos to the linearized scaffold vector as follows: Components

Final concentration

T4 ligase buffer

1

Digested scaffold vector

50 ng

Annealed oligos (see step 3 in Subheading 3.1.3)

Thrice the molar concentration of the vector

T4 ligase

2.5 U

Nuclease-free water

Up to 10 μL

9. Incubate the mixture at 22  C for 1–2 h or at 16  C overnight. 10. Use 3–6 μL of ligated product to transform the E. coli (TOP-10 or DH5-α) competent cells as per the standard protocol. 11. Plate the transformed cells to the Luria Agar selection plate (with appropriate antibiotics) for the selection of recombinants and incubate overnight at 37  C in bacteriological incubator. 12. To confirm the transformation, pick 5–10 colonies and perform colony-PCR using one target and one vector specific forward and reverse primer. Check the PCR reaction on 1.2% Agarose gel, if the transformation is successful, DNA band at the corresponding molecular weight of amplicon size will be obtained. 13. Inoculate the individual PCR positive colonies into 5 mL LB medium and incubate overnight at 37  C with constant agitation.

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14. Extract the plasmid using Plasmid miniprep kit as per the manufacturer’s instructions. 15. To verify the recombinants harboring correct sgRNA, sequence the isolated plasmid DNA using vector specific forward and reverse primer. 3.1.4 In Vitro Transcription of sgRNA

1. Amplify the cloned sgRNA construct using T7-spacer forward and sgRNA specific reverse primers by PCR as mentioned below: Components

Final concentration

Pfu buffer

1

dNTP

1 mM (0.25 mM each)

T7-spacer primer F

0.5 μM

sgRNA primer R

0.5 μM

Pfu polymerase

1.25 U

sgRNA construct (see step 15 in Subheading 3.1.3) 0.1–0.5 ng Up to 50 μL

Nuclease-free water

2. Incubate the reaction into the thermocycler with the following conditions: Step 1: Denaturation

95  C, 5 min

Step 2: Denaturation Step 3: Annealing Step 4: Extension

95  C, 20 s 55-65  C (depends on Tm of primer used), 20 s 72  C, 20 s

Step 5: Final extension

72  C, 5 min

Steps 2–4 35 cycles

3. Run the PCR product on 1.2% agarose gel in TAE buffer. A single band of corresponding size of sgRNA sequence + scaffold RNA sequence is expected. 4. Excise the DNA band from the agarose gel and purify using gel extraction kit as per the manufacturer’s instructions. Quantify the eluted DNA using NanoDrop. 5. Using High-yield RNA synthesis kit (see Subheading 2.1.2), set up the reaction at room temperature as mentioned below:

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Components

Final concentration

Reaction Buffer

1

dNTP

1 M (250 mM each)

Template DNA (step 4 in Subheading 3.1.4)

0.5–1 μg

T7 RNA Polymerase mix

2 μL

Nuclease-free water

Up to 20 μL

6. Incubate the above PCR reaction in thermocycler at 37  C for 3 h. 7. To degrade the template DNA, give DNase I treatment for 1–2 h at 37  C. 8. Add 500 μL of freshly prepared 70% (v/v) ethanol and incubate the reaction at 20  C overnight for RNA precipitation. 9. Centrifuge at 12,000  g for 10 min at 4  C and discard the supernatant. Wash the RNA pellet twice with 500 μL of 70% (v/v) ethanol. Let it air-dry at room temperature. 10. Resuspend the RNA pellet in 50 μL of nuclease-free water and quantify the RNA concentration using NanoDrop and store at 80  C till further use. 3.1.5 Production of Cas9 Protein

Cas9 protein-mediated editing is more efficient as it rapidly degrades in the cell, thus reducing the chances of off-targets. Commercially produced Cas9 protein can be used, however, it has higher glycerol which makes the protein sticky on the carrier particles, reducing the efficiency of delivery to immature embryo cells. Therefore, Cas9 protein may be expressed in the lab for improved delivery of protein as described earlier [19].

3.1.6 Validation of Cas9 RNPs Activity

Before proceeding for biolistic delivery of Cas9 RNPs in planta for gene editing, first the activity of developed RNPs should be confirmed by in vitro (on PCR amplified target) and in vivo (on isolated protoplasts) assays.

In Vitro Assay

1. Amplify the targeted genomic DNA sequence (complementary to designed sgRNA) with the target specific primers using PCR. Separate the amplified product on 2% agarose gel in TAE buffer; cut and purify the single amplicon using gel extraction kit following the manufacturer’s instructions. 2. To assess the activity of CRISPR/Cas9 RNPs, prepare the reaction (see Note 8) as follows:

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Components

Final concentration

Cas9 Reaction Buffer

1

PCR purified DNA (see step 1)

100 ng

sgRNA

1 μg

Cas9 protein

1 μg

Nuclease-free water

Up to 20 μL

3. Mix well and incubate at 37  C for 1 h. 4. To terminate the reaction (dissociation of RNP complex) heat at 65  C for 10 min. 5. Run the reaction on 2% agarose gel in TAE buffer. The cleavage activity of Cas9 RNPs is confirmed if two bands of corresponding size are observed. In Vivo Assay (Using Protoplast)

1. Isolate the wheat protoplast following standard protocol [24]. 20–30 wheat seedlings can generate ~1  107 protoplasts. 2. PEG-mediated Transfection: Gently mix 20 μg of Cas9 protein (see step 1 in Subheading 2.1.3) and sgRNA (see step 10 in Subheading 3.1.4) each in a fresh 1.5 mL tube. Then, add protoplast (5  105 cells) and 240 μL of freshly prepared transfection solution (see step 3 in Subheading 2.1.3). Incubate the mixture in the dark for 20 min. 3. Add 900 μL of Solution 1 to stop the transfection. 4. Centrifuge at 100  g (to avoid rupture of protoplast) for 3 min at 25  C, discard the supernatant and resuspend the protoplast pellet into 1 mL of Solution 1 (see step 2 in Subheading 2.1.3). Incubate the tube in the dark at 23  C for 2 days. 5. After 2 days, harvest the protoplast by centrifugation at 12,000  g for 2 min, at room temperature and discard the supernatant. 6. Extract the DNA from transfected protoplast by the CTAB method [25]. Genomic DNA should be stored at 20  C till further use (see Note 9).

Assessment of Cas9 RNP Activity

CRISPR/Cas9 RNP activity can be assessed by either PCR/RE assay or T7EI assay. If the target sequence has a restriction enzyme site just upstream to PAM site, then perform PCR/RE assay, otherwise T7EI assay may be done.

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1. PCR/RE assay: (a) Set the PCR reaction as follows: Components

Final concentration

Fast Pfu Buffer

1

dNTP

1 mM (0.25 mM each)

Primer F

0.4 μM

Primer R

0.4 μM

DNA template (see step 6 of in vivo assay)

15–30 ng

Fast Pfu polymerase

2.5 U

Nuclease-free water

Up to 50 μL

(b) Incubate the above reaction in thermocycler with the following conditions: Step 1: Denaturation

95  C, 5 min

Step 2: Denaturation Step 3: Annealing Step 4: Extension

95  C, 20 s 55–65  C (depends on Tm of primer used), 20 s 72  C, 20 s

Step 5: Final extension

72  C, 5 min

Steps 2–4 35 cycles

(c) Set up the digestion reaction as follows along with a negative control (see Note 10): Components

Final concentration

Fast digest Buffer

1

PCR product (see step 1 (b) above)

0.5–1 μg

Restriction enzyme

1U

Nuclease-free water

Up to 50 μL

(d) Gently but thoroughly mix and incubate the reaction at 37  C for 1 h. Separate the digested product on 2% agarose gel in TAE buffer. This would yield an uncleaved band in the mutated DNA (Cas9 edited sample), while two bands in the case of wild type. This is because in Cas9treated sample restriction site will be edited (indel mutation), resulting in undigested intact band.

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(e) Using below relation, estimate the indel frequency [24]: a  100 Indel ð%Þ ¼ aþbþc where a is the intensity of undigested band (can be measured by ImageJ software), b and c are the intensities of digested products. Indel frequency >10% is recommended for stable plant transformation. 2. T7EI assay: (a) Prepare the reaction as follows: Components

Final concentration

T7EI Buffer

1

PCR product (see step 1 (b) above)

5 μL

Nuclease-free water

Up to 10 μL

(b) Incubate the above reaction into the thermocycler as: Denaturation at 95  C for 5 min followed by a touchdown PCR program of 95–15  C (with gradual decrease of 10  C per min). (c) Perform T7EI nuclease digestion along with digested wild-type sample as negative control as follows: Components

Final concentration

Heteroduplex from (see step 2 (b) above)

10 μL

T7EI

2.5 U

Nuclease-free water

Up to 10 μL

(d) Mix gently but thoroughly and incubate at 37  C. Check the reaction on 2% agarose gel in TAE buffer. Getting the digested bands in CRISPR/Cas9 treated sample (see Note 11) and not in the wild type, it is a good indicative of indel mutation. (e) Use the below relation [24] to calculate indel frequency: rffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi! bþc Indel ð%Þ ¼ 1  1   100 aþbþc where a is the intensity of undigested band, b and c are the intensities of digested products. (f) Identification of mutation by sequencing: Clone the PCR product to any blunt-end cloning vector and sequence the plasmid isolated from the recombinant plasmid. Follow standard protocols for cloning, transformation, and plasmid extraction.

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1. In vitro expression of sgRNA is mentioned above (see step 10 of Subheading 3.1.4). To form Cas9 RNP complex, purified Cas9 protein and guide RNA are mixed in 1 reaction buffer in a molar ratio of 5:1. 2. Mixture tubes should be prepared for ten deliveries as follows: Components

Final concentration

Cas9 reaction Buffer

1

sgRNA

20 μg

Cas9 protein

20 μg

Nuclease-free water

Up to 100 μL

3. Incubate at 25  C for 10 min. 3.2 Biolistic Delivery of CRISPR/Cas9 RNPs and Mutant Screening 3.2.1 Wheat Growth Conditions

3.2.2 Harvesting Spikes and Sterilization

Grow wheat seeds under controlled conditions in greenhouse while maintaining 16 h photoperiod. To facilitate efficient germination, day and night temperature should be maintained at 18  C and 12  C, respectively for 7–10 days. Vernalize the wheat seedlings for ~3 weeks at 4  C in cold house and then transfer to the greenhouse for 3 weeks. During jointing and earing stages, the day/night temperature should be maintained at 28  C/18  C. 1. For biolistic delivery of CRISPR/Cas9 RNPs, it is essential to harvest immature embryos at appropriate stage. Usually spikes ~14 days post anthesis are ideal (see Note 12). 2. Dip the spikes in 70% (v/v) ethanol for 30–60 s. Collect the caryopses of middle region of the spike, leaving behind the inner and outer caryopses, as they generally have smaller embryos due to asynchronous growth. 3. Surface sterilize the caryopses by washing with 70% (v/v) ethanol for ~45 s followed by soaking in 2.5% (w/v) sodium hypochlorite for 15–20 min with gentle shaking on platform shaker (~60 rpm). Wash the caryopses thoroughly but gently six times with sterile distilled or RO water under aseptic conditions and dry on filter paper.

3.2.3 Isolation of Immature Embryo

1. Collect immature embryos from caryopses using a sharp sterilized scalpel under optical microscope, carefully. Embryos that have slightly translucent scutellum and nearly opaque axis are more responsive for bombardment. Before biolistic bombardment, incubate the embryos in high-osmotic medium (see step 1 of Subheading 2.2.4) at least for 3–4 h (up to 2 days) at 23  C to improve recovery of embryos.

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3.2.4 Preparation of Gold Particle Suspension

1. Wash 40 mg gold particles (0.6 μm) with 1 mL absolute ethanol thoroughly by vortexing in a fresh 1.5 mL microcentrifuge tube. Sonicate in water bath for 90 s, short-spin (~5 s), and discard the supernatant. 2. Repeat the above washing step with absolute, 70% ethanol, and nuclease-free water, respectively. 3. Add 1 mL of nuclease-free water, vortex thoroughly to resuspend the gold particles. Make aliquots of 50 μL gold particle suspension in 1.5 mL microfuge tubes (see Note 13). The aliquots are best if used before 6 months when stored at 20  C.

3.2.5 Coating of Cas9 RNPs on Gold Nanoparticles

1. Take 50 μL of well suspended gold nanoparticles (see Subheading 3.2.4). Spread 15 μL of gently but thoroughly mixed Cas9 RNP mixture (see step 3 of Subheading 3.1.7) over the central region of macrocarrier, and let it air-dry on an undisturbed surface (see Note 14) for 1–2 h at 25  C.

3.2.6 Biolistic Delivery of Cas9 RNPs

At present, biolistic approach is the most preferred and successful approach of DNA delivery in wheat [26–28]. 1. Take ~100 embryos (see Subheading 3.2.3) per shot of bombardment using PDS-1000/He particle bombardment system (Bio-Rad) as per the manufacturer’s instructions (see Note 15), with target distance of 6 cm, pressurized helium at rupture pressure of 1100 psi under vacuum of 28 mm Hg (see Note 16). Detailed information about bombardment procedure in wheat tissue is described elsewhere [29]. Repeat the bombardment procedure with at least 3–4 plates. 2. For successful transformation keep the bombarded embryos in high-osmotic medium (see item 1 under Subheading 2.2.4) in dark at 23  C overnight.

3.2.7 Verification of Cas9 RNP Delivery

1. Two days after transformation, randomly select ~100 bombarded embryos for genomic DNA isolation by CTAB method [25]. 2. PCR-amplify the target gene copies using genome-specific primers followed by separation on 1.2% agarose gel. PCR reaction and conditions are the same as described under PCR/RE assay of Subheading 3.1.6. Using gel extraction kit, purify the PCR product and elute in 30 μL nuclease-free water. 3. Use the purified PCR products obtained in the previous step to perform a secondary PCR amplification as described in step 1a, b of PCR/RE assay with the same set of genome-specific primers but containing additional unique 6-nucleotide barcode sequence at 50 end.

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4. Separate all 50-μL products on 1.2% agarose gel in TAE buffer and excise the band of expected size (~110–130 bp) for subsequent purification using the gel extraction kit. 5. Make an equimolar pool of above PCR product (~200 ng each) and sequence it. 6. Measure the mutation frequency using online tools like Cas-Analyzer [30]. Detection of targeted mutagenesis confirms the successful delivery of Cas9 RNPs into explants. 3.2.8 Tissue Culture Regeneration of Bombarded Embryos

1. Incubate bombarded embryos derived from Subheading 3.2.6 in the recovery medium (see item 2 of Subheading 2.2.4) plates at 23  C in dark for 2 weeks for callus induction (see Note 17). After 2–3 weeks of incubation 4–6 mm calli will be formed. 2. Transfer the healthy calli (see Note 18) to the regeneration medium (see item 3 of Subheading 2.2.4) at 23  C for another 2–3 weeks with 16 h light and 8 h dark photoperiod. 3. By the end of 3 weeks green leaves should have developed in some of the calli. Carefully detach and separate the green tips in 2–3 parts using sterile forceps. Transfer each leaf or divided leaf section to fresh regeneration media and incubate in growth conditions mentioned in step 2 of Subheading 3.2.8 for 2 weeks. Usually 2–4 plantlets are developed per bombarded embryo. 4. For root induction, transfer the regenerated plantlets to rooting media (see item 4 under Subheading 2.2.4) and incubate in growth conditions mentioned in step 2 of Subheading 3.2.8 for 1–2 weeks. Plantlets of above 5 cm tall may be used for mutant screening. 5. Transfer the ~10 cm tall plantlets (see Note 19) with welldeveloped root system to Jiffy pots and incubate in high humidity (to acclimatize) conditions for 12–14 days in GM containment chamber. Make sure to rinse the roots gently with fresh water to remove media prior to transferring. As the plants grow taller, transfer them to larger pots (13-cm diameter) and grow in GM controlled chambers till maturity (which depends upon the variety).

3.2.9 Mutant Screening

1. Cut and pool (5–10 mg) leaf tissues from 3 to 4 plantlets obtained from step 4 of Subheading 3.2.8 and extract genomic DNA by CTAB method [25] or using plant DNA extraction kit (see Note 20). 2. Using common conserved primers, simultaneously PCR-amplify all the three genome targets as follows:

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Component

Final amount/concentration

Genomic DNA (see step 1 of Subheading 3.2.9)

40–50 ng

Master mix

1

Conserved primer F

1 μM

Conserved primer R

1 μM

Nuclease-free water

Up to 10 μL

3. Perform the PCR with following conditions: one step of denaturation at 94  C for 3 min; 35 cycles of denaturation at 94  C for 30 s, annealing at Tm for 30 s and extension at 72  C for 30 s followed by a final extension at 72  C for 2 min. 4. Separate 5 μL of PCR product on 1.2% agarose gel in TAE. Cut-off the single expected band and purify using gel extraction kit. 5. Digest the purified product using suitable restriction enzyme as per manufacturer’s instructions. 6. Separate the digested product on a 2.0% agarose gel in TAE buffer. Number of mutants should be 3–4 per 100 samples. 7. Select the plantlets from the pool that produced single band in PCR digestion (i.e., remain undigested) to fresh rooting media and grow for 5–7 days. If samples are sufficient for sampling, this step may be skipped. 8. Select individual plantlets showing positive results (single band after restriction) from the pool and repeat screening by PCR/RE assay using conserved primers as mentioned in steps 1–7 of Subheading 3.2.9. 9. Now amplify DNA sample of the mutants (showing single band in restriction) using target-specific primers. 10. Confirm the mutation type by sequencing as per standard protocol.

4

Notes 1. Use of Cas9 protein for genome editing eliminates the use of selection gene and possibilities of random genome integration. Such transgene-free plants would be more acceptable to public. Cas9 protein is nonheritable and thus prevents probable editing in subsequent generations. 2. Cold stratification improves germination rate in wheat.

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3. It is advised to grow the seeds at an interval of 1–2 weeks to harvest regular batches of immature embryos for bombardment. 4. A drop of mild surfactant like Tween-20 can also be added. 5. Gold particles are suitable because they are of homogeneous size and chemically inert to affect DNA and cell components. Appropriate size of particles is essential for adequate momentum to penetrate the embryo cells. 6. When preparing vitamin/amino acid for customized MS media, filter-sterilize using 0.2-μm filters. 7. In the entire tissue culture procedure no antibiotic is used for selection of transformed embryos. Selection-free method overcomes the tedious labor, time, and cost required for the optimization of antibiotic type and concentration which varies with the variety. Instead, a mixed-pool screening approach followed by high-throughput sequencing is implemented. 8. For HDR (homology directed repair), 50 μM of ssDNA oligos should be added in the reaction mixture. 9. Make sure the extracted DNA is of high quality, as enzymatic (nuclease) activity is highly affected by impurities. 10. Use PCR-amplified target from wild (nontransformed plant) as negative control. 11. T7EI nuclease identifies and cleaves mismatch in the doublestranded DNA. In Cas9 mutated sample, mismatch will be present in heteroduplex which is cleaved by T7EI endonuclease, while in the wild type sample there will be no mismatch in the duplex and hence not cleaved. 12. Immature embryos are the most appropriate target as they are competent for transformation, spreads evenly on media, and a variety of media have shown enhanced regeneration [31–33]. 13. Gold particles settles quickly, mix by pipetting before aliquoting each tube to ensure homogeneous distribution of particles. 14. Avoid keeping the macrocarrier on any vibrating surfaces, as it may lead to agglomeration (aggregation of particles). 15. Particle bombardment procedure needs to be performed under sterile conditions. Therefore, the gun’s chamber and its components are sterilized by spraying 70% (v/v) ethanol, while, macrocarriers, rupture discs and other parts are sterilized using absolute ethanol (for not more than 5 min) and allow to dry for ~5 min. 16. In each biolistic experiment, bombarded and nonbombarded controls should be used to check the efficiency of regeneration. Nonbombarded controls are used to monitor regeneration of donor cells (immature embryo). Bombarded controls (gold

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INDEX A Acetohydroxy acid synthase (AHAS) ............................. 14 Agargel® ..............................................232, 233, 240, 242 Agrobacterium ......................................................... 13, 40, 91, 108, 126, 164, 177, 201, 217, 230, 251, 264, 282, 296, 309 Allergy.............................................................................. 13 Alpha-amylase...................................................11, 15, 118 Arabidopsis thaliana ....................................................... 52 Ascorbic acid ........................................................ 231–233 Axial element (AE)....................................................22, 25

B Bar (bialaphos resistance) .................................... 179, 190, 242, 252, 254, 257, 258 Barley ................................................................... 4, 23, 45, 85, 132, 150, 202, 265, 282 B-, C-, and γ-hordeins .................................................... 11 Binary vectors ....................................................41, 43, 51, 52, 164, 178, 283, 284, 287 Biolistic delivery .................................................. 129, 131, 134, 135, 144–146, 148, 150, 153, 154, 177–193, 202–203, 309–328 Biolistics........................................................ 45, 108, 125, 145, 163, 177, 198, 217, 229, 251, 264, 288, 296, 309 Biolistic transformation .......................................... 45, 46, 110–113, 118, 127, 129, 131, 163–175, 251, 252, 256, 257, 259, 263–277, 288, 310, 312 Bionanotechnology ..................................... 141, 142, 144 Biotechnology .................................................39, 97, 135, 141, 142, 154, 198 Bivalents................................................. 20, 21, 25–29, 31 Bombyx mori ..............................................................23, 26 Bouquet stage.................................................................. 24 Brewery.......................................................................... 7, 8

C Calcium alginate beads ................................................... 48 Callus induction medium .................................... 220, 286 Carbon nanotubes (CNTs) ...........................50, 144, 146 Carlsberg Laboratory................................ 5–8, 12, 14, 24 Caryopsis ....................................................................... 283

Cauliflower mosaic virus (CaMV).......................... 41, 74, 165, 179, 203, 252, 283 Celiac disease ................................................................... 13 Celiac-safe wheat .......................................................13, 14 Cell suspension...................................................... 50, 126, 127, 131, 164, 218, 219 Chiasmata ..............................................20, 27, 28, 30, 31 Chitosan (CS)................................................................ 150 Chloroplast transformation .................................... 46, 47, 53, 118, 145 Chromosome pairing .................................. 12, 20, 22, 24 Clustered regularly interspaced short palindromic repeats/CRISPR associated protein 9 (CRISPR/ Cas9) ..................................................9, 13, 54–56, 198, 199, 205–209, 272, 295, 297, 309–328 Copper sulfate .....................................180, 201, 232, 242 Coprinus.................................................23, 25, 26, 28, 30 Creeping bentgrass (Agrostis stolonifera) .................................... 81, 91, 251–256 Cre–lox system ........................................................ 54, 153 CRISPR interference (CRISPRi) ................................... 56 CRISPR RNA (crRNA) .................................55, 205, 206

D DEMETER ............................................................. 13, 272 Developmentally regulated..........................70, 73, 91–99 Dicamba........................................................253–255, 288 2,4-Dichlorophenoxy acetic acid (2,4-D)............................................ 165, 180, 181, 200, 220, 223, 225, 231, 233, 288, 299, 303, 314 Diolistics ........................................................................ 297 Discosoma sp. red fluorescent protein (DsRed) .................................................... 299–301 Diter von Wettstein (DvW)........................... 3–16, 19–32 DNA delivery ....................................................43, 47, 49, 51, 110, 148, 164–166, 177–193, 197–209, 229, 251–259, 296, 305, 306, 324 Double-strand break (DSB) ................................... 27, 30, 46, 55, 56, 205, 206, 209, 273, 310 Dre2 ........................................................... 9, 13, 272–276 Droplet Digital PCR ..................................................... 225 Drosophila melanogaster............................................23, 30 Dynamic light scattering (DLS) ................................... 148

Sachin Rustgi and Hong Luo (eds.), Biolistic DNA Delivery in Plants: Methods and Protocols, Methods in Molecular Biology, vol. 2124, https://doi.org/10.1007/978-1-0716-0356-7, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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332 Index

IN

PLANTS: METHODS

AND

E Electrocompetent cells......................................... 285, 287 Electron microscopy (EM) ................................ 21, 22, 24 Electroporation-mediated plant transformation (EPT) ......................................... 51 Embryogenic callus ............................................. 113, 132, 171, 189, 198, 199, 202, 218–221, 226, 235, 239, 248, 253 Endochitinase ..............................................................9, 12 Endosperm ........................................................ 11, 13–15, 80, 84–88, 110, 186, 204, 244, 272, 302 Epigenetics ................................................................11, 13 Explants ............................................................... 164, 186, 198, 217, 230, 252, 264, 283, 325 Expression control ............................................70, 74, 81, 84–86, 97, 131

F Food safety ...................................................................... 13 Furled leaf ............................................................. 219, 220 Fusarium ............................................................. 12, 84, 91

G Gene delivery................................................ 42, 130, 132, 144, 163, 217–226 Gene editing ........................................................ 230, 244, 310, 316, 319 Gene-gun............................................................. 181, 297, 298, 300, 302 Genetic engineering ............................................ 7, 16, 40, 52, 118, 205, 252, 264 Genetic transformation ..................................... 13, 39–57, 107–118, 141–154, 164, 177, 198–200, 202, 229–249, 252, 256, 264, 265, 267, 282, 284, 295, 309 Genome editing ..............................................55–57, 126, 134, 135, 153, 164, 178, 198, 205–208, 296, 310, 311, 326 β-glucanase ...................................................................... 12 β-glucans.......................................................................... 15 β-glucuronidase (GUS)................................ 70, 131, 165, 179, 188, 189, 204, 244, 283, 291, 306 Glufosinate ammonium (PPT)........................... 232, 242, 248, 252, 253, 257, 258 Gluten ..................................................................... 13, 272 Glutenases........................................................................ 13 GM/GMO ................................................... 16, 230, 235, 240, 249, 325 Gold microcarriers .............................................. 234, 236, 237, 246 Gold nanoparticles (AuNPs) ..................... 142, 146–148, 150, 153, 311, 314, 324 Grain storage proteins .................................................... 11

PROTOCOLS Green fluorescent protein (GFP) ..............................9, 70, 80, 86, 87, 95, 132, 133, 150, 152, 153, 244, 284, 286–288, 304, 305 Guide RNA (gRNA) ............................................. 56, 206, 273, 275, 310, 315, 323

H Hi II genotype ....................................115, 179, 190, 299 Homoeologs............................................ 27–29, 272, 273 Homologous recombination (HR)........................ 20, 45, 47, 54–55, 272, 273 Hordeum vulgare....................................... 23, 56, 87, 284 Hygromycin......................................................... 165, 167, 171–173, 175, 203, 226, 242, 252, 286, 289 Hygromycin phosphotransferase gene (hph) ............... 252

I Imidazolinone ................................................................. 14 Indole-3-butyric acid (IBA) ............................... 200, 201, 231, 233 Interlockings................................................ 21, 23, 25, 28 Intracellular delivery ................................... 295, 296, 302 In vitro ...............................................................39, 40, 49, 111, 113, 117, 127, 164, 174, 277, 310, 312, 313, 318–320, 323

L Lipids and liposomes .................................................... 142 L7 Macrosalts ....................................................... 231, 232 L Microsalts .......................................................... 231–233

M Macrocarrier holders .................................. 168, 181, 185, 187, 192, 193, 223, 234, 237–239, 270, 289, 290, 300, 303 Macrocarriers........................................................ 108, 168 Magenta vessels ................................................... 231, 233, 235, 240, 241 Magnetic iron oxide nanoparticles (MIONPs) ................................................ 142, 146 Mature embryos ............................................................ 164 Meiosis ...............................................................20, 23, 24, 28, 31, 32, 81 Mesoporous silica nanoparticles (MSNs) ........................................... 144, 146–148, 150, 152, 153, 297 Microbiolistics ............................................. 150, 151, 153 Microbombardment.....................................125, 133–135 Microcarrier launch assembly (MLA) ................ 185, 187, 188, 192, 193, 223, 237, 238 Microcarriers .............................................. 108, 116, 118, 187, 192, 221–223, 226, 236–238, 246–248, 256, 270

BIOLISTIC DNA DELIVERY Microinjection-mediated plant transformation .......51, 52 Micro-particle bombardment............................. 125–127, 130–132, 134 Microprojectiles..................................................... 46, 108, 125, 163, 168–170, 181, 183–185, 187, 188, 198, 223, 251, 269–271, 297, 300, 303, 304 Microspore embryogenesis........................................... 277 Modified MS vitamins......................................... 179, 181, 182, 231, 232 Molecular breeding ....................................................... 199 MS vitamins .......................................................... 231, 233 Multivalents ........................................................ 22, 27–29 Murashige and Skoog (MS) medium ................ 116, 167, 200, 219, 220, 231, 254, 255, 314

N Nanobiolistics ...............................................135, 141–154 Nanoparticles................................................ 50, 135, 142, 202, 296, 314 Neomycin .................................................... 203, 226, 295 Neomycin phosphotransferase ............................ 221, 295 Nonhomologous end joining (NHEJ) ........................................ 55, 56, 205, 310 Nontransgenic plants .................................................... 153 Northern blot................................................................ 258 Nuclear localization signals (NLS)..................44, 45, 206 Nucleic acid delivery ................................... 134, 142, 309 Nutritional quality .......................................................... 13 N6 vitamin stock .................................179, 181, 299, 300

O Organelle transformation ........................... 118, 153, 221 Oryza sativa......................................................56, 86, 165 Osmotic medium for maize.......................................... 299

P Pairing homoeologous 1 (Ph1) ...................................28, 29 Particle bombardment ................................. 42, 108, 127, 177, 202, 222, 229, 251, 264, 290, 309 PDS-1000/He particle delivery system ............ 163, 168, 177, 178, 181, 185, 189, 236, 256 Phosphinothricin tripeptide (PPT) .................... 232, 233, 239, 240, 242, 248, 252–254, 257, 258 Photosynthetically active radiation (PAR) ......................................................... 26, 233, 248, 249, 313 Picloram................................................................ 232, 233 Plant genetic engineering ...................................... 55, 282 Plant genetic transformations..................................39–57, 107–118, 144–146, 154, 163, 198, 202 Plantlets ........................................................ 92, 113, 115, 152, 166, 171, 175, 190, 193, 198, 201, 203, 208, 219, 224, 235, 239–241, 249, 271, 277, 286, 289, 325, 326

IN

PLANTS: METHODS

AND

PROTOCOLS Index 333

Plasmids ........................................................ 40, 108, 129, 144, 164, 177, 217, 270, 281, 304, 312 Pollen tube-mediated transformation (PTT) ..........49, 50 Polyethylene glycol (PEG) ................................... 47, 109, 126, 229, 264, 296, 313 Polymerase chain reaction (PCR) ........................ 40, 129, 133, 164, 173, 219, 224–226, 240, 249, 258, 273, 274, 276, 312, 313, 315, 317–320, 322, 324, 326 Proanthocyanidin ............................................... 14, 15, 80 Programmable nucleases..............................264, 309–328 Promoters ..........................................................11, 44, 69, 127, 165, 179, 203, 218, 252, 272, 283, 312 Protein delivery ...................................142, 154, 295–306 Proteolistics ..................................................135, 295–306 Protospacer Adjacent Motif (PAM) ..................... 56, 206, 310, 311, 315, 320 Pseudomonas ......................................................... 9, 13, 91 Pyrrolnitrin ..................................................................9, 13

R Recombination nodules............................................21, 30 Reduced-immunogenicity ...................................... 11, 14, 272–276 Regeneration medium (RZP5), )....................... 189, 201, 233, 240, 241, 254, 255, 265, 269, 271, 286, 314, 325 Repeat Variable Di-residues (RVDs).............................. 55 Rhizoctonia ..................................................................... 12 Ribonucleoproteins (RNPs) ............................... 135, 142, 145, 178, 206, 230, 244, 295, 297, 310–326, 328 RNA interference (RNAi) .................................... 13, 134, 199, 204, 205 Rooting medium (RP5)...................................... 166, 168, 171, 174, 233, 235, 240, 241, 249, 271 Rupture disc retaining cap (RDRC) .................. 185, 187, 188, 192, 193, 223, 237, 238, 247, 248 Rupture discs .............................................. 187, 234, 237, 238, 243, 246–248, 327

S Saccharomyces cerevisiae....................................23, 47, 127 Scutellum ............................................................. 170, 178, 186–188, 235, 244, 286, 299, 302, 303, 323 Selectable markers ........................................ 54, 165, 178, 179, 190, 191, 202, 203, 209, 218, 221, 222, 234, 240, 242, 245, 248, 252, 257, 283 Silicon carbide (SiC) whisker....................................49, 50 Silver nitrate ............................................... 117, 180, 181, 232, 242, 299, 300, 303 Single-guide RNA (sgRNA)................................. 56, 206, 209, 272, 310–313, 315–318, 320, 323 Single-walled carbon nanotubes (SWCNTs) ................................................ 144, 146

BIOLISTIC DNA DELIVERY

334 Index

IN

PLANTS: METHODS

AND

Small interfering RNA (siRNA) ......................... 129, 135, 144, 204 Sonication .............................................................. 48, 245, 264, 265 Sordaria macrospora........................................................ 25 Sorghum .................................................. 52, 56, 197–209 Southern blot ....................................................... 225, 258 Spermidine.................................................. 153, 180, 184, 221, 222, 226, 234, 236, 243, 246, 256, 266, 270, 290 Stopping screens......................................... 168, 174, 177, 179, 181, 185, 187, 188, 192, 193, 221, 223, 234, 237–239, 247, 248, 289, 290, 300, 302, 303 Sugarcane.........................................................45, 54, 118, 197, 204, 217–226, 251, 259 Synaptonemal complex (SC) ....................................21–31

T Terminal deoxynucleotide transferase dUTP nick end labeling (TUNEL) ............................................ 150 Tissue culture ....................................................39, 49, 52, 53, 57, 107, 110–113, 115–117, 164, 165, 179, 190, 191, 198–202, 206, 207, 224, 225, 230, 239, 241, 242, 245, 249, 258, 264, 266, 277, 325, 327 Tissue-specific....................................13, 69–97, 205, 272 Topoisomerase ................................................................ 25 Totipotency ............................................. 39, 40, 164, 198 Trans-activating CRISPR RNA (tracrRNA).................................................. 56, 205 Transcription activator-like effector nucleases (TALEN) ..................................................... 54, 55, 134, 205, 310 Transferred DNA (T-DNA) ....................................40–45, 47, 54, 108, 113, 117, 130, 282, 309 Transformation efficiency (TE) .............................. 47, 48, 50–52, 54, 112, 113, 115, 116, 145, 148, 150, 153, 154, 173, 179, 198, 202, 209, 217–219, 226, 230, 243, 245, 248, 265, 283, 292 Transgenes ........................................................... 9, 13, 15, 43, 47, 48, 51, 54, 69, 77, 84, 86, 88, 91, 94, 97, 107–109, 126–132, 135, 150, 153, 164, 173, 174, 178, 203, 204, 219, 224, 225, 258, 259

PROTOCOLS Transgenic plants...............................................39, 41, 42, 49–51, 56, 70, 77, 79, 81, 85, 91, 94, 95, 98, 108, 109, 111, 112, 118, 125, 127, 150, 164–166, 198, 203, 207, 225, 241, 245 Transient gene expression....................... 46, 73, 125–135 Transposome ........................................................ 295, 297 Trichoderma..................................................................... 12 Triticum aestivum .............................................23, 27, 56, 85, 229–249, 263, 289, 310, 313 Tumour-inducing principle (TIP).................................. 40 Tungsten....................................................... 46, 108, 149, 154, 163, 177, 202, 221, 226, 303 Type-II callus ...................................................... 178, 186, 189, 190, 193, 256

U Ultrastructural analysis ................................................... 27

V vir genes ............................................................. 42–44, 52 Virus-induced gene silencing (VIGS) ................... 41, 134

W Wheat.................................................................11, 45, 79, 112, 126, 145, 198, 229, 251, 263, 282, 310 WUS and BABY BOOM ................................................. 52

X β-xylanase.....................................................................9, 14

Y Yeasts................................7, 12, 23, 29, 31, 46, 285, 312

Z Zea mays............................................................23, 56, 113 Zeatin........................................................... 200, 201, 233 Zeta potential (ZP) ....................................................... 148 Zinc-finger nucleases (ZFN) .................................. 54, 55, 205, 310