Transgenic Wheat, Barley and Oats: Production and Characterization Protocols

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Transgenic Wheat, Barley and Oats

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

For other titles published in this series, go to www.springer.com/series/7651

METHODS

IN

MOLECULAR BIOLOGY™

Transgenic Wheat, Barley and Oats Production and Characterization Protocols

Edited by

Huw D. Jones and Peter R. Shewry Centre for Crop Genetic Improvement, Rothamsted Research, Hertfordshire, UK

Editors Huw D. Jones Centre for Crop Genetic Improvement Rothamsted Research Harpenden Hertfordshire, AL5 2JQ, UK

Peter R. Shewry Centre for Crop Genetic Improvement Rothamsted Research Harpenden Hertfordshire, AL5 2JQ, UK

ISBN: 978-1-58829-961-1 e-ISBN: 978-1-59745-379-0 ISSN: 1064-3745 e-ISSN: 1940-6029 DOI: 10.1007/978-1-59745-379-0 Library of Congress Control Number: 2008933591 © Humana Press, a part of Springer Science+Business Media, LLC 2009 All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Humana Press, c/o Springer Science + Business Media, LLC, 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. While the advice and information in this book are believed to be true and accurate at the date of going to press, neither the authors nor the editors nor the publisher can accept any legal responsibility for any errors or omissions that may be made. The publisher makes no warranty, express or implied, with respect to the material contained herein. Cover illustration: Derived from Figure 1B in Chapter 3 Printed on acid-free paper 9 8 7 6 5 4 3 2 1 springer.com

Preface Understanding the physical and genetic structure of cereal genomes and how defined coding and non-coding regions interact with the environment to determine a phenotype are key to the future of plant breeding and agriculture. The production and characterisation of transgenic plants is a powerful reverse genetic strategy increasingly used in cereals research to ascribe function to defined DNA sequences. However, the techniques and resources required to conduct these investigations have, until recently, been difficult to achieve or totally lacking in wheat, barley and oat. This book brings together the latest protocols for the transformation, regeneration and selection using both biolistic and Agrobacterium tumefaciens appropriate for these three species. It includes two chapters describing in vitro Agrobacterium co-cultivation, one leading to germ line transformation with no need for tissue culture-based regeneration. In addition, it has several chapters dedicated to the manipulation of gene expression and characterisation of the recombinant locus and transgenic plants. Finally, it tackles the issues of GM risk assessment, field trials and substantial equivalence in terms of transcriptomics, proteomics and metabolomics. Although this book is dedicated to the temperate small grain cereals wheat, barley and oats, many of the techniques described could be readily adapted for other cereals or plants generally. We thank all the contributing authors for their timely and informative chapters, the staff of Humana Press, especially John Walker for their guidance, and Helen Jenkins for her proof-reading, word processing and administrative support.

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

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PART I. INTRODUCTION 1. Transgenic Wheat, Barley and Oats: Production and Characterization. . . . . . . Paul A. Lazzeri and Huw D. Jones

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PART II. TRANSFORMATION AND REGENERATION 2. Selection of Transformed Plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Huw D. Jones and Caroline A. Sparks 3. Reporter Genes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Alison Huttly 4. Biolistics Transformation of Wheat. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Caroline A. Sparks and Huw D. Jones 5. Agrobacterium-Mediated Transformation of Bread and Durum Wheat Using Freshly Isolated Immature Embryos. . . . . . . . . . . Huixia Wu, Angela Doherty, and Huw D. Jones 6. Floral Transformation of Wheat . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sujata Agarwal, Star Loar, Camille Steber, and Janice Zale 7. Highly Efficient Agrobacterium-Mediated Transformation of Wheat via In Planta Inoculation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Thierry Risacher, Melanie Craze, Sarah Bowden, Wyatt Paul, and Tina Barsby 8: Barley Transformation Using Biolistic Techniques . . . . . . . . . . . . . . . . . . . . Wendy A. Harwood and Mark A. Smedley 9. Barley Transformation Using Agrobacterium-Mediated Techniques . . . . . . . Wendy A. Harwood, Joanne G. Bartlett, Silvia C. Alves, Matthew Perry, Mark A. Smedley, Nicola Leyland, and John W. Snape 10. Transformation of Oats and Its Application to Improving Osmotic Stress Tolerance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Shahina B. Maqbool, Heng Zhong, Hesham F. Oraby, and Mariam B. Sticklen PART III. GENE AND PROTEIN EXPRESSION 11. Promoter Sequences for Defining Transgene Expression . . . . . . . . . . . . . . . Huw D. Jones and Caroline A. Sparks 12. Down-Regulation of Gene Expression by RNA-Induced Gene Silencing. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Silvia Travella and Beat Keller

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PART IV. CHARACTERISATION OF TRANSGENIC PLANTS 13. Gene Insertion Patterns and Sites. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Philippe Vain and Vera Thole 14. Fluorescent In Situ Hybridization to Detect Transgene Integration into Plant Genomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Trude Schwartzacher 15. Establishing Substantial Equivalence: Transcriptomics . . . . . . . . . . . . . . . . . María Marcela Baudo, Stephen J. Powers, Rowan A.C. Mitchell, and Peter R. Shewry 16. Establishing Substantial Equivalence: Proteomics . . . . . . . . . . . . . . . . . . . . . Alison Lovegrove, Louise Salt, and Peter R. Shewry 17. Establishing Substantial Equivalence: Metabolomics .. . . . . . . . . . . . . . . . . . Michael H. Beale, Jane L. Ward, and John M. Baker 18. Design and Management of Field Trials of Transgenic Cereals . . . . . . . . . . . Zoltán Bedo˝, Mariann Rakszegi, and László Láng 19. GM Risk Assessment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Penny A.C. Sparrow

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273 289 305 315

PART V. CONCLUSIONS 20. Transgenic Wheat, Barley and Oats: Future Prospects. . . . . . . . . . . . . . . . . . . 333 Jim M. Dunwell Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 347

Contributors SUJATA AGARWAL • Department of Plant Sciences, University of Tennessee, Knoxville, TN, USA SILVIA C. ALVES • Department of Crop Genetics, John Innes Centre, Colney, Norwich, UK JOHN M. BAKER • National Centre for Plant and Microbial Metabolomics, Rothamsted Research, Harpenden, Hertfordshire, UK TINA BARSBY • NIAB, Cambridge, UK JOANNE G. BARTLETT • Department of Crop Genetics, John Innes Centre, Colney, Norwich, UK MARÍA MARCELA BAUDO • Centre for Crop Genetic Improvement, Department of Plant Sciences, Rothamsted Research, Harpenden, Hertfordshire, UK MICHAEL H. BEALE • National Centre for Plant and Microbial Metabolomics, Rothamsted Research, Harpenden, Hertfordshire, UK ZOLTÁN BEDO˝ • Agricultural Research Institute of the Hungarian Academy of Sciences, Hungary SARAH BOWDEN • NIAB, Cambridge, UK MELANIE CRAZE • NIAB, Cambridge, UK ANGELA DOHERTY • Centre for Crop Genetic Improvement, Department of Plant Sciences, Rothamsted Research, Harpenden, Hertfordshire, UK JIM M. DUNWELL • School of Biological Sciences, University of Reading, Reading, Berkshire, UK WENDY A. HARWOOD • Department of Crop Genetics, John Innes Centre, Colney, Norwich, UK ALISON HUTTLY • Centre for Crop Genetic Improvement, Department of Plant Sciences, Rothamsted Research, Harpenden, Hertfordshire, UK HUW D. JONES • Centre for Crop Genetic Improvement, Department of Plant Sciences, Rothamsted Research, Harpenden, Hertfordshire, UK BEAT KELLER • Institute of Plant Biology, University of Zürich, Zürich, Switzerland LÁSZLÓ LÁNG • Agricultural Research Institute of the Hungarian Academy of Sciences, Hungary PAUL A. LAZZERI • Agrasys S.L., Barcelona, Spain NICOLA LEYLAND • Department of Crop Genetics, John Innes Centre, Colney, Norwich, UK STAR LOAR • Department of Plant Sciences, University of Tennessee, Knoxville, TN, USA ALISON LOVEGROVE • Centre for Crop Genetic Improvement, Department of Plant Sciences, Rothamsted Research, Harpenden, Hertfordshire, UK SHAHINA B. MAQBOOL • Department of Biology, Syracuse University, Syracuse, NY, USA

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ROWAN A.C. MITCHELL • Centre for Crop Genetic Improvement, Department of Biomathematics and Bioinformatics, Rothamsted Research, Harpenden, Hertfordshire, UK HESHAM F. ORABY • Department of Horticulture, Michigan State University, East Lansing, MI, USA WYATT PAUL • Biogemma, Clermont-Ferrand Cedex 2, France MATTHEW PERRY • Department of Crop Genetics, John Innes Centre, Colney, Norwich, UK STEPHEN J. POWERS • Centre for Mathematical and Computational Biology, Department of Biomathematics and Bioinformatics, Rothamsted Research, Harpenden, Hertfordshire, UK M ARIANN RAKSZEGI • Agricultural Research Institute of the Hungarian Academy of Sciences, Hungary THIERRY RISACHER • Biogemma, Clermont-Ferrand Cedex 2, France LOUISE SALT • Structuring Food for Health Group, Institute of Food Research, Colney, Norwich, UK TRUDE SCHWARZACHER • Department of Biology, University of Leicester, Leicester, UK PETER R. SHEWRY • Centre for Crop Genetic Improvement, Department of Plant Science, Rothamsted Research, Harpenden, Hertfordshire, UK MARK A. SMEDLEY • Department of Crop Genetics, John Innes Centre, Colney, Norwich, UK JOHN W. SNAPE • Department of Crop Genetics, John Innes Centre, Colney, Norwich, UK CAROLINE A. SPARKS • Centre for Crop Genetic Improvement, Department of Plant Sciences, Rothamsted Research, Harpenden, Hertfordshire, UK PENELOPE A.C. SPARROW • John Innes Centre, Colney, Norwich, UK CAMILLE STEBER • USDA/ARS Wheat Genetics, Quality, Physiology and Plant Disease Unit, Pullman, WA, USA MARIAM B. STICKLEN • Department of Crop and Soil Sciences, Michigan State University, East Lansing, MI, USA VERA THOLE • Department of Crop Genetics, John Innes Centre, Colney, Norwich, UK SILVIA TRAVELLA • Institute of Plant Biology, University of Zürich, Zürich, Switzerland PHILIPPE VAIN • Department of Crop Genetics, John Innes Centre, Colney, Norwich, UK JANE L. WARD • National Centre for Plant and Microbial Metabolomics, Rothamsted Research, Harpenden, Hertfordshire, UK HUIXIA WU • Genetic Resources and Enhancement Unit, CIMMYT, Mexico DF, Mexico JANICE ZALE • Department of Plant Sciences, University of Tennessee, Knoxville, TN, USA HENG ZHONG • SABRI, Research Triangle Park, NC, USA

Chapter 1 Transgenic Wheat, Barley and Oats: Production and Characterization Paul A. Lazzeri and Huw D. Jones Abstract Ever since the first developments in plant transformation technology using model plant species in the early 1980s, there has been a body of plant science research devoted to adapting these techniques to the transformation of crop plants. For some crop species progress was relatively rapid, but in other crop groups such as the small grain cereals, which were not readily amenable to culture in vitro and were not natural hosts to Agrobacterium, it has taken nearly two decades to develop reliable and robust transformation methods. In the following chapters of this book, transformation procedures for small grain cereals are presented, together with methods for gene and protein expression and the characterization of transgenic plants. In this introductory chapter we try to put these later chapters into context, giving an overview of the development of transformation technology for small grain cereals, discussing some of the pros and cons of the techniques and what limitations still exist. Key words: Small grain cereals, transformation, biolistics, Agrobacterium, tissue culture, regeneration, selection, promoters, reporter genes.

1. Development of Biolistic Transformation Methodology and Application in Small Grain Cereals

Biolistic transformation methods, involving the coating of DNA onto microscopic particles and the subsequent shooting of these particles into the cells of living tissue, were first applied in plants in the late 1980s (1), and in 1989 the first fertile transgenic maize plants produced by this method were reported (2) (Fig. 1). This demonstration was a major turning point in the field of genetic modification of cereal crops, similar to the demonstration in the early 1980s that Agrobacterium tumefaciens

Huw D. Jones and Peter R. Shewry (eds.), Methods in Molecular Biology, Transgenic Wheat, Barley and Oats, vol. 478 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-379-0_1

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Fig. 1. Time line of first reports of transformation of major cereal species. Biolistic (black), Agrobacterium (grey).

could be used to generate stable transgenic plants in those crop species susceptible to infection (3). The advent of biolistic transformation was very significant to cereal genetic modification at the time because a decade of attempts to transform cereal species with Agrobacterium had not resulted in the production of transformed plants, and it was generally believed that cereals were highly recalcitrant to Agrobacterium and that T-DNA insertion did not occur (4). It has subsequently been shown that this incompatibility does not exist, and Agrobacterium-mediated transformation protocols have been developed for all the major cereals (Fig. 1) (discussed in Section 3 and in Chapters 5–7 and 9 in this book), but in the early 1990s the primary obstacle to the application of GM techniques in cereals was the lack of efficient transformation technology (5). The lack of success in transforming cereals with Agrobacterium had stimulated research on the development of a range of alternative direct gene transfer (DGT) techniques, including imbibition, macroand microinjection of DNA, electroporation, electrophoresis, ultrasound and laser-mediated DNA uptake, the use of silicon carbide microfibres and protoplast transformation. DNA transfer into cells was demonstrated using all these techniques, and

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electroporation, silicon carbide fibres and protoplast transformation were used to generate populations of transgenic plants, particularly in rice and maize, but they were dependent on specific types of cell culture systems and were highly technically demanding (6). In contrast, it was recognized immediately that biolistic transformation gave the potential to deliver DNA to many different cell types, including those tissues explanted directly from plants and in vitro cultured cells. This offered the possibility of targeting gene delivery to cell types known to be able to regenerate plants, such as embryogenic callus cultures or embryonic tissues such as scutellum. This was particularly important for the small grain cereals such as barley, wheat and oats, in which it is very difficult to establish regenerable cell suspension cultures which were the preferred target for other DGT methods. A further characteristic of biolistic transformation that was quickly recognized was that it was highly suited to comparative studies of transient expression, in which the level of expression from different gene constructs could easily be compared by bombarding replicate cultures and subsequently assessing the expression of a scorable marker gene either by a biochemical assay or by counting expression spots for a visual marker such as beta-glucuroniduase (GUS). Transient expression assays were an important tool in the early development of stable transformation protocols for cereals, as they allowed the optimization of the various technical parameters for DNA delivery into target tissues and the identification of promoter sequences that gave high levels of expression. In the first years of biolistic transformation, a number of different particle acceleration systems were tested, including devices that used explosive charges, high-pressure pulses of nitrogen, helium or air, or electrostatic discharge (7). Most of these instruments were made in private workshops and were not commercially available, making it difficult to compare and transfer results between different research groups. However, when the PDS1000/He particle gun became commercially available from BioRad, it was found to be the most versatile and efficient machine, and it became accepted as the standard device (8). In the early experiments on the biolistic transformation of small grain cereal crops, the target tissue was generally callus cultures, following the results in maize from the bombardment of embryogenic suspension cells (2). In these experiments, parameters affecting gene delivery assessed by transient expression assays were analyzed (9, 10) and stably transformed callus lines and some plants were recovered (11, 12), but it was recognized that callus cultures were not the ideal target material, as their regeneration potential was not high and decreased with successive subcultures. Instead of callus cultures, several laboratories switched to immature embryo or scutellum cultures as target material. The use of

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these primary embryogenic cultures resulted in the production of fertile transgenic plants by several groups in the early 1990s in wheat (13–15) and barley (16, 17, 18), although in oats the first transgenic plants were recovered from bombarding callus cultures (19) (Fig. 1). Immature embryo or scutellum cultures remain the preferred target material for biolistic transformation of wheat and barley, largely because they have the highest regeneration capacity of the different culture systems available. They have the disadvantage that donor plants must be grown through flowering to early seed development to maintain a supply of target explants and this has prompted the investigation of alternative target tissues. Transgenic plants have been recovered from the bombardment of a number of other explant and culture types, including microspore cultures (19, 20), immature inflorescences (21, 22), mature seed-derived cultures (23, 24), leaf base cultures (25) and meristem cultures (26). Although several of these target tissues, such as seed-derived cultures or meristem cultures, are much more convenient to use than immature embryos or scutella, the levels of transformation efficiency obtained are generally significantly reduced and these systems are not widely used. Since the initial demonstrations of transgenic wheat, barley and oat plants from biolistic transformation, two of the main areas of research have been to increase the efficiency of transformation systems, as well as to achieve reliable high-throughput procedures suitable for applied genetic modification projects and to develop procedures applicable to broader ranges of genotypes, including elite varieties. In the first area, significant progress has been made by the analysis and optimization of the component parameters of culture and regeneration procedures (27, 28), bombardment conditions (29, 30) and selection protocols (31, 32). In the second area of developing procedures applicable in elite cereal varieties, progress has been made in developing optimized protocols that allow the transformation of germplasm not previously pre-selected for response (33, 34), but genotypic limitation on transformation efficiency is still a major factor in cereal transformation. This aspect is discussed in Section 6. Biolistic transformation was the first method to deliver transgenic plants in the small grain cereals and it has been used extensively throughout for more than a decade of basic and applied research and is a now a standard capability in laboratories working in cereal biotechnology. At the level of production of transgenic lines intended for commercialization as varieties, it has the disadvantage compared to Agrobacterium-mediated transformation that it tends to produce more complex transgene integration patterns (discussed in Section 4), which may lead to instability in gene expression and heritability, but it remains an important research tool.

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2. Development of Agrobacterium Transformation Methodology and Application in Small Grain Cereals

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The notion that plants could be engineered by introducing defined lengths of foreign DNA was born partly from the discovery that bacteria of the genus Agrobacterium possessed the ability to transfer genes responsible for crown gall disease. It is now 100 years since this soil bacterium was implicated as the causative agent of crown gall symptoms (35), although the molecular basis of this inter-kingdom gene transfer was not elucidated until 70 years later (36). Crown gall disease is caused by the transfer of a portion of the Ti plasmid (the T-DNA) into host tissue and the expression of genes encoding auxins, cytokinins and enzymes that direct the synthesis of amino acids or sugar derivatives resulting in plant cell proliferation and tumorous growth (reviewed in (37–40)). As well as the T-DNA itself, two other genetic regions are necessary for successful tumour formation. Chromosomally located chv and att genes play a role in recognition of and binding to susceptible plant cells and several virulence (vir) genes, carried on the Ti plasmid, are required for T-DNA processing and transfer. A breakthrough in the use of A. tumefaciens as a tool for genetic engineering came with the observation that no physical linkage between the T-DNA and the other genetic components was needed (41, 42). Thus, binary vector systems were developed in which the Ti plasmid was rendered non-oncogenic (disarmed) by removing the phytohormone biosynthetic genes, and smaller (binary) plasmids, capable of replicating in both E. coli and Agrobacterium, were designed to carry the T-DNA with convenient multiple cloning sites for the insertion of foreign DNA (43–47). However, the removal of the oncogenes from the T-DNA resulted in the loss of a scoreable phenotype which hitherto had been used to indicate successful transfer and expression of bacterial genes (48, 49). Thus, the use of binary vectors and disarmed Agrobacterium strains necessitated the development new marker genes to select successful transformation events. The first to be utilized was the neo (nptII) gene encoding neomycin phosphotransferase II and conferring the ability of plant cells to grow on medium containing aminoglycoside antibiotic G418 (50, 51), although others soon followed, including genes conferring resistance to the antibiotics hygromycin, methotrexate or the herbicide bialaphos (40) (see Chapter 2). Early attempts at plant transformation utilized isolated Ti plasmids physically transferred into protoplasts or the in vitro transformation of cultured cells or tumours using the expression of opines or antibiotic resistance genes with plant promoters as markers (50, 52–54). The first transgenic plants that possessed a recombinant T-DNA, which was not only expressed but also inherited by its progeny, were reported in 1984 (55, 56).

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Five years later, 25 plant species had been transformed using A. tumefaciens (reviewed in (57), although the only monocot reported was asparagus (58), and cereals were generally considered to be outside the host range for Agrobacterium. Factors thought to be important for successful cereal transformation such as the presence of vir genes on helper or binary plasmids, explant type, inclusion of vir gene inducers, etc. were investigated and optimized. By 1995, there was good evidence of heritable Agrobacterium transformation in rice (59, 60) and maize (61) with reports of transgene expression or tumour formation in over 20 other monocot species (reviewed in (62)). A decade of further improvements to factors such as Agrobacterium strain/binary vector combination, inoculation and co-cultivation conditions has led to successful Agrobacterium transformation of Japonica, Indica and Javanica rice, spring and winter varieties of wheat and barley, hybrid maize, sorghum, rye and several forage and turf grasses (reviewed in (63)) (Fig. 1): although to date we are unaware of any reports of Agrobacterium transformation of oat or millet.

3. Pros and Cons of Direct Versus Agrobacterium DNA Delivery

The merits and limitations of Agrobacterium and physical/direct methods of DNA delivery fall into two categories. First, and more significant, are the considerations of molecular genetic properties exhibited by the transgenic events generated, and second, there are the differing requirements of the protocols themselves, in terms of specialist equipment and expertise required, genotype dependency, throughput and efficiency of transformation, etc. It is generally accepted that, irrespective of the DNA delivery method, transgenes integrate into the plant nuclear genome through illegitimate recombination, aided to some extent by the cell’s own DNA repair machinery, although micro-homology has been implicated in some T-DNA integrations (64) and many data indicate a preference for transcriptionally active regions of the genome (reviewed in (65–67)). Many studies of transgene molecular integration patterns have been carried out and some general trends are emerging. However, differences between experiments in terms of plant species transformed, Agrobacterium strain/vector combinations used, bombardment parameters and tissue culture protocols, etc. make direct comparisons difficult. It is clear that both DNA delivery methods can produce plants with multiple copies of rearranged transgene. However, many authors highlight the advantages of Agrobacterium transformation in terms of generally more simple integration patterns (reviewed

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in (65, 68–70)). For instance, in a direct comparison between rice plants obtained using Agrobacterium or biolistics, it was found that the average number of transgene copies was 1.8 and 2.7, respectively (71). In addition, the percentage of plants containing intact cassettes was higher and the line-to-line variation in GUS expression was lower with Agrobacterium (71). A similar comparison in barley revealed that all the Agrobacterium-generated lines possessed between one and three transgene copies, while 60% of the lines derived by particle bombardment integrated more than eight copies (72). In maize, Agrobacterium-mediated transformants had lower transgene copies and higher, more stable gene expression than their bombardment-derived counterparts (73). Analysis of wheat Agrobacterium lines in different laboratories showed that between 30 and 68% were single-copy events (74–78). In contrast, molecular analysis of wheat lines obtained using biolistics showed a tendency for high transgene copy number and insertional rearrangements (79, 80), an observation also seen in oat (81, 82). Counterbalancing these observations are the numerous reports of unwanted, binary vector backbone DNA found in plants generated using Agrobacterium. In wheat, approximately two-thirds of the lines analysed contained some DNA outside the T-DNA borders (75), with equivalent values for barley of 48% (83) and 45–70% in the various reports for rice (84– 86). However, suppression of non-T-DNA backbone sequences in rice has been demonstrated by the use of a binary vector with multiple left border repeats (87). In an alternative approach to ‘clean’ transformation, extraneous backbone sequence can be physically removed from the DNA prior to using the direct delivery method. For instance, linear, minimal expression cassettes have been introduced into rice using biolistics, which were found to give simpler integration events when compared to equivalent, supercoiled plasmid lines (88, 89). As can be seen in the various chapters in this book, the transformation protocols for cereals are all relatively complex and specialized, regardless of whether biolistics or Agrobacterium are used to deliver the DNA. However, significant differences exist with implications for throughput or efficiencies. Depending on the vectors systems available, the cloning of a candidate gene into an appropriate binary vector, and the transfer of this vector into a suitable Agrobacterium strain, inevitably involves more steps compared with the usual practice of co-bombarding the candidate gene and selectable marker cassette on separate plasmids. However, once the Agrobacterium strain is made, scaling up transformation throughput is relatively easy compared with the operation of the biolistic device which can handle up to only 50 explant pieces per shot and represents a bottleneck. An advantage of biolistics is that it is a physical process that exploits common DNA repair mechanisms and does not rely on the specific biological

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interactions necessary for successful T-DNA transfer between Agrobacterium and host plant cell. For many species, Agrobacterium is only suitable for a limited range of genotypes, whereas biolistics is more genotype independent and can be used to transfer DNA to a wide range of single cell or tissue types including those traditionally considered as hard to reach such as dividing cells in the apical meristem (reviewed in (90)).

4. Pros and Cons of Germ Line Versus Tissue Culture Based Methods

As discussed in Sections 1–3, the first robust and reliable transformation methods that were developed for small grain cereals were based on the use of particle bombardment or Agrobacterium to deliver genes into cells of embryogenic tissue cultures, followed by the regeneration of plants from transgenic cells. While these methods can work at efficiencies suitable for large-scale commercial-crop GM programmes, they do have some inherent disadvantages, which means that there is continued interest in developing alternative transformation methods. Their principal disadvantage is the reliance on good tissue culture response from cereal varieties to be transformed, i.e. it must be possible to initiate regenerable cultures at an acceptable frequency and the response of these cultures must be sufficiently robust to tolerate the stresses of gene delivery and selection of transformants without losing regeneration capacity. As discussed in Section 6, there is very significant genotypic dependence on culture response in cereals, the practical result of which is that still today many elite varieties cannot be transformed at acceptable frequencies. A second disadvantage of culture-based transformation methods is the potential for unwanted culture-induced variation (somaclonal variation) among regenerated plants. Plants regenerated from cultured somatic tissues may show a range of variation including chromosomal aberrations, point mutations, activation of transposable elements, epigenetic variation and physiological disturbances (5). The incidence of this variation can be reduced by targeting primary explants, minimizing culture time and other measures, but there still exists a background frequency of ‘off’ plants among populations of transformants. To avoid tissue culture, considerable effort has been devoted to the development of methods for the direct transformation of germ line cells such as meristem cells, microspores or ovules that would develop directly into plants. Limited success was achieved until it was demonstrated in Arabidopsis that it was possible to generate transformants by germinating seeds in the presence of Agrobacterium (91), and subsequently that infiltration of floral

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tissues with Agrobacterium could also produce transformants (92). More recently, in planta transformation methods, targeting germ line cells in seeds or developing inflorescences, have been shown to work in wheat ((93–95) and Chapter 6). This is an important development, as germ line transformation methods potentially offer several advantages over culture-based transformation. Genotypic dependence on regeneration capability and somaclonal variation among transformants should not be limitations with these methods, and the labour-intensive steps of target explant preparation and culture initiation are also avoided. These methods may also have the potential to be performed under clean glasshouse rather than aseptic laboratory conditions, although, paradoxically, the requirements for work to be performed under GM containment conditions may be easier to achieve in the laboratory than in a glasshouse. Despite the advantages of in planta transformation methods, it is still too early to say if they will prove generally applicable. There is so far not enough experience to know whether there will also be genotypic limitations and what efficiency levels will routinely be achieved. In addition, with inflorescence transformation, plant growth environment and developmental stage will influence efficiency and it may not be facile to grow plants under the conditions required.

5. GenotypeDependency, Regeneration and TransformationEnhancing Genes

Genotypic variation in response in vitro and in regeneration capacity is seen in all plant cell culture systems and this clearly has implications for the efficiency of tissue-culture-based transformation technology. As a group, the cereal species were long regarded as being very difficult to manipulate and regenerate in vitro (5), and although regeneration systems starting from a range of explants, including immature embryos, leaf and meristem tissues and, more recently, mature seeds have been developed for all of the major cereal species, in all cases there are genotypic restrictions on the application of these systems and they are generally really efficient only in a limited range of amenable genotypes. Genotypic dependence on regeneration efficiency, in turn, limits the application of transformation technology because in all tissue-culture-based transformation systems, using either Agrobacterium or DGT, a primary factor that influences transformation efficiency is culture response and regeneration capacity. Much effort has been expended in optimizing tissue culture and transformation procedures and efficiencies have steadily increased, but genotype dependence remains an important limitation. This factor is of lesser importance for research applications, where

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it may be acceptable to work in amenable ‘model’ genotypes, but for commercial applications elite varieties must be targeted. The approach taken in many agbiotech companies to cope with genotype dependency is to establish high-throughput transformation pipelines using amenable genotypes and subsequently to cross selected events into elite breeding lines for further development. While this provides a workable solution, there is still strong interest in reducing genotypic dependence; this is a driver for the development of in planta transformation techniques to avoid genotype effects on culturability in vitro and for research towards understanding the factors controlling genotypic variation in regeneration and transformation efficiency. The genetics of regeneration response in culture has been examined in several cereal species, and in particular in rice and maize QTLs (quantitative trait loci) for culture initiation and regeneration have been identified (96, 97). Isogenic lines for QTLs affecting culture response have been developed and the potential for breeding for culture response is clear (96). In some cases, the major genes have been identified from culture response QTLs; in rice, for example, variation in the expression of a ferritin-nitrite reductase gene was shown to underlie the difference in regeneration capability between responsive and non-responsive varieties (98). In parallel with whole plant genetic approaches, gene expression profiling is being used to identify key genes involved in the processes of tissue culture initiation and regeneration. Most progress has been made to date in Arabidopsis, where numbers of genes showing differential expression during shoot and root morphogenesis have been identified (99) and candidate genes involved in the regulation of these processes are being evaluated (100). Similar approaches are being followed in cereal species to identify genes involved in regeneration in culture (101). If such genes can be identified, they may be useful for the development of markers for regeneration capability, and it may be possible to improve the regeneration and transformation potential of elite breeding lines by their introgression, either by crossing or via transformation.

6. Regulating Gene Expression Using Heterologous Promoters

One of the challenges of using a transgenic approach to modify a plant phenotype is to utilize cis-regulatory elements that result in the desired patterns of gene expression. In the broadest sense, there are many regulatory features of a gene that effect expression in cis, including enhancer elements that may be located far up or down stream, the promoter itself including the core sequences, response elements, etc., 5´ and 3´ untranslated regions (UTRs), introns,

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polyadenylation signals, etc. However, more work has been done on the characterization of promoter and intron sequences than the other features, and in practice the choice of promoter is the major single factor in determining transgene expression. It is tempting to assume that the promoter of a gene with a well-defined expression pattern in one species can be used to mimic, via transformation, that same expression profile in another species. However, while there are many examples of cross-species, or even cross-kingdom promoter activities, there can be major, unpredictable changes in the specific expression patterns. It is common practice to test new promoter configurations in transformation experiments using reporter gene constructs (see Chapter 3) prior to using them for trait manipulation. Chapter 11 describes in detail the current state of knowledge regarding constitutive, tissue-specific and inducible promoters for cereal transformation and includes a table of all promoters known to have been previously used transgenic wheat, oats or barley.

7. Substantial Equivalence An issue that is inherent to the use of genetic modification technology to alter cereal crop traits is the possibility that apart from the expression of the inserted genes other unintended changes in gene expression may be caused, which might have unwanted effects, either at the level of agronomic performance or on nutritional characteristics. There are possible mechanisms by which transgene insertion may induce unintended variation; current transformation methods, either Agrobacterium-based or particle bombardment, involve random insertion into the recipient genome, which may disrupt native genes, or the expression of the transgene may have pleiotropic effects on endogenous gene expression. Our current experience from the past 25 years of production of transgenic plants is that the frequency of such effects is very low, as would be expected, and the potential for pleiotropic effects as a result of genetic modification is not a phenomenon that is confined to transgenic technology but may also occur in conventional breeding. However, transgenic crops are subjected to a level of scrutiny not applied to varieties produced by other genetic modification methods such as chemical or radiationinduced mutagenesis, which may induce widespread genomic disruption, but whose use in plant breeding over nearly a century has not resulted in important food safety problems. In particular, the safety of transgenic food stuffs has been targeted by NGOs and other groups fundamentally opposed to the use of transgenic crops in agriculture, and this has raised the level of concern among consumer groups and regulatory bodies.

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The need for standardized evaluation methods for ensuring the safety of foods derived from GM crops was recognized in the early 1990s and was considered by the World Health Organization/ Food and Agricultural Organization (WHO/FAO) (1991) (102) and the Organization of Economic Co-operation and Development (OECD), and in 1993 the OECD published a document entitled “Safety Evaluation of Foods Derived from Modern Biotechnology: Concepts and Principles” (103), which proposed the concept of ‘substantial equivalence’ as an approach to the assessment of the safety of foodstuffs derived from GM crops. The basis of substantial equivalence is that if the composition of a novel (GM) foodstuff does not differ significantly from that of the same type of foodstuff derived from standard crop varieties, then there is no reason to consider that the novel foodstuff is unsafe. There has been considerable debate on the definition and interpretation of substantial equivalence and as to its suitability as the basis of evaluation of the safety of GM foods (104, 105), and the way in which the concept has been applied in European Union (EU) food safety assessments and by the Food and Drug Administration in the United States is rather different (106). However, substantial equivalence is now generally accepted as the starting point for novel food safety evaluation, with the proviso that it is not a food safety assessment per se, but an approach to evaluate differences between a novel food and its standard counterpart, to be followed up by further analyses as needed to establish the extent and significance for food safety of those differences detected, with particular focus on constituents known to have implications for health, such as allergenic or toxic molecules (106). One of the early criticisms applied the use of substantial equivalence in the evaluation of the safety of GM foods was that the biochemical and nutritional profiles of many conventional foods was not well established, and surprisingly little was understood of the extent of variation between different varieties and to what extent environmental effects modified food composition. At the technical level, unless an approach of analysing certain selected groups of food constituents (storage proteins, for example) was taken, it was technically very difficult to use standard biochemical assay techniques to look for changes in hundreds of different metabolites. In recent years, however, there have been very significant advances in technologies for large-scale profiling of biological samples at the levels of gene expression (transcriptome), protein expression (proteome) and metabolites (metabolome) (covered in detail in Chapters 16–18), which have made it possible to make comparisons between novel and reference samples at the ‘global’ level, rather than by the analysis of changes in selected groups of genes, proteins or metabolites. These new profiling techniques, and in particular metabolic profiling using

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nuclear magnetic resonance (NMR) methodology, are now being adopted as the primary approach for the evaluation of substantial equivalence between standard and GM crops and the foodstuffs derived from them (107). In the cereals, recent publications demonstrate their application in maize (108, 109), rice (110) and wheat (107).

8. Use of Selectable and Reporter Genes

The low efficiencies of most plant transformation methods require a system for identifying and facilitating the survival of those few cells that incorporate the transgene over the vast majority that do not. Such selectable markers can be grouped depending upon whether they confer positive or negative selection characteristics and whether they are conditional on the presence of additional media components. Commonly used genes are those that encode enzymes conferring the ability to grow in the presence of antibiotics or herbicides, but many different selection strategies have been employed in the production of transgenic cereals, which are reviewed in (111, 112) and discussed further in Chapter 2. Non-selectable, reporter or scorable marker genes are commonly used in transformation experiments to visually detect transgenics, to optimize protocols or to study the effect of specific regulatory sequences on gene expression. Commonly used visual reporter genes are uidA, Luc and gfp, encoding the enzymes beta-glucuronidase, luciferase and the green fluorescent protein, respectively, although others have been also applied successfully. The merits of these reporter genes in the context of small grain cereals are described in Chapter 3.

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88. Fu, X. D., Duc, L. T., Fontana, S., Bong, B. B., Tinjuangjun, P., Sudhakar, D., Twyman, R. M., Christou, P. and Kohli, A. (2000) Linear transgene constructs lacking vector backbone sequences generate low-copynumber transgenic plants with simple integration patterns. Transgen Res. 9, 11–19. 89. Agrawal, P. K., Kohli, A., Twyman, R. M. and Christou, P. (2005) Transformation of plants with multiple cassettes generates simple transgene integration patterns and high expression levels. Mol Breed. 16, 247–260. 90. Altpeter, F., Baisakh, N., Beachy, R., Bock, R., Capell, T., Christou, P., Daniell, H., Datta, K., Datta, S., Dix, P. J., Fauquet, C., Huang, N., Kohli, A., Mooibroek, H., Nicholson, L., Nguyen, T. T., Nugent, G., Raemakers, K., Romano, A., Somers, D. A., Stoger, E., Taylor, N. and Visser, R. (2005) Particle bombardment and the genetic enhancement of crops: myths and realities. Mol Breed. 15, 305–327. 91. Feldmann, K. A. and Marks, M. D. (1987) Agrobacterium-mediated transformation of germinating seeds of Arabidopsis thaliana – a non-tissue culture approach. Mol GenGenet. 208, 1–9. 92. Bechtold, N., Ellis, J. and Pelletier, G. (1993) In planta Agrobacterium-mediated Ge.e-transfer by infiltration of adult Arabidopsis thaliana plants. Comptes Rendus De L Academie Des Sciences Serie Iii-Sciences De La Vie-Life Sciences 316, 1194–1199. 93. Craze, M. and Risacher, T. (2000) Plant Transformation Method. Patent No. WO 00/63398. 94. Zale, J. M. and Steber, C. M. (2006) In planta transformation of wheat as a genomics tool, in Proceedings of the Plant and Animal Genomics XIV Conference, Jan 14–18, 2006, San Diego, USA. 95. Supartana, P., Shimizu, T., Nogawa, M., Shioiri, H., Nakajima, T., Haramoto, N., Nozue, M. and Kojima, M. (2006) Development of simple and efficient in Planta transformation method for wheat (Triticum aestivum L.) using Agrobacterium tumefaciens. JBiosciBioeng. 102, 162–170. 96. Taguchi-Shiobara, F., Yamamoto, T., Yano, M. and Oka, S.(2006)Mapping QTLs that control the performance of rice tissue culture and evaluation of derived near-isogenic lines. TheorApplGenet. 112, 968–976. 97. Krakowsky, M. D., Lee, M., Garay, L., Woodman-Clikeman, W., Long, M. J., Sharopova, N., Frame, B. and Wang, K. (2006) Quantitative trait loci for callus initiation and totipotency in maize (Zea mays L.). Theor Appl Genet. 113, 821–830.

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98. Nishimura, A., Ashikari, M., Lin, S., Takashi, T., Angeles, E. R., Yamamoto, T. and Matsuoka, M. (2005) Isolation of a rice regeneration quantitative trait loci gene and its application to transformation systems. P. N.A.S.USA 102, 11940–11944. 99. Che, P., Lall, S., Nettleton, D. and Howell, S. H. (2006) Gene expression programs during shoot, root, and callus development in Arabidopsis tissue culture. Plant Physiol. 141, 620–637. 100. DeCook, R., Lall, S., Nettleton, D. and Howell, S. H. (2006) Genetic regulation of gene expression during shoot development in Arabidopsis. Genetics172, 1155–1164. 101. Che, P., Love, T. M., Frame, B. R., Wang, K., Carriquiry, A. L. and Howell, S. H. (2006) Gene expression patterns during somatic embryo development and germination in maize Hi II callus cultures. Plant MolBiol. 62, 1–14. 102. Joint FAO/WHO Consultation on the Assessment of Biotechnology in Food Production and Processing as Related to Food Safety (1990) Geneva S, Strategies for assessing the safety of foods produced by biotechnology: report of a joint FAO/WHO consultation, Geneva, 5–10 November 1990. 103. OECD (1993) Organisation for Economic Co-operation and Development. Safety Evaluation of Foods Derived by Modern Biotechnology – Concepts and Principles, OECD, Paris. 104. Kuiper, H. A., Kleter, G. A., Noteborn, H. and Kok, E. J. (2001) Assessment of the food safety issues related to genetically modified foods. Plant J. 27, 503–528. 105. Konig, A., Cockburn, A., Crevel, R. W. R., Debruyne, E., Grafstroem, R., Hammerling, U., Kimber, I., Knudsen, I., Kuiper, H. A.,

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Peijnenburg, A., Penninks, A. H., Poulsen, M., Schauzu, M. and Wal, J. M. (2004) Assessment of the safety of foods derived from genetically modified (GM) crops. Food ChemToxicol. 42, 1047–1088. Levidow, L., Murphy, J. and Carr, S. (2007) Recasting “substantial equivalence”: transatlantic governance of GM food. Science TechnolHuman Values32, 53–91. Baker, J. M., Hawkins, N. D., Ward, J. L., Lovegrove, A., Napier, J. A., Shewry, P. R. and Beale, M. H. (2006) A metabolomic study of substantial equivalence of fieldgrown genetically modified wheat. Plant Bio tech J. 4, 381–392. Manetti, C., Bianchetti, C., Bizzari, M., Casciani, L., Castro, C., D’Ascenzo, G., Delfini, M., Di Cocco, M. E., Lagana, A., Miccheli, A., Motto, M. and Conti, F. (2004) NMRbased metabonomic study of transgenic maize. Phytochem. 65, 3187–3198. Herman, R. A., Storer, N. P., Phillips, A. M., Prochaska, L. M. and Windels, P. (2007) Compositional assessment of event DAS59122-7 maize using substantial equivalence. Regul Toxicol Pharmacol. 47, 37–47. Oberdoerfer, R. B., Shillito, R. D., De Beuckeleer, M. and Mitten, D. H. (2005) Rice (Oryza sativa L.) containing the bar gene is compositionally equivalent to the nontransgenic counterpart. JAgrFood Chem. 53, 1457–1465. Miki, B. and McHugh, S. (2004) Selectable marker genes in transgenic plants: applications, alternatives and biosafety. JBiotechnol. 107, 193–232. Wilmink, A. and Dons, J. J. M. (1993) Selective agents and marker genes for use in transformation of monocotyledonous plants. Plant Mol Biol Report. 11, 165–185.

Chapter 2 Selection of Transformed Plants Huw D. Jones and Caroline A. Sparks Abstract The low frequency and randomness of transgene integration into host cells, combined with the significant challenges of recovering whole plants from those rare events, makes the use of selectable marker genes routine in plant transformation experiments. For research applications that are unlikely to be grown in the field, strong herbicide- or antibiotic resistance is commonly used. Here we use genes conferring resistance to glufosinate herbicides as an example of a selectable marker in wheat transformation by either Agrobacterium or biolistics. Key words: Selectable marker genes, selection, herbicide, antibiotic, Basta, bar, pat, glufosinate ammonium, G418.

1. Introduction Dominant selectable marker genes have been pivotal in the development of transformation systems in all organisms. Selectable genes encode proteins that enable the identification of rare events by allowing genetically transformed cells to grow preferentially compared to the vast majority of untransformed ones. Usually the selection system comprises two components: a chemical agent, such as a herbicide, an antibiotic or a specific carbon source which is added to the tissue culture medium; and a gene incorporated into the transformation cassette, conferring selective advantage under those particular media conditions. Selection agents and the relevant marker genes that are commonly used in cereal transformation are listed in Table 1. Traditionally, plant transformation methods utilized genes such as bar, epsps, nptII and hpt, which confer to regenerating cells the ability to survive otherwise lethal concentrations of herbicides such as those based on glufosinate Huw D. Jones and Peter R. Shewry (eds.), Methods in Molecular Biology, Transgenic Wheat, Barley and Oats, vol. 478 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-379-0_2

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or glyphosate, or antibiotics such as aminoglycosides. Recently, alternative strategies have been adopted using genes that give a metabolic advantage to the transformed cells by allowing preferential access to sugars or amino acid sources. These latter methods are often called ‘positive selection’ (see Note 2) and are considered more benign both for the health and safety of laboratory scientists and for the environment if the plants are grown in the field. An example of this type of selection is the pmi (phosphomannose isomerase) gene, which facilitates the utilization of mannose as a carbon source and prevents the inhibition of glycolysis caused by an accumulation of mannose-6-phosphate.

Table 1 Selectable marker genes used for the generation of stably transformed wheat, barley or oat plants (see Note 1 for full gene names) Species and DNA delivery method

Selectable marker gene

Selection agent

Sample references

Ubi1 plus first intron

L-Phosphinothricin: Bialaphos, Basta, etc.

(13, 14, 18, 19)

bar

CaMV 35S with or without Adh1 intron1

L-Phosphinothricin: Bialaphos, Basta etc.

(20, 21, 22)

hpt

CaMV 35S

Hygromycin B

(23)

CP4 epsps and gox

Duplicated CaMV 35S

Wheat biolistics bar

Promoter driving selectable marker gene

(24)

cah

Ubi1 plus first intron

Cyanamide

(25)

nptII

Enhanced CaMV 35S with HSP70 intron

G418

(26)

pmi (manA)

Ubi1 plus first intron and others

Mannose

(27)

nptII

Enhanced CaMV 35S with HSP70 intron

G418

(28)

CP4 epsps

Various promoters

Glyphosate

(29, 30)

hpt

CaMV 35S

Hygromycin B

(31)

bar

Ubi1 plus first intron

L-Phosphinothricin: Bialaphos, Basta, etc.

(17, 32, 33)

pmi (manA)

Ubi1 plus first intron and others

Mannose

(34, 35)

Barley PEG

nptII

Act1 plus first intron

G418

(36)

Barley CaCl2

nptII

CaMV 35S plus Adh1 intron1

G418

(37)

Wheat Agrobacterium

(continued)

Selection of Transformed Plants

25

Table 1 (continued) Species and DNA delivery method

Selectable marker gene

Selection agent

Sample references

Ubi1plus transit peptide from ssRbcs

Lysine plus threonine

(38)

bar

Ubi1 plus first intron

Bialaphos

(39–41)

bar

CaMV 35S with or without Adh1 intron1

Bialaphos/Basta

(42–43)

nptII

nos

G418

(44)

nptII

CaMV 35S

Dot blot at rooting stage

(45)

codA

Act1 plus first intron

5-Fluorocytosine

(46)

hpt

CaMV 35S plus Adh1 intron1

Hygromycin B

(47)

Plant promoter

Mannose

(34)

bar

Ubi1 plus first intron

Bialaphos

(48–50)

pat

CaMV 35S

Bialaphos

(51)

hpt

CaMV 35S plus hpt intron

Hygromycin B

(50, 52)

nptII

CaMV 35S plus Adh1 intron1

Paromomycin

(53, 54)

bar

CaMV 35S with or without Adh1 intron1

L-Phosphinothricin: Bialaphos, Basta, etc.

(55, 56)

bar

Ubi1 plus first intron

Bialaphos

(57)

hpt

Rice Act1 plus first intron

Hygromycin B

(58)

Barley biolistics lysC

Barley Agrobac- pmi (manA) terium

Oat biolistics

Promoter driving selectable marker gene

In a departure from conventional approaches, cereal transformations have been achieved using scorable markers such as GFP or luc either alone or in conjunction with conventional chemical selection (1–8). In addition, low numbers of transgenic wheat plants were obtained without the use of any selectable markers, relying instead on polymerase chain reaction (PCR) screening (9). Various chapters in this book describe biolistics or Agrobacterium transformation and mention the specific selection regime suitable for that species. Here, we provide a detailed description specifically on the selection steps using bar or pat marker genes (see Note 3) and the selection agent phosphinothricin (PPT), the active ingredient in several commercial glufosinate ammonium-based broad spectrum herbicides (see Note 4). We use wheat as an example, but this selection regime can easily be adapted for barley or oats (see Note 5).

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2. Materials 1. Suitable plasmid construct containing the bar or pat gene under the control of a strong promoter to ensure gene expression in tissue culture phase. A constitutive promoter will also allow selection to be applied at germination or adult growth stages (see Note 6) 2. Immature embryos that have been bombarded or have undergone co-cultivation with Agrobacterium (see Note 7) 3. Callus induction medium (×2) (see Note 8): 200 ml MS macrosalts (×10), 2 ml L7 microsalts (see (10) for list of components) (×1,000), 20 ml/l ferrous sulfate chelate solution (×100), 200 mg/l myo-inositol, 0.2 mg/l thiamine HCl, 1.0 mg/l nicotinic acid, 1.0 mg/l pyridoxine HCl. For biolistics induction medium add 750 mg/l L-glutamine (see Note 9), 150 mg/l L-proline, 100 mg/l L-asparagine, 180 g/l sucrose (see Note 10); or for Agrobacterium induction medium add 1 g/l L-glutamine, 200 mg/l casein hydrolysate, 3.9 g/l MES and 80 g/l maltose (see Note 11). Adjust pH to 5.7 with 5 M NaOH or KOH. Osmolarity should be within the range 800–1,100 mOsM. Filter sterilize (see Note 12) 4. Regeneration medium (×2) (see Note 8): 200 ml L7 macrosalts (×10), 2 ml L7 microsalts (see (10) for list of components) (×1,000), 20 ml/l ferrous sulfate chelate solution (×100), 400 mg/l myo-inositol, 20 mg/l thiamine HCl, 2 mg/l nicotinic acid, 2 mg/l pyridoxine HCl, 2 mg/l Ca-pantothenate, 2 mg/l L-ascorbic acid and 60 g/l maltose. Adjust pH to 5.7 with 5 M NaOH or KOH. Osmolarity should be within the range 269–298 mOsM. Filter sterilize (see Note 12) 5. Gelling agents (Sigma-Aldrich, Poole, UK): Agargel (×2), 10 g/l; or Phytagel (×2) 4 g/l. Autoclave to sterilize (see Note 13) 6. Glufosinate ammonium (PPT) (see Note 14): 10 mg/ml in water. Filter sterilize (see Note 12) 7. DNA extraction reagents and equipment (see Note 15) 8. PCR and electrophoresis apparatus and consumables. Primers to detect bar gene in transgenic plants: Bar1 GTC TGC ACC ATC GTC AAC C and Bar2 GAA GTC CAG CTG CCA GAA AC 9. Commercial herbicide (e.g. Basta, Challenge, Harvest) for leaf painting: dilute in 0.1% Tween 20 to give 0.2 and 2 g/l of active ingredient PPT (see Note 16)

Selection of Transformed Plants

27

3. Methods The methods described below have been developed over 10 years of usage and are based on work described in (11–17). The initiation of embryogenic callus from immature wheat scutella and regeneration of shoots (approximately 10 callus pieces per Petri dish) are performed in a tissue culture room at 22°C under a 12-h photoperiod (see Note 17). Selection pressure can be first applied during the callus initiation phase (considered as ‘early selection’) from the first round of regeneration, or more conventionally, from the second round of regeneration onwards (see Note 18). The regeneration response is evaluated every 3 or 4 weeks at each transfer cycle by scoring regenerants with only shoots, only roots or shoots and roots. When choosing a selection agent for a specific plant variety, it is important to test for natural resistance of the target plant material by first performing a kill curve experiment to establish the optimum concentration to use in transformation studies (see Note 19). 3.1. Growth and Selection of Transformed Plants

1. Establish callus by culturing immature scutella containing putatively transformed cells on callus induction medium supplemented with 0.5 mg/l 2,4-D (see Note 20) and 10 mg/l AgNO3 (see Note 21). For Agrobacterium-treated cultures, also add 2 mg/l Picloram and 160 mg/l Timentin (see Note 22). Incubation during the callus induction phase is carried out in the dark (see Note 17). Selection agent, glufosinate ammonium at 2–6 mg/l, can be added at this stage (see Note 18). 2. After 3 or 4 weeks (see Note 23), transfer callus cultures from induction medium to first round of regeneration medium supplemented with 0.1 mg/l 2,4-D (see Note 20), 25 mg/l CuSO4 (see Note 24) and 5 mg/l zeatin, and continue culture in the light (see Note 17). For Agrobacterium-treated cultures, also add 160 mg/l Timentin (see Note 22). Selection agent, glufosinate ammonium, at 2–6 mg/l can be added at this stage (see Note 18). 3. Evaluate the regeneration/selection response after 3–4 weeks (see Note 25). Transfer regenerating calli to a second round of regeneration medium supplemented with 5 mg/l zeatin (see Note 26) and 2–6 mg/l of the selection agent glufosinate ammonium and culture for a further 3–4 weeks (see Notes 27 and 28). Half of the tissues from a non-transformed control plate should be cultured on regeneration medium with selection and the other half without selection to provide selected and non-selected controls (see Note 7 and Fig. 1).

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Fig. 1. Wheat calli on regeneration medium without selection (top), with selection at 6 mg/l PPT (bottom).

4. Evaluate the regeneration and selection responses after 3–4 weeks (see Note 25). 5. To ensure a low percentage of escapes, transfer only sections with healthy, green shoots and/or plantlets to regeneration medium supplemented with 4–6 mg/l glufosinate ammonium in Magenta vessels (Sigma-Aldrich) (see Note 29) and culture for a further 3–4 weeks (see Note 30). 6. Once the plants are of sufficient size (10–15 cm) and have established a reasonable root system, transfer putative transgenic plants to compost in 8 cm square plastic pots (see Note 31) and grow on in a GM containment glasshouse (see Note 32). 3.2. Analysis of Transgenic Plants 3.2.1. PCR Analysis

1. When the plants are suitably established (three to four leaves), a leaf sample approximately 2 cm in length can be taken from which to extract genomic DNA (see Note 33). Place leaf samples immediately in liquid nitrogen. 2. Grind leaf samples to powder in liquid nitrogen and extract genomic DNA using, for example, the Wizard genomic DNA purification kit (see Note 15). 3. In order to establish whether the plant is transformed, the DNA is analysed by PCR using the bar gene primers 1 and 2 (see above) at an annealing temperature of 57°C.

Selection of Transformed Plants

29

4. Re-pot confirmed PCR positive plants into 13 cm diameter pots and continue growth in GM containment glasshouse (see Note 32). 3.2.2. Herbicide Leaf Paint Assay

Later in development, transgenic plants can be analysed further using the herbicide leaf paint assay (17). 1. Prepare dilutions from the herbicide stock (e.g. Challenge) using 0.1% Tween to give two different concentrations of the active ingredient PPT: 2 and 0.2 g/l (see Note 16). 2. For each plant to be tested, select three approximately equally sized, healthy-looking leaves, from separate tillers where possible, avoiding the flag leaf. Tag the chosen leaves on the stem immediately below to designate 0.2 g/l PPT, 2 g/l PPT and control (0.1% Tween) treatments (see Note 34). 3. Mark each leaf with biro half way along its length. Paint the upper surface of the distal half of the leaf with the appropriate solution using a cotton bud (see Note 35). 4. Leave the plants for 7 days for the herbicide to take effect before assessing for resistance/susceptibility. 5. Score each treated leaf according to the percentage desiccation suffered over the area painted with the herbicide solution, and the percentage of the proximal region of the leaf that has been affected by travel of the herbicide (see Fig. 2a). 6. A plant can be considered transgenic if no symptoms of injury are detected when 0.2 g/l and 2 g/l PPT are applied (see Fig. 2b). If a plant shows resistance at the lower concentration only, it should be confirmed to be transgenic by other means.

Fig. 2. (a) Diagram showing scoring for herbicide leaf paint assay. (b) Leaves painted with Challenge herbicide (2 g/l PPT) showing desiccation/beaching effect on distal portion of leaf (controls, left) compared to resistant leaf (right).

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Jones and Sparks

4. Notes 1. – bar – Basta resistance, encodes phosphinothricin acetyltransferase isolated from Streptomyces hygroscopicus. – pat – encodes phosphinothricin acetyltransferase isolated from Streptomyces viridochromogenes. – cah – encodes cayanamide hydratase isolated from a soil fungus Myrothecium verrucaria. – hpt (hph, aph-IV) – encodes hygromycin phosphotransferase isolated from Escherichia coli. – epsps – encodes 5-enolpyruvylshikimate-3-phosphate synthase (EPSPS) isolated from Agrobacterium sp. Strain CP4. – gox – encodes glyphosate oxidoreductase isolated from Ochrobactrum anthropi. – nptII – encodes neomycin phosphotransferase isolated from Escherichia coli transposon Tn5. – pmi (manA) – encodes phosphomannose isomerase isolated from Escherichia coli. – lysC – encodes a lysine feedback desensitised aspartate kinase-III (AK-III) isolated from Escherichia coli. – codA – encodes a cytosine deaminase isolated from Escherichia coli. 2. Many authors refer to the toxin-based strategies (e.g. antibiotic and herbicide) as ‘negative selection’ and the nutrient conversion strategies (e.g. sugar isomerase and alternative amino acids) as ‘positive selection’. However, this nomenclature clashes with others, particularly in the non-plant literature, which use the term positive selection to all approaches that promote survival of the transformed cells and reserve the term negative selection for approaches that kill the transformed cells. The latter definitions appear to be more widely used. 3. Two genes (from different Streptomyces species) that encode the enzyme phosphinothricin N-acetyltransferase have been used to confer resistance to PPT in transformation experiments. The Basta resistance (bar) gene was isolated from Streptomyces hygroscopicus and the pat gene was isolated from Streptomyces viridochromogenes. 4. The L-isomer of PPT is available in various commercial formulations, including Basta, Bialaphos, Ignite, Rely, Finale, Challenge, Liberty and Harvest.

Selection of Transformed Plants

31

5. Alternative varieties or species may require modifications to the media detailed here. Modifications may include the choice of basal salts (MS or L7), the type of sugars (sucrose or maltose) and their concentration and/or the type and level of hormones. 6. Many plasmids have been constructed to contain the bar or pat genes. Examples that have been widely used include: – pAHC20 and pAHC25 (59) which incorporate the bar gene under the control of the maize Ubi1 promoter + intron and nos terminator – pBARGUS (60) which incorporates the bar gene under the control of the CaMV 35S promoter plus Adh 1 intron and nos terminator – pDBI (22) which incorporates the bar gene under the control of the CaMV 35S promoter and CaMV 35S terminator. – pCB30 series (61), which are binary plasmids, that incorporate the bar gene under the control of the nos promoter and nos terminator – pAL156 and others in the pGreen series (62), which are binary plasmids, that incorporate the bar gene under the control of the maize Ubi1 promoter + intron and nos terminator 7. Various control plates should be included within each experiment: untreated to monitor the development/regeneration of donor tissue; treated but without DNA and unselected to monitor tissue culture response following transformation; and treated with DNA and selected to monitor the effects of the selection on regeneration. 8. All solutions should be made using reverse osmosis, polished water with a purity of 18.2 MΩ/cm. Sterile stock solutions of media components are prepared; those kept at 4°C can be stored for 1–2 months and those at −20°C should remain effective for at least a year, provided no freeze/thawing has occurred. 9. Dissolve L-glutamine at pH 9.0 before mixing with the other components. 10. Sucrose (9%) partially plasmolyses the cells and this may increase their ability to withstand bombardment. However, this is variety- and species dependent and 3% sucrose is often a suitable alternative, e.g. for T. turgidum ssp. durum scutella. 11. Although the induction medium described here for Agrobacterium-treated embryos differs in composition from that used for biolistics, the same induction medium can be used if more convenient, with similar success.

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12. Filter sterilization is carried out using 0.2 µm filter size. For large volumes use MediaKap (NBS Biologicals Ltd., Cambridgeshire, UK), and for smaller volumes use a Nalgene syringe filter (Fisher Scientific, UK). 13. Agargel is the gelling agent for the induction medium for culture of embryos from biolistics and for all regeneration media. Phytagel is used for the induction medium for culture of Agrobacterium-treated embryos. Both are prepared at double concentration, autoclaved at 121°C for 20 min and either used straight away or re-melted in a microwave oven before adding to filter-sterilized, double concentration medium. Supplements are added prior to pouring in Petri dishes or Magentas. To avoid non-uniform solidification and difficulties when remelting, the agargel or phytagel solution should be shaken well both before and after autoclaving. 14. Glufosinate ammonium is synthetically produced phosphinothricin (PPT) bound to ammonium (Greyhound Chromatography and Allied Chemicals, Cheshire, UK), and is the active component in herbicides such as Basta and Challenge. Bialaphos (phosphinothricylanalylanaline, sodium) (Melford Laboratories Ltd.) is a successful alternative selection agent when used at 3–5 mg/l. 15. Various kits are commercially available for extraction of genomic DNA. The Wizard genomic DNA purification kit (Promega) with one to two approximately 2 cm lengths of leaf material typically yields 10–50 µg DNA which is more than sufficient for PCR. However, if greater quantities of DNA are required, e.g. for Southern analysis, use the CTAB (cetyl-trimethylammonium bromide) method (63). 16. Different herbicides may have different concentrations of active ingredient (PPT) in the formulation: e.g. Basta 200 g/l and Challenge 150 g/l. The amount of herbicide diluted is therefore calculated accordingly to give 0.2 and 2 g/l concentrations of the active ingredient PPT. Gloves should be worn when dealing with these herbicides. 17. Incubation is carried out in a controlled-environment room with the room temperature set to give 22°C at the level of the cultures under the lights. A 12-h photoperiod is provided by cool white fluorescent tubes emitting lighting levels of approximately 250 µmol/m2/s PAR. For the callus induction phase, trays are covered with foil to provide darkness. 18. Selection is generally applied at the second round of regeneration and subsequent transfers. However, selection can be introduced earlier, i.e. at callus induction or at the first round of regeneration. Earlier selection can lead to fewer calli and/or plantlets requiring transfer at each stage but with the associated disadvantage that some transformants may be

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lost if they are not strong enough to survive selection early on. Lower concentrations of selective agent could therefore be used if included in the early stages. 19. The selection agent should be used at a concentration that is known to fully inhibit the growth of non-transformed explants. However, the concentration should be gauged according to the development of the cultures at each transfer stage. Generally, use within the range 2–6 mg/l glufosinate ammonium (PPT). 20. Picloram (Sigma-Aldrich) at 2–6 mg/l can be used as an alternative auxin (64, 65). 21. AgNO3 is added to the induction medium to promote embryogenesis but 10 mg/l silver thiosulfate (a mix of silver nitrate and sodium thiosulfate) can be used as an alternative. Both are photosensitive, so the stock solutions and any media plates containing them should be kept in the dark. 22. For cultures having undergone Agrobacterium co-cultivation, Timentin (Ticarcillin/Clavulanic (15:1), Melford Laboratories Ltd.) should be added to all media at 160 mg/l to prevent Agrobacterium overgrowth. 23. The induction period for somatic embryogenesis is usually 3–5 weeks; however, the explants should be observed regularly to check for contamination. Judgement and experience are required to monitor development in order to determine the best time for transfer to regeneration medium; transfer is carried out when the embryogenic callus has mature somatic embryos some of which may just be forming small shoots. At this stage, whole calli should be transferred without division, placing approximately 10 calli per plate. 24. Copper sulfate is a stress-inducing agent (similar to silver nitrate) used to promote shooting. The preferred copper sulfate concentration is 100 µM (25 mg/l), but if too much shooting occurs, 50 µM can be used. 25. Calli can be assessed for shoot and root formation at each subculture to monitor the response in tissue culture and the effects of selection once applied. 26. Zeatin may only need to be added during the first round of regeneration; it is not as critical at this second stage. 27. As most embryogenic calli give rise to shoots and some roots, it is important that the roots are pushed fully into the selection medium. This should lower the risk of escapes (i.e. plantlets forming that are not transgenic). Deeper dishes can be used at this stage by using the upturned base of another Petri dish as the lid. This provides greater height for growth of shoots. 28. The number of calli per 9-cm Petri dish can be reduced at this stage if the regenerating calli to be transferred are large.

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If a callus divides naturally, each of the callus pieces should be documented in order to trace plants with possible clonal origin. 29. If several plants are generated from same callus, this should be carefully recorded as the plants may be of clonal origin. 30. If many green plantlets are still surviving at this stage, the higher selection concentration of PPT can be used (6 mg/l). 31. Tissue cultured plantlets have little or no waxy cuticle, so are particularly prone to desiccation after transfer to soil. Place newly potted transgenic plants in a propagator for 1–2 weeks to maintain high humidity in order for the cuticle to develop. 32. Glasshouse conditions are 18–20°C day and 14–16°C night temperatures with a 16-h photoperiod provided by natural light supplemented with banks of Son T 400W sodium lamps (Osram Ltd.) giving 400–1,000 µmol/m2/s PAR. 33. The leaf sample should not be taken from an emerging leaf but from one already unfurled. 34. Chosen plants should be watered properly before herbicide application. 35. The application should be quite firm to ensure coating and some penetration of the solution into the leaf. All the control leaves should be painted first followed by the lower concentration, then the higher concentration of herbicide, to minimize carry over of the herbicide to other leaves from gloves, etc.

Acknowledgements Rothamsted Research receives grant-aided support from the Biotechnology and Biological Sciences Research Council (BBSRC) of the UK. References 1. Fang, Y. D., Akula, C. and Altpeter, F. (2002) Agrobacterium-mediated barley (Hordeum vulgare L.) transformation using green fluorescent protein as a visual marker and sequence analysis of the T-DNA :: barley genomic DNA junctions. J. Plant Physiol. 159, 1131–1138 . 2. Murray, F., Brettell, R., Matthews, P., Bishop, D. and Jacobsen J. (2004) Comparison of Agrobacterium-mediated transformation of four barley cultivars using the GFP and GUS reporter genes. Plant Cell Reports 22:397–402 3. Jordan M. C. (2000). Green fluorescent protein as a visual marker for wheat transformation. Plant Cell Reports 19, 1069–1075.

4. McCormac, A. C., Wu, H. X., Bao, M. Z., Wang, Y. B., Xu, R. J., Elliott, M. C. and Chen, D. F. (1998) The use of visual marker genes as cell-specific reporters of Agrobacterium-mediated T-DNA delivery to wheat (Triticum aestivum L.) and barley (Hordeum vulgare L.). Euphytica 99, 17–25. 5. Kaeppler, H. F., Menon, G. K., Skadsen, R. W., Nuutila, A. M. and Carlson, A. R. (2000) Transgenic oat plants via visual selection of cells expressing green fluorescent protein. Plant Cell Reports 19, 661–666. 6. Ahlandsberg, S., Sathish, P., Sun, C. X. and Jansson, C. (1999) Green fluorescent protein as a reporter system in the trans-

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formation of barley cultivars. Physiologia Plantarum 107, 194–200. Harwood, W. A., Ross, S. M., Bulley, S. M., Travella, S., Busch, B., Harden, J. and Snape, J. W. (2002) Use of the firefly luciferase gene in a barley (Hordeum vulgare) transformation system. Plant Cell Reports 21, 320–326. Huber, M., Hahn, R. and Hess, D. (2002) High transformation frequencies obtained from a commercial wheat (Triticum aestivum L. cv. ‘Combi’) by microbombardment of immature embryos followed by GFP screening combined with PPT selection. Mol. Breeding 10, 19–30. Permingeat, H. R., Alvarez, M. L., Cervigni, G. D. L., Ravizzini, R. A. and Vallejos, R. H. (2003) Stable wheat transformation obtained without selectable markers. Pl. Mol. Biol. 52, 415–419. Sparks, C. A. and Jones, H. D. (2004) Transformation of wheat by biolistics, in Transgenic Crops of the World - Essential Protocols, (Curtis, I. P., ed.), Kluwer Academic Publishers, Dordrecht, pp. 19–35. Barcelo, P., Rasco-Gaunt, S., Thorpe, C. and Lazzeri, P. A. (2001) Transformation and gene expression, in Advances in Botanical Research Incorporating Advances in Plant Pathology Vol. 34, (Shewry, P. R., Lazzeri, P. A. and Edwards, K. J., eds.), pp. 59–126. Goodwin, J., Pastori, G., Davey, M. and Jones, H. D. (2004) Selectable markers: antibiotic and herbicide resistance, in Transgenic Plants: Methods and Protocols, (Pena, L. ed.,), Humana Press, Totowa, NJ. Pastori, G. M., Wilkinson, M. D., Steele, S. H., Sparks, C. A., Jones, H. D. and Parry, M. A. J. (2001) Age-dependent transformation frequency in elite wheat varieties. J. Exp. Bot. 52, 857–863. Rasco-Gaunt, S., Riley, A., Cannell, M., Barcelo, P. and Lazzeri, P. A. (2001) Procedures allowing the transformation of a range of European elite wheat (Triticum aestivum L.) varieties via particle bombardment. J. Exp. Bot. 52, 865–874. Rasco-Gaunt, S., Riley, A., Lazzeri, P. and Barcelo, P. (1999) A facile method for screening for phosphinothricin (PPT)resistant transgenic wheats. Mol. Breeding 5, 255–262. Jones, H. D., Doherty, A. and Wu, H. (2005) Review of methodologies and a protocol for the Agrobacterium-mediated transformation of wheat. Plant Methods 1, 5. Wu, H., Sparks, C., Amoah, B. and Jones H. D. (2003) Factors influencing successful

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Agrobacterium-mediated genetic transformation of wheat. Plant Cell Rep. 21, 659– 668. Iser, M., Fettig, S., Scheyhing, F., Viertel, K. and Hess, D. (1999) Genotype-dependent stable genetic transformation in German spring wheat varieties selected for high regeneration potential. J. Pl. Physiol. 154, 509–516. Weeks, J. T., Anderson, O. D. and Blechl, A. E. (1993) Rapid production of multiple independent lines of fertile transgenic wheat (Triticum aestivum). Pl. Physiol. 102, 1077–1084. Vasil, V., Castillo, A. M., Fromm, M. E. and Vasil, I. K. (1992) Herbicide resistant fertile transgenic wheat plants obtained by microprojectile bombardment of regenerable embryogenic callus. Bio-Technol. 10, 667–674. Nehra, N. S., Chibbar, R. N., Leung, N., Caswell, K., Mallard, C., Steinhauer, L., Baga, M. and Kartha, K. K. (1994) Selffertile transgenic wheat plants regenerated from isolated scutellar tissues following microprojectile bombardment with two distinct gene constructs. Plant J. 5, 285–297. Becker, D., Brettschneider, R. and Lorz, H. (1994) Fertile transgenic wheat from microprojectile bombardment of scutellar tissue. Plant J. 5, 299–307. Ortiz, J. P. A., Reggiardo, M. I., Ravizzini, R. A., Altabe, S. G., Cervigni, G. D. L., Spitteler, M. A., Morata, M. M., Elias, F. E. and Vallejos, R. H. (1996) Hygromycin resistance as an efficient selectable marker for wheat stable transformation. Plant Cell Rep. 15, 877–881. Zhou, H., Arrowsmith, J. W., Fromm, M. E., Hironaka, C. M., Taylor, M. L., Rodriguez, D., Pajeau, M. E., Brown, S. M., Santino, C. G. and Fry, J. E. (1995) Glyphosate-tolerant CP4 and GOX genes as a selectable marker in wheat transformation. Plant Cell Rep. 15, 159–163. Weeks, J. T., Koshiyama, K. Y., MaierGreiner, U., Schaeffner, T. and Anderson, O. D. (2000) Wheat transformation using cyanamide as a new selective agent. Crop Sci. 40, 1749–1754. Pastori, G. M., Huttly, A., West, J., Sparks, C., Pieters, A., Luna, C. M., Jones, H. D. and Foyer, C. H. (2007) The maize Activator/Dissociation system is functional in hexaploid wheat through successive generations. Funct. Pl. Biol. 34, 835–843. Gadaleta, A., Giancaspro, A., Blechl, A. and Blanco, A. (2006) Phosphomannose iso-

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Jones and Sparks merase, pmi, as a selectable marker gene for durum wheat transformation. J. Cereal Sci. 43, 31–37. Cheng, M., Fry, J. E., Pang, S. Z., Zhou, H. P., Hironaka, C. M., Duncan, D. R., Conner, T. W. and Wan, Y. C. (1997) Genetic transformation of wheat mediated by Agrobacterium tumefaciens. Plant Physiol. 115, 971–980. Hu, T., Metz, S., Chay, C., Zhou, H. P., Biest, N., Chen, G., Cheng, M., Feng, X., Radionenko, M., Lu, F. and Fry, J. (2003) Agrobacterium-mediated large-scale transformation of wheat (Triticum aestivum L.) using glyphosate selection. Plant Cell Reports 21, 1010–1019. Cheng, M., Hu, T. C., Layton, J., Liu, C. N. and Fry, J. E. (2003) Desiccation of plant tissues post-Agrobacterium infection enhances T-DNA delivery and increases stable transformation efficiency in wheat. In Vitro Cell. Develop. Biol. - Plant 39, 595–604. Mitic, N., Nikolic, R., Ninkovic, S., MiljusDjukic, J. and Neskovic, M. (2004) Agrobacterium-mediated transformation and plant regeneration of Triticum aestivum L. Biologia Plantarum 48, 179–184. Wu, H., Doherty, A. and Jones, H. D. (2007) Efficient and rapid Agrobacteriummediated transformation of durum wheat (Triticum turgidum L. ssp durum) using additional virulence genes. Transgen. Res. In Press. Khanna, H. K. and Daggard, G. E. (2003) Agrobacterium tumefaciens-mediated transformation of wheat using a superbinary vector and a polyamine-supplemented regeneration medium. Plant Cell Rep. 21, 429–436. Reed, J., Privalle, L., Powell, M. L., Meghji, M., Dawson, J., Dunder, E., Suttie, J., Wenck, A., Launis, K., Kramer, C., Chang, Y. F., Hansen, G. and Wright, M. (2001) Phosphomannose isomerase: an efficient selectable marker for plant transformation. In Vitro Cell. Develop. Biol. - Plant 37, 127–132. Wright, M., Dawson, J., Dunder, E., Suttie, J., Reed, J., Kramer, C., Chang, Y., Novitzky, R., Wang, H. and Artim-Moore, L. (2001) Efficient biolistic transformation of maize (Zea mays L.) and wheat (Triticum aestivum L.) using the phosphomannose isomerase gene, pmi, as the selectable marker. Plant Cell Rep. 20, 429–436. Funatsuki, H., Kuroda, H., Kihara, M., Lazzeri, P. A., Muller, E., Lorz, H. and Kishinami, I. (1995) Fertile transgenic bar-

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bacterial codA- and cytochrome P450 genemediated selection. Plant J. 19, 719–726. Hagio, T., Hirabayashi, T., Machii, H. and Tomotsune, H. (1995) Production of fertile transgenic barley (Hordeum vulgare L.) plant using the hygromycin-resistance marker. Plant Cell Rep. 14, 329–334. Tingay, S., McElroy, D., Kalla, R., Fieg, S., Wang, M. B., Thornton, S. and Brettell, R. (1997) Agrobacterium tumefaciens-mediated barley transformation. Plant J. 11, 1369–1376. Travella, S., Ross, S. M., Harden, J., Everett, C., Snape, J. W. and Harwood, W. A. (2005) A comparison of transgenic barley lines produced by particle bombardment and Agrobacterium-mediated techniques. Plant Cell Rep. 23, 780–789. Shrawat, A. K., Becker, D. and Lorz, H. (2007) Agrobacterium tumefaciens-mediated genetic transformation of barley (Hordeum vulgare L.). Plant Sci. 172, 281–290. Kumlehn, J., Serazetdinova, L., Hensel, G., Becker, D. and Loerz, H. (2006) Genetic transformation of barley (Hordeum vulgare L.) via infection of androgenetic pollen cultures with Agrobacterium tumefaciens. Plant Biotech. J.l 4, 251–261. Wang, M. B., Abbott, D. C., Upadhyaya, N. M., Jacobsen, J. V. and Waterhouse, P. M. (2001) Agrobacterium tumefaciens-mediated transformation of an elite Australian barley cultivar with virus resistance and reporter genes. Aust. J. Pl. Physiol. 28, 149–156. Torbert, K. A., Rines, H. W. and Somers, D. A. (1995) Use of paromomycin as a selective agent for oat transformation. Plant Cell Rep.s 14, 635–640. Torbert, K. A., Rines, H. W. and Somers, D. A. (1998) Transformation of oat using mature embryo-derived tissue cultures. Crop Sci. 38, 226–231. Somers, D. A., Rines, H. W., Gu, W., Kaeppler, H. F. and Bushnell, W. R. (1992) Fertile transgenic oat plants. Bio-Technol. 10, 1589–1594. Maqbool, S. B., Zhong, H., El-Maghraby, Y., Ahmad, A., Chai, B., Wang, W., Sabzikar, R. and Sticklen, M. B. (2002) Competence of oat (Avena sativa L.) shoot apical meristems for integrative transformation, inher-

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Chapter 3 Reporter Genes Alison Huttly Abstract Reporter genes have been widely used in plant molecular biology, typically to discern patterns of gene expression, but also as markers of transformed cells during stable transformation procedures. The ideal marker gene would be expected to display characteristics such as ease and cheapness of use, lack of toxicity, and robustness; and the most commonly used ones – GUS, GFP, LUC, and C1 + R/B (anthocyanin accumulation) exhibit most if not all of these properties. Each, however, differs in potentially important ways, and before deciding which to use it is important to consider carefully your particular set of experiments and the plant tissue you will be using. In this chapter, I will introduce each marker, outline protocols for their use, and discuss their strengths and weaknesses. Key words: Reporter genes, GUS, LUC, GFP, C1 + R/B, anthocyanin accumulation.

1. Introduction 1.1. UidA (GUS)

The potential for using the E. coli gene UidA, encoding the hydrolyase β-glucuronidase (EC.3.2.1.31) (GUS), as a scorable and quantitative marker in plants was first realized during the 1980s by R.A. Jefferson. The availability of a range of commercial β-glucuronides, which yield coloured or fluorescent products on hydrolysis, coupled to low endogenous levels of similar enzymatic activity in plants and their ease of handling ensured its rapid and widespread use in many plant species including cereals (1, 2). UidA has a monomeric molecular weight of approximately 68,200 Da and most probably functions as a tetramer. The enzyme, which requires no co-factors, has a pH optimum between 5.2 and 8.0 and is tolerant to a range of ionic conditions, and

Huw D. Jones and Peter R. Shewry (eds.), Methods in Molecular Biology, Transgenic Wheat, Barley and Oats, vol. 478 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-379-0_3

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detergents allow it to be easily assayed in crude extracts or whole tissues. Because of this tolerance and its stability, GUS is easily quantified, a major strength, but the need to bring the generally membrane-impermeable substrates in contact with the enzyme means that GUS is not usually measured or visualized in live, intact tissue. This is potentially less of a problem when used as a reporter for gene expression studies but has distinctly more limited appeal in a transformation marker. However, at least one protocol does suggest that non-destructive in situ detection can be achieved (3). There are two main methods used to detect GUS activity in transgenic material: either it is quantified in plant extracts using a fluorogenic substrate, or histochemical methods are employed on whole tissues. A number of fluorogenic substrates are commercially avaible but 4-methyl-umbelliferyl-β-D-glucuronide (MUG), which when cleaved by GUS yields the fluorescent product 4-methylumbelliferone (MU), has generally been the preferred substrate. For cellular localization of GUS gene expression, where absolute quantification of GUS activity is not required, the histochemical substrate 5-bromo-4-chloro-3-indolyl-β-D-glucuronide (X-gluc) is mostly used. UidA catalyses the cleavage of this substrate, generating the colourless and soluble product 5-bromo4-chloroindole. This, in turn, is readily oxidized and dimerized to form insoluble indigo, which is dark blue in colour. A great advantage of this substrate is that the indigo is easily visible by eye, needing no specialized equipment, although close observation of tissues requires at least a low-power light microscope. A second major advantage in using GUS/X-gluc is that the indigo product is stable through tissue fixation and embedding steps, which can make tissue-specific identification of promoter expression patterns relatively straightforward. 1.2. Luciferase (LUC)

The luciferase gene from the firefly Photinus pyralis (LUC) (EC.1.13.12.7) encodes a 62 kD protein. In a two-step reaction, it combines ATP with the substrate luciferin to yield an enzymebound intermediate complex, which then undergoes oxidative decarboxylation to produce the highly luminescent compound oxyluciferin (OL), which is capable of releasing a photon at 562 nm. Step 1 LH2+MgATP+LUC

LUC–LH2–AMP+MgPPi

Step 2 LUC–LH2–AMP+O2+OH–

LUC–OL+CO 2 +AMP+ light+H2O

Bioluminescence (meaning light produced by a chemical reaction that originates within an organism) has been found in several species and the enzymes involved are collectively known as luciferases; however, LUX from bacteria and RLUC from the

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coral Renilla are unrelated to firefly LUC and importantly require different substrates. The firefly enzyme has one of the highest known quantum efficiencies of all luciferases, but for use in plants and animals it was still found necessary for the wild-type gene to be modified to generate brighter forms (4, 5). LUC is detected in one of two ways. For direct enzyme measurements in cell extracts, either an illuminometer or scintillation counter is employed. The oxyluciferin product is released only slowly from the enzyme complex, so at high ATP concentrations the reaction gives rise to an initial (<10 s) flash of light emission that decays to a low, constant level as the enzyme slowly turns over. This can present a significant difficulty in measuring the light emission since it is essential that the delay between starting the reaction and measuring is constant and for a fixed time. Fortunately, several substances, such as Coenzyme A (CoA) or nucleotide analogues, have been found to change the initial flash of light production into a linear one that lasts for at least a minute (6), presumably as a result of increasing the release of the oxyluciferin and turnover of the LUC enzyme. Most LUC assays now make use of such compounds. It is also possible to make in planta measurements of LUC but these require the use of high-sensitivity charge-coupled device (CCD) imaging systems. At neutral pH luciferin carries a negative charge, and it was assumed therefore not to be membrane permeable, but in practice this appears not to be the case. Simply spraying plants with a solution of luciferin has proved to be sufficient in most cases (7). Levels of CoA are, however, low in plants and following an initial spraying of luciferin there will be a burst of light produced from the accumulated LUC similar to the reaction on extracts carried out in the absence of CoA. LUC protein is quite stable but it loses catalytic activity after only a few enzymatic cycles with the result that after repeated spraying with luciferin eventually light production becomes dependent on de novo LUC biosynthesis. At this point, the light produced reflects the steady-state production of the enzyme. This feature of the LUC reporter system has been used to great advantage in work studying real-time changes in gene expression, for example helping to understand the circadian rhythms of plants and animals (8, 9). Even with the use of strong constitutive promoters, the light produced by plants transformed with LUC constructs is very low, requiring complete darkness and very sensitive instrumentation for its detection. Possibly because of the specialist equipment needed for detection, LUC seems less favoured as a reporter in plants than GUS or GFP, except where dynamic changes in gene expression are being studied. It is often also the method of choice for transient expression analysis as part of a dual reporter system with GUS (10).

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In comparison with fluorescent proteins, light emitted from bioluminescence is very dim, therefore long exposure times are required which can limit temporal and spatial resolution. For fast events and sub-cellular imaging, a fluorescent protein may therefore be a more suitable reporter. However, LUC often has a lower signal-to-noise ratio since there are extremely low levels of background bioluminescence in plant material. Another advantage is that since bioluminescent reactions do not require exogenous illumination, bleaching of the reporter does not occur nor are tissues subjected to damaging short wavelength light. For this reason, the technique of bioluminescence resonance energy transfer (BRET) with LUC as the photon donor is gaining popularity in the study of protein interactions (11). In cereals, its use has mainly been limited either to transient expression studies, or as a marker for stably transformed cells and regenerated plants in barley and wheat transformation experiments (12, 13); in planta imaging has only been reported for rice (14). 1.3. GFP, Its Variants, and Other Fluorescence Proteins

Green fluorescent protein (GFP) is a stable 27 kD fluorescent protein from the jellyfish Aequorea victoria. Fluorescence is an optical phenomenon in which a molecule absorbs a photon of light to go to an excited electronic state, and upon returning to the ground state emits a second photon; since between these two events some energy is lost as molecular vibrations or heat, the emitted photon has a lower energy and therefore a longer wavelength. The aborption and emission spectra and the difference between them, known as the Stokes shift, is directly dependent on the structure of the chromophore. Wild-type GFP has two absorption maxima at 395 and 475 nm corresponding to the protonated and deprotonated states of the chromophore and corresponding emission maxima at 503 and 508 nm. Fluorescence emission of GFP does not require external substrates or cofactors other than O2, as the fluorescence derives from an internal p-hydroxybenzylidene-imidazo-lidinone chromophore generated by the cyclization and oxidation of a Ser-Tyr-Gly sequence at amino acid residues 65–67 in the protein. Detection of GFP in living cells therefore simply requires a source of blue light and a means of detecting green light, making GFP an ideal choice for in vivo monitoring of gene expression and protein interactions (15). Initial experiments using the native GFP were, however, disappointing because even when driven by strong constitutive promoters, little or no fluorescence was observed in heterologous hosts. Fortunately, this provoked a number of labs into generating modified versions of the wild-type protein to yield forms with greatly increased brightness and altered spectral properties and it is these new GFP variants that are routinely used as successful reporter genes in many organisms including plants (16, 17).

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Some of the induced amino acid mutations in GFP, Thr203 for example, gave rise to a variant whose absorption and emission maxima at 514 and 527 nm, respectively, were sufficiently shifted towards the red end of the spectrum for this protein to become known as a yellow fluorescent protein (YFP). Yet other mutations yielded blue (BFPs) and cyan (CFPs) forms of GFP. Initially, these different spectral fluorescence proteins (FPs) were generated at the expense of characteristics such as brightness, pH sensitivity, or stability; but improved forms of all the spectral classes of GFP are now available (16). A. victoria is in fact only one of a number of organisms to produce fluorescent proteins and now several genes from these species have also been isolated, such as DsRed from Discosoma sp. which has a red emission maxim (16). In order to detect any FP, it is necessary to invest in a suitable fluorescence microscope that is capable of imaging the distribution of a single molecular species based on the properties of its absorption and emission spectra. To do this, it will need a powerful light source, either a mercury or xenon lamp in wide-field or (epi-fluorescent) microscopes, or lasers in confocal microscopes. It will also require appropriate filters and a sensitive means of capturing the image against a dark background. An epi-fluorescence microscope may be all that is needed to view material satisfactorily, but generally for studying thicker samples a confocal microscope will become necessary. With the careful selection of variants and appropriate filters as well as software to deconvolute the fluorescence signals generated, it is possible to detect up to four different FPs in one plant cell. As FPs are tolerant to both N ′and C ′ terminal protein fusions (with and without linker amino acids), they are particularly useful in studying the subcellular location of proteins and protein–protein interactions. Increasingly, techniques such as fluorescence resonance energy transfer (FRET), fluorescence recovery after photobleaching (FRAP), fluorecence lifetime imaging (FLIM), and bimolecular fluorescent complementation (BiFC) are being successfully used in plants including cereals (18, 19). Genetically encoded indicators for Ca2+ coupled to FRET are not yet common in plants, but successful examples are to be found in the literature (19). FPs are also becoming the method of choice as a reporter during stable transformation experiments (particularly for cereals) as a means to improve transformation efficiencies and as part of the selection method (20, 21, 22, 23). Although FPs are not often quantified in extracts, it is possible to do this using a spectrofluorimeter and comparison to known quanties of fluorescence produced from the purified FP (24). 1.4. C1 + R/B: Anthocyanin Accumulation

Anthocyanins are water-soluble flavonoid pigments that can accumulate to high levels in the vacuole of the cell. They are often localized in the outer cell layers of many higher plant

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organs but most noticeably in flowers and fruits; hence their name ‘anthos’ meaning flower and ‘kyanos’ meaning blue (in fact, they range in colour from red to blue according to chemical structure, pH, and concentration). Anthocyanins are just one subgroup of the diverse family of aromatic flavonoid compounds derived from the phenylpropanoid pathway, other branches of which result in the production of the colourless chalones, flavones, flavonols, flavandiols, aurones, and isoflavonoids, as well as the brown pigmented oxidized proanthocyanidins (condensed tannins) and red phlobaphenes (25, 26). Within each subgroup, different plants and tissues can produce a range of compounds with differing substitutions around the basic aromatic ring structures. Anthocyanins provide visual cues for pollination and seed dispersal and have roles in defence and UV protection, such that natural accumulation occurs in response to temperature, water, and light stress. Most of the structural enzymes are found in the cytoplasm, possibly as part of multi-enzyme complexes, while the final products are transported into and stored in the vacuole. Their production is therefore also linked with cell morphogenesis and with the biogenesis and physiology of vacuoles (26, 27, 28, 29). With such a strong cell-autonomous visual detection, which requires no additional substrates, is not essential for development, and is non-toxic, anthocyanin accumulation has long been considered as a useful reporter system for both transient expression studies and stable transformations. Plants lacking anthocyanins through a genetic block in one of the structural genes of the pathway provide the easiest form of system when the missing gene is reintroduced by transformation (30), but the need to produce the necessary genetic stocks first precludes widespread use of this strategy. Instead, it was found that anthocyanin accumulation could be increased in wild-type plants by manipulation of the genes that regulate the pathway; for, despite numerous enzymic steps being involved in the production of anthocyanins, control of the pathway is dependent on a much smaller number of regulatory genes. In maize, members of two classes of transcription factors (bHLH and MYB) are primarily involved (29). For the production of anthocyanins to occur in a cell, there is a requirement for both MYB and bHLH factors to be present in the nucleus, as these appear to act as a heterodimer to promote transcription of the structural genes. Maize contains a small paralogous set of these factors with different members being involved in the control of anthocyanin accumulation in different parts of the plant. Hence, in the embryo and the aleurone layer of the endosperm, the MYB gene C1 is responsible for anthocyanin accumulation, whereas in the rest of the plant it is the closely related MYB gene PL (31). More duplication and

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specialization have taken place in the bHLH genes found at the R and B loci (32). It was established, however, that the paralogous factors were functionally redundant such that overexpression of one factor could substitute for the lack of another in a different tissue. Furthermore, the maize regulatory proteins could activate anthocycanin production in heterologous hosts such as arabidopsis and tobacco (33, 34, 35), suggesting that the essential components of the flavonoid pathway and its regulation are conserved between species. Even though further work has implicated the involvement of other different transcription factors in the expression of the complete phenylpropanoid pathway, to effect anthocyanin accumulation it is generally a matter of overexpressing representatives of the MYB and bHLH factors (36). In some instances it is sufficient to overexpress either the MYB or the bHLH factor; in others both are required. This presumably reflects either the level of endogenous MYB or bHLH factors in any cell or the inability of certain combinations of host and introduced factors to interact correctly. However, while not all cell types respond to MYB/bHLH overexpression with accumulation of anthocyanins, others may do so only at particular stages of their development. In cereals, for example, anthocycanin naturally accumulates in the outer layer of the endosperm (the aleurone), but overexpression of C1 + R-S or C1 + R-Lc in the developing grain of rice or wheat fails to result in accumulation of anthocyanins in the inner starchy endosperm cells ((37), Huttly unpublished), despite the use of highly active constitutive promoters. In rice, other noncoloured flavonoids were detected in these cells and increased expression of at least some of the structural genes of the phenylpropanoid pathway was detected, thereby suggesting that only a part of the pathway had been activated (37). There may be several reasons for a lack of apparent anthocyanin accumulation, including the flow of metabolites through other parts of the phenylpropanoid pathway, and the state of cellular and/or vacuolar differentiation. This presents a serious limitation to the use of anthocycanin accumulation as a reporter system when wishing to study tissue-specific promoter analysis, since lack of coloured pigmentation in a particular cell type may not be due to absence of promoter activity. However, anthocyanin accumulation has been used as a visual transformation marker, helping to improve cereal transformation protocols and track transgenic plants (38, 39, 40): the commercial unacceptability of dark red transgenic plants though seems to have limited widespread use to research orientated projects. Where anthocyanin accumulation has proved most useful is in transient expression analyses, since it is possible to score individual positive cells easily

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by counting (41). It can be used as either the reference or the report system with any of the other reporter genes described above: GUS, LUC or GFP.

2. Materials 2.1. Constructs

1. UidA: Numerous vectors containing the UidA gene are available either commercially or on request from users’ laboratories (see Note 1). More recently, vectors containing glucuronidase genes isolated from Staphylococcus and Penicillium species, under the names GUSPlus and penGUS, respectively, have become available and are reported to be up to 10 times more active in plants (42). 2. LUC: The LUC + gene available from Promega (4) is reported to be 50 times more active in light production than the native form when tested in wheat and maize (5) because of the removal of cryptic splicing, glycosylation and poly A addition sites, and improved codon usage. The C′ terminal sequence directing expression of the protein to the peroxisome is also modified in this version of the enzyme to prevent targeting to this organelle. LUC + has proved amenable to both N′ and C′ protein fusions (4, 11). 3. FPs: Numerous FP gene constructs are available from Clontech and from the labs in which the variants have been generated (16, 17). This includes FPs targeted to sub-cellular locations such as the nucleus, endoplasmic reticum, and plasmalemma (18, 19). FPs tested in cereals have mainly been restricted to sGFP S65T in barley and wheat (20, 21), or mGFP5-er in rice (43). Both these variants contain modifications to improve codon usage, removal of a cryptic intron for optimal expression, as well as amino acid changes that have increased maturation speed, quantum yield, and photostability, all of which increase brightness over the wild-type enzyme (16, 17). The FPs EYFP, mcitrineYFP, ECFP, and mRFP have all also been used successfully in plants (18, 19), and there is one report of five coral reef FPs from Anemonia, Discosoma, and Zoanthus (44) in a range of plants including cereals (see Note 2). The choice of which FPs to use depends largely on the type of experiment and particular plant or tissues to be studied. Reviews detailing the physical properties of currently available FPs in addition to their absorption

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and emission spectra are useful sources of information (16, 18) (see Note 3). 4. Anthocyanin regulatory genes: The most common regulatory genes to be used in manipulating anthocyanin production in transgenic plants have been the maize MYB C1 gene and either the maize R-Lc or B-peru bHLH genes with a range of different promoters being used to drive their expression (35, 38, 45, 46). Clones of these genes can be obtained from the laboratories reporting their cloning or use. To use anthocyanin accumulation as a marker in a new system, it will be necessary to overexpress both types of transcription factor by default or by testing whether one will suffice.

2.2. Media

1. Fluorogenic Gus assay using MUG 1. GUS Buffer: 50 mM sodium phosphate buffer pH 7.0; 10 mM EDTA.Na2; 0.1% Triton-X-100; 0.1% sarcosyl; 10 mM DTT (see Note 4) 2. 3M: 0.4 M mannitol,15 mM MgCl2, 0.1% MES pH 5.4 3. 0.2 M Na2CO3 (anhydrous). 2. Histochemical GUS assay 1. HGUS buffer: 100 mM sodium phosphate buffer pH 7.0; 10 mM EDTA; 0.1% Triton X-100. Add prior to use: 1. 1 mM (0.5 mg/ml) X-gluc sodium trihydrate (MW 498.7) (stock 10 mg/ml in water kept at −20°C in the dark) 2. 1 mM potassium ferricyanide, 1 mM potassium ferrocyanide (see Note 5) 3. 10 µg/ml chloroamphenicol (stock 10 mg/ml EtOH), 4 µg/ml nystatin (stock 4 mg/ml EtOH) (see Note 6) 4. 10–100 mM ascorbate (see Note 7). 3. LUC assay 1. Luciferase lysis buffer: 100 mM potassium phosphate buffer pH 7.8; 1 mM EDTA; 10% glycerol; 1% Triton X-100; 7 mM 2-mercaptoethanol 2. 3 M: 0.4 M mannitol, 20 mM MgCl2, 0.1% MES pH 5.4 3. Luciferase assay buffer: 20 mM Tricine pH 7.8; 5 mM MgCl2; 0.1 mM EDTA; 3.3 mM DTT; 270 µM coenzyme A; 500 µM luciferin; 500 µM ATP (store in aliquots at −70°C) (see Notes 8 and 9).

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3. Methods 1. b-Gluronidase Detection Using 4-Methyl-Umbelliferyl b-D-Glucuronide

GUS cleaves MUG to yield the fluorescent product MU but fluorescence of MU is maximal only when its hydroxyl group is ionized, and therefore it is necessary to bring the pH of the assay above 8, which has the advantage of simultaneously stopping the reaction. The MU product can then be detected in a fluorimeter and quantified relative to known MU standards or to a standard curve produced using purified GUS protein. A number of alternative fluorogenic substrates are commercially available, but not commonly used, although resorufin glucuronide with an excitation at 570 nm and emission at 590 nm and maximal fluorescence at neutral pH could be a useful alternative to MUG. Methods for using alternative substrates can be found in (47). 1. Tissue preparation: whole tissues 1 Remove a small (5–50 mg) sample of tissue leaf, root, callus, etc. The amount required will need to be determined for your material (see Note 10). 2. Homogenize tissues in (50–200 µl) GUS buffer in Eppendorf tubes. 3. Pellet cell debris by spinning 15 min at 16,000 × g, 4°C. 4. Determine the protein concentration of the extract (see Note 11). 5. Dilute the tissue extract with GUS buffer to a concentration of 0.1 mg total protein/ml. 6. Use extracts immediately or store at −80°C (see Note 12). 2. Protoplasts 1. Collect transformed protoplasts in (1 × 106) aliquots by centrifugation for 1 min at 50 × g and re-suspend in 2 ml of 3 M buffer (seeNote 13). 2. Remove an aliquot of protoplasts to count with a haemocytometer. 3. Centrifuge the remainder for 1 min at 50 × g and re-suspend in 150 µl of GUS buffer. Vortex briefly (or pass through a 26G needle) and leave to stand for 5 min. 4. Pelletize cell debris by spinning for 15 min at 16,000 × g, 4°C. Determine the protein content of the extract and dilute as for whole tissue samples (step 5).

3.2. Flurogenic MUG Assay

1.Place 50 and 25 µl samples of tissue extract in wells of an opaque 96-well microtiterplate (black or white). Include

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untransformed tissue controls. Bring to a total volume of 200 µl with GUS buffer and mix. 2. To start the reaction, add 100 µl of GUS buffer containing 2.5 mg/ml MUG (final concentration 2 mM), mix, and cover. 3. Place assays in an incubator at 37°C and allow them to equilibrate for 5–10 min. Remove a 50 µl aliquot and further aliquots after suitable time intervals (10, 20, 30 min, or hourly) into 200 µl aliquots of 0.2 M Na2CO3 in a second microtiterplate to terminate the reaction. 4. Measure the fluorescence from the MU product using a platereading fluorimeter using excitation at 365 nm, and emission at 455 nm. Check that the samples produce linear kinetics and that dilutions of protein extracts and duplicate samples correspond (see Notes 14 and 15). 5. GUS activity can be expressed as pmoles MU/min/mg protein, in which case standards of MU in the linear range of the fluorimeter (e.g. 0–5 µM) should be diluted into 0.2 M Na2CO3 and included in the plate. Solutions of MU, 1 mM, may be stored at +4°C wrapped in foil, and check for precipitation of the Na2CO2 before use. Alternatively, activity can be expressed as fluorescence units (flu)/ng protein/min. To convert this to the amount of UiDA protein, it will be necessary to generate a standard curve using purified GUS enzyme (type VII-A from E. coli, Sigma). Dilute the GUS enzyme in GUS buffer to give 0.1 ng/µl working strength. Add between 0 and 2 ng of enzyme in total volumes of 50 µl GUS buffer and assay as above. 6. Some plant tissues have high levels of endogenous glucuronidase activity. It is still possible to measure GUS activity in these tissues using modified reaction conditions (see Note 16) 3.3. Histochemical GUS Assay

There are many, slight variations to the basic histochemical protocol for GUS detection using 5-bromo-4-chloro-3-indolyl-βD-glucuronide (X-gluc) as substrate in the literature (47). The differences mainly revolve around the methods to deal with two potential problems of the X-gluc reaction, namely the impermeability of the glucuronide substrate, making tissue penetration an issue, and the solubility of the colourless intermediate 5-bromo4-chlorindole that can diffuse away from the site of GUS activity. Both affect the accuracy with which the indigo product reflects the location of the enzyme in tissues and cells. With any new transgenic material, it will be necessary to observe the effects of inclusion of a pre-fixation step and addition of the ferri and ferrocyanide salts, which catalyse oxidation of the 5-bromo-4chlorindole product. Untransformed control tissue should also be studied in order to gauge the level of endogenous glucuronidase

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activity present; this is important because although generally very low, some tissues, particularly pollen but also developing seeds and fruits, can exhibit significant levels (48). A number of papers have outlined methods to reduce or remove this endogenous activity if necessary ((49), see Note 16). Because of the problems associated with X-gluc, the use of alternative substrates has been explored. Cleavage of the β-glucuronide from Sudan-β-D-glucuronides results in the liberation of an insoluble Sudan dye, so no diffusion artefacts are generated, and in tests the latest Sudan IV variant indicated it may be a valuable histochemical glucuronide substrate (50). Similarly, a fluorogenic β-glucuronidase substrate has also been developed that is relatively membrane permeable. ImaGene Green C12FDGLcU gives rise to a 5-dodecanoylaminofluorescein (Abs 495 nm, Em 518) that is retained by cells. A few papers have shown the use of this substrate but it has not been widely employed (2, 51). Figure 1a shows the results of using the basic histochemical protocol on Arabidopsis leaf material stably transformed with a promoter–GUS construct. The material was not prefixed, but 1 mM ferri and ferrocyanide were included in the buffer. The image was captured using a Leica DFC300FX digital camera attached to a Leica MZ8 stereomicroscope. 1. Place tissue (see Note 17) in sufficient freshly made HGUS buffer (containing X-gluc and other additions) to submerge the material completely. The assay can be performed in the wells of a microtiter plate or in tubes.

Fig. 1. (a) Histochemical detection of GUS activity directed by the promoter from a highly expressed Brassica napus gene in a stably transformed Arabidopsis leaf. (b) In planta detection of the same promoter driving the LUC gene in the upper part of the Arabidopsis stem, cauline leaf and developing floral buds. (c) The rice actin promoter driving sGFP S65T in a germinating wheat seed. (d) Transient anthocycanin accumulation in developing wheat aleurone cells bombarded with a wheat 8S globulin promoter driving the R-Lc and C1 genes from maize.

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Fig. 1. (continued)

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2. Apply vacuum for 5–10 min and release (see Note 18). 3. Wrap in an aluminium foil and place at 37°C for up to 48 h (see Note 19). 4. Wash material in 100 mM phosphate buffer, pH 7.0. 5. To clear material for observation, incubate for 15–30 min in 25% EtOH (see Note 20), 15–30 min in 50% EtOH, and 30 min in 75% EtOH. 6. Finally, incubate in several changes of 90% EtOH at 37°C 2–4 h or until all chlorophyll has been removed (see Note 21). 7. Samples can be kept indefinitely in 90% EtOH in tightly capped tubes in the dark at room temperature, but the level of EtOH should be monitored regularly. 3.4. Detection of LUC Activity in Plant Extracts)

The reaction catalysed by LUC is dependent on several different parameters including temperature, pH, concentration of components, and metal ions. For this reason, the standard protocols use a known volume of Tricine buffer, pH 7.8, including Mg2+ ions, DTT, and EDTA, and are carried out at room temperature. 1. Whole tissues 1. Freeze plant tissue (5–50 mg) in liquid N2 and grind to fine powder. 2. Re-suspend at room temperature in 100–400 µl lysis buffer with further homogenization. 3. Remove the debris by centrifugation for 15 min at 16,000 × g, 4°C. 4. Determine protein concentration of extracts (seeNote 22). 5. Use extracts immediately or store at −80°C. 2. Protoplasts 1. Harvest (1 × 106 protoplasts) by centrifugation at 50 × g for 4 min. 2. Re-suspend in 2 ml of 3 M buffer, and spin as above. 3. Re-suspend in 400 µl lysis buffer, and vortex briefly (or pass through a 26G needle). 4. Pelletize the cell debris by centrifugation for 15 min, at 4°C and 16,000 × g. Determine the protein concentration. 3. Luciferase assay There are many different luminometers available. The most sophisticated, and therefore the most expensive, ones are based on a 96-well format and are fully automated. Below is the protocol for using a plate-reading luminometer (see Note 23). Different instruments vary in sensitivity and signal-tonoise ratios but can detect down to 1 fg of luciferase. For each instrument, it will be important to determine the linear range

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of light detection and to produce a standard curve. It is also possible to measure bioluminescence using a liquid scintillation counter (see Note 24). 1. Place 20 µl aliquots of plant extract into wells of an opaque microtiterplate. Include the untransformed tissue extract as control. 2. Programme the luminometer to dispense 100 µl of the Luciferase assay buffer (see Note 25) and to perform a 2 s measurement delay followed by a 10 s measurement read for luciferase activity. The light intensity of the reaction is nearly constant for about 1 min and then decays slowly, with a half-life of approximately 10 min. The read time may be shortened if sufficient light is produced. 3. Express activity as fluorescence units (flu)/10 s/mg or relate to purified LUC obtained from a standard curve (see Note 26). 4. Transient assays are often performed with a second (internal control) reporter construct such as GUS and results normalized in comparison to the control reporter expression. It should be noted that GUS is reported to be fully active when plant tissue is extracted in luciferase lysis buffer, whereas GUS buffer adversely affects LUC. Secondly, it should be demonstrated that the control neither interferes with the expression of the test construct nor is affected by the culture conditions. Typically, expression from the control plasmids is kept low by mixing in the ratio 1:10 with the test construct prior to bombardment or electroporation. It is also to be remembered that transient experiments will require multiple repartitions to provide statistical relevance. 3.5. In Planta Measurement of LUC Using a Cooled CCD Camera

For in planta measurement of luciferase, penetration of the luciferin into live tissues is an issue that needs to be addressed. Simply spraying plants with a solution of luciferin has proved to be sufficient in most cases (7). However, some tissues, such as seeds in the late stages of development, do not readily take up luciferin, and require wounding (see Note 27) to help substrate penetration (52, 53). Apparent lack of LUC activity in planta should ideally be verified through alternative means using tissue extracts or in situ hybridisation (7, 54). Since wounding will affect the luciferin–O2 equilibrium reached even after repeat-spraying of luciferin, experiments designed to look at wounding-affected gene expression may be difficult to interpret using LUC. Although O2 and Mg2+ are generally not limiting for luciferase activity, under specific conditions such as submergence of plant tissue, O2 levels can become so with a consequential drop in light production (7). Similarly, cell suspension cultures grown at

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different levels of oxygenation have been shown to have differing levels of LUC activity detectable. 1. Preparation of material 1. Using a fine aerosol spray, a 1–5 mM solution of D-luciferin in water is applied directly onto whole plants, wetting them completely (see Note 27). Aliquots of 100 mM D-lucifierin in water should be kept at −20°C and diluted freshly before use. 2. Repeat the above spraying with luciferin at 24, 16, 4, and 2 h before imaging (see Note 28). 2. Detection of LUC in planta The easiest method of capturing the low levels of light emitted is by X-ray or photographic film. Whole plants expressing a luciferase construct have been imaged by pressing against X-ray films to determine the location of expression, although resolution is not high in such images. Hence, for most experiments it will be necessary to purchase a sensitive CCD camera system. There are two types of CCD devices that are generally used: the slow-scan liquid nitrogen-cooled CCD camera, and the intensified CCD camera. The cooled camera types generally offer higher spatial resolution (9) but they have a higher sensitivity to wavelengths longer than 500 nm (see Note 29). In either camera system, a silicon wafer is employed that emits electrons when struck by photons. Electrons emitted from a single pixel are quantified electronically and an image created. The number of electrons emitted is expressed in grey scale units (gsu). Software such as MetaMorph can translate these gsu values into a pseudo-colour image within set boundaries. Quantification of light levels within defined areas can then be made, but in practice in whole plant images this can be difficult as the orientation of leaves and other organs can affect the signal detected. Figure 1b shows the upper floral bolt showing the stem, cauline leaf, and developing floral buds from an Arabidopsis plant transformed with a promoter–LUC construct. The image was captured with a 3 min exposure using a Princeton Instruments slow-scan liquid nitrogen-cooled CCD camera with a Nikon 50 mm lens and shows a pseudo-colour image in which blue is set for low intensity pixels and red for high intensity produced using MetaMorph software. 1. Set up the camera by cooling to −120°C working temperature with liquid nitrogen (see Note 30). 2. Approximately 10–20 min before capturing data, spray the material with luciferin. 3. Place plants, or isolated leaves, stems, roots on the camera stage with the back lamp on, focus the camera, and capture an image as reference.

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4. Turn the light off and leave material in complete darkness for 5–10 min (see Note 31). 5. Capture data for between 1 and 30 min (see Note 32). 6. Calculate the image intensity using an imaging software (see Note 33). 3.6. Microscopic Detection of Fluorescent Proteins

Material expressing FPs is generally viewed initially with an epifluorescence microscope. In some cases this will be all that is needed, particularly if events in the epidermis are being studied or if deconvolution software is available to remove the out-offocus fluorescence collected (18). However, for studying thicker samples, confocal microscopy then often becomes necessary since it has the ability to produce thin (0.5–1.5 µm) optical sections through fluorescent specimens up to a thickness of 50 µm or more. It is also useful in situations where multiple FPs are being imaged, since multiple emission wavelengths can be detected concurrently (see Note 34). A major disadvantage a confocal microscope is in its use of lasers to illuminate specimens. As there is only a limited range of laser excitation wavelengths commonly available, it will often be necessary to compromise on excitation efficiency: depending on the chromophore’s physical characteristics, a 25% excitation efficiency may be adequate but 80% or greater would be regarded as optimal. Selection of the correct filter sets (see Note 35) is highly important, whatever microscope is in use since suboptimal filters can lead to reduced apparent perceived brightness of the FP. This could in turn result in the need for increased exposure times and illumination intensities, which not only could affect photostability of the FP, since all FPs will photobleach in time, but also adversely affect cell viability. A number of different companies sell filter sets that are accommodated by a range of microscopes. Since they should be selected to best suit the particular variant of FP being used, it can be expected that several different sets will be required (16, 55). A significant problem associated with the use of FPs as reporters is that a range of metabolites and structural components of cells can exhibit natural fluorescence: generally referred to as auto-fluorescence. Auto-fluorescence spectra unfortunately can be broad, often encompassing most of the visible range, and so overlapping inevitably with emission wavelengths of GFPs and other FPs. In plant samples, the most important sources of autofluorescence are chlorophyll, polyphenols, lignins, cellulose, and vacuolar contents, but culture media and some fixation methods can often be unwanted sources of intense auto-fluorescence. Careful selection of a particular FP variant and filter sets can alleviate many of the problems encountered in individual tissues (see Note 36). It will also be essential to compare transgenic to

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non-transgenic material particularly when expression is low and the researcher is inexperienced. To enhance detection of FPs, many groups have used constructs that target the FP to subcellular locations, the benefit being a concentration of the signal into a distinctly recognizable pattern as opposed to a low-level diffuse fluorescence, which may not be greatly different in spectral characteristics to the auto-fluorescence of the sample (see Note 37). A second method of enhancement is to use genetic amplification. FPs differ from other reporter systems in that there is no built in enzymatic amplification step in its detection, which can mean that it is a less sensitive detection method than GUS or LUC. Genetic amplification in which the promoter of interest is fused to a trans-activator, for example gal4:vP16, and the FP under control of cis-elements to which the trans-activator will bind is one way of producing an amplification step (43). Figure 1c shows a germinating wheat seed of a plant stably transformed with a rice actin promoter–GFP construct. The GFP used was the sGFP S65T variant, and the image was captured using a Leica DFC300FX digital camera attached to a Leica epifluoresecence MZFl III stereomicroscope using a Leica GFP2 filter set (see Note 42). 1. View material initially using an epi-fluorescence microscope. 2. Switch on the mercury lamp (see Note 38), first checking that the light stop is in place. 3. Place live tissues (whole mount or sectioned material) on the microscope stage (see Notes 39–42). 4. Focus the microscope using transmitted light. 5. Select filter set (see Note 43), remove light block from the mercury lamp, and view sample (see Note 44). 6. Capture images as required (see Note 45). 7. If necessary, view material using a confocal microscope (see Note 46). 3.7. Quantification of Anthocyanin Expression

1. Transient expression studies Following bombardment of tissues, transformed cells will begin to appear stained dark red or occasionally dark blue after about 12 h of incubation. The colour often appears spread throughout the vacuole, giving the appearance that the whole cell is red. The colour will intensify for several days and can remain stable for 1–2 weeks. For some cells, the anthocyanin pigment is sequestered in densely coloured vacuolar inclusions or AVIs (27), or it can slowly become so localized over time, and in these cases individual cells are more difficult to score. Although, visible by eye if sufficient cells are accumu-

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lating anthocycanin, scoring of the number of transformed cells by counting will require a low-power stereomicroscope. Figure 1d shows developing aleurone cells from the wheat cultivar Cadenza transiently expressing the maize genes C1 and RLc, both driven by a wheat 8S globulin promoter. Anthocyanin accumulation was visible only when both constructs were bombarded together in equimolar quantities and the image taken after 3 days incubation. Anthocyanin accumulation was visible only when both constructs were bombarded together. The image was captured using a Leica DFC300FX digital camera attached to a Leica MZ8 stereomicroscope. 2. Quantification of anthocyanins in plant extracts by absorption spectra It is possible to quantify anthocyanins by a number of methods including extraction and absorption at 535 nm (56) or by image analysis (57). Quantification and identification of individual anthocyanins can be achieved by high-performance liquid chromatography (HPLC) and liquid chromatographymass spectrometry (LC-MS) using the same extracts once concentrated by evaporation (58). 1. Homogenize tissue, 200 mg in 3 ml of acidified ethanol or methanol (ethanol:HCl 0.1N 85:15 v/v, pH 1.0). 2. Adjust pH to 1.0 with 4N HCl and shake vigorously for 30 min, re-adjust pH to 1.0 if necessary. 3. Repeat shaking for 30 min, then centrifuge at 27,000 × g for 15 min, 5°C 4. Leave extract at 5°C for 48 h, centrifuge as in step 3, and pass through a 0.45 µm filter. 5. Make up to 3 ml with acidified ethanol. 6. Measure total anthocycanin content by absorption at 535 nm against a reagent blank. Quantify the amount of anthocyanin present in the sample against a standard curve prepared using a suitable commercially available standard anthocyanin such as cyanine-3-O-glucoside. 3. Quantification of anthocyanin by image analysis Quantification of anthocycanin can be achieved by measuring the pigment intensity using image analysis. 1. Place 1 µl drops of a known series of cyanidin-3-O-glucoside (0–400 µg/ml) concentrations, prepared in acidified ethanol, on slides. Capture images with a digital camera. Colour intensity is quantified with suitable software and the correlation coefficient calculated between pigment intensity and cyanidin-3-O-glucoside concentration. 2. Image the samples under the same conditions as above.

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3.8. Final Words

When using any reporter to monitor gene or protein expression, it is important to remember that you are creating an artificial situation. Any promoter that has been isolated, cloned into a plasmid, and reintroduced into the genome is now in a genomic context different to the endogenous gene. This is still true even if all of the endogenous cis-elements required for its expression, which could reside in the 5′ upstream or 3′ downstream regions to the gene, within its coding region including the introns, or even within other nearby genes, are present within your cloned sequences. Such isolated promoters can be influenced by cryptic cis-elements present in the plasmid sequences and the new genome position, and by other strong promoters within the constructs, such as the CaMV35s or maize ubiquinin (Ubi) promoters that are often used to drive the transformation selection marker. Reporter gene expression patterns should therefore be supported by other data, such as in situ hybridization, and in fact many journals will not report data based solely on reporter gene promoter constructs. Another problem relates to the level of expression of the reporter gene. Several papers have suggested that there is an inverse relationship between high levels of reporter gene expression and regeneration in transformation procedures. It is not clear whether this points to toxicity of the reporter or suppression of the cell’s potential to regenerate, or simply a reflection of the fact that cells capable of high level gene expression are in a different development phase. The net effect is that selection based solely on reporter gene expression has given rise to lower transformation efficiencies (21, 22, 40, 59). High levels of reporter gene expression can also result in gene silencing in some individuals. Silencing can be tissue- or cell specific, often leading to a range of patterns of reporter gene expression in siblings over and above differences that might relate to detection thresholds, and it can become more pronounced in subsequence generations. For this reason it is prudent to study between 10 and 20 independent transformants carrying a particular gene construct. High levels of expression may also lead to silencing or ectopic expression of GFP within cells or to the aggregation of chimeric proteins still able to fluoresce; here work needs to be verified by immunocytochemistry techniques, and working with lines showing a range of expression levels may also be informative. It should also be noted that different cells can have vastly different internal structure and volume; therefore the same level of gene expression in one type of cell compared to a second can give rise to apparently different levels of reporter or concentration of reporter within a cell. Good background knowledge of plant physiology should prevent overinterpretation of data in this case.

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4. Notes 1. Over time, several modifications to the native E. coli gene have been made, including enhancement of the translation initiation site and removal of glycosylation sites to allow processing through the secretary system (1, 60). For use with Agrobacterium delivery systems, it is best to employ one of the many UidA versions into which an intron has been introduced, as this will prevent low-level mis-expression of the gene in contaminating bacteria following co-cultivation. The UidA protein is tolerant of N′ and C′ terminal fusions and some vectors are designed to facilitate this (42, 61). 2. The excitation spectra of all wild-type FPs change with protein concentration, suggesting that some form of interaction is occurring, but while GFPs are categorized as weakly dimeric, FPs from some of the other organisms appear to be strictly oligometric, which initially seemed to cause significant problems in their use. Fortunately, further mutation screens have meant that monomeric forms are now generally available for these proteins (16). 3. Knowledge of the extinction coefficient (amount of light absorbed), quantum yield (% of light emitted as fluorescence), and relative photobleaching (photodestruction of the chromophore in the presence of O2) rate of the FP can be very important. For example, FPs with low extinction coefficients and quantum yields but with high photobleaching will require higher light intensities for detection and lead to high free-radical generation, both of which can damage cells and cause significant changes in cell development over time. Similarly, shorter wavelengths are of higher energy and therefore more damaging, limiting the use of BFPs. For techniques such as FRET and BiFC the emission spectrum of the donor must overlap significantly with the absorption spectra of the acceptor while overlaps between the two aborption and emission spectra should be minimized, and for this reason YFPs and CFPs have often been used as partners. 4. GUS is generally very tolerant to the reaction conditions but it is most active in the presence of thiol reducing agents (β-mercaptoethanol or DTT), and since it may be inhibited by some divalent heavy metal ions (Cu2+, Zn2+), EDTA is generally included in assay buffers (1). 5. Inclusion of ferri and ferrocyanide salts in the incubation accelerates formation of the insoluble indigo product, thus minimizing diffusion of the 5-bromo-4-chloroindole away

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from the site of enzyme activity and providing a more precise and intense stain. However, it also inhibits the glucuronidase; hence, a compromise needs to be sought and an optimal concentration, typically between 0.5 and 10 mM, determined empirically. At low levels of GUS expression, the ferri and ferrocyanide salts can reduce the enzyme activity to a level that is undetectable. With any new material, it is best to include an assay omitting the salts altogether for comparison. 6. General antibiotics can be added to the assay to prevent bacterial and/or fungal growth during long assay incubations, as these organisms have very active β-glucuronidases. 7. If the tissue is prone to oxidative browning, addition of 10– 100 mM ascorbate to the HGUS buffer can alleviate this. 8. When using LUC assays to measure the amount of enzyme in a sample, as opposed to measuring ATP concentrations, maximum sensitivity is required and hence high levels of ATP are required. Under these conditions, the reaction gives the initial (<10 s) flash of light emission that decays rapidly to a low, constant level. Under these conditions, the reaction needs to be initiated while within the counting chamber of an illuminometer and mixing of the initializing component to be rapid and complete. To produce a reaction where light is emitted linearly for at least a minute, the inclusion of high levels of CoA are required (6). 9. It is possible to buy commercial LUC assay kits, and the buffers here are essentially very similar to these. 10. It is possible to freeze material in liquid nitrogen and store at −80°C. 11. Use a suitable proprietary protein assay. 12. It is generally better to process extracts immediately, as storage of some frozen samples at −80°C can lead to random changes in GUS activity. 13. It may be necessary to first wash protoplasts free of culture media components which can interfere with the assay. Remember to include an untransformed aliquot. 14. The assay is generally found to be very stable and few problems are encountered, allowing in some cases very low levels of GUS to be detected. However, such circumstances increase the likelihood of non-enzymic cleavage of the substrate, or allow low levels of endogenous glucuronidase activity to be detected. Similarly, in some tissues certain compounds can mask the fluorescence through either intrinsic fluorescence or absorption or inhibit the reaction. To test these possibilities, appropriate untransformed controls should be included and a reconstruction assay should

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be carried out. To do this, add the equivalents of 0, 1, 2, and 5 µg protein of an untransformed control plant extract to 0.2–1.0 ng of purified GUS enzyme. Assay as above and plot the fluorescence against GUS enzyme with or without plant extract. If the plant extract significantly effects the activity of the purified enzyme, reducing the amount of extract used or the reaction times may resolve the problem, or partial purification of the extract may be necessary (see Note 15) if high endogenous levels of glucuronidase are present in the tissue (see Note 16). 15. Some protoplasts and tissues produce phenolic compounds which can interfere with the fluorescence of MU (62). In this case, add 5–10 mg of acid-washed Polyclar AT to the GUS buffer used for tissue extraction. Prepare a 1.0 ml spin column of Sephadex G25 equilibrated in GUS buffer in a 1.5 ml Eppendorf tube as follows: puncture the base of the Eppendorf tube with a syringe needle and layer 20–30 µl of glass ballotini beads (0.1–0.2 mm diameter) at the bottom of the tube. Fill the tube with a suspension of Sephadex G25 equilibrated in GUS buffer over the beads and place in a small centrifuge tube such that it is suspended at the top of the tube. Centrifuge for 4 min at 450 × g and exchange the centrifuge tube for a clean one, and the column is then ready for use. Layer the protoplast extract on the top of the Sephadex bed and spin through the column by centrifuging for 4 min at 450 × g. Add a further 50 µl of GUS buffer through the column and spin for 4 min at 450 × g. It may also be necessary to increase the concentration of the NaCO3 required to stop the MUG assay from 0.2 to 1 M. 16. An initial report suggested that addition of methanol to the reaction buffer (up to 20% (v/v)) was effective for inhibition of endogenous GUS activity, but for some tissues this has proved insufficient. Detailed comparisons between the endogenous and UidA activity have since determined that they can usually be distinguished by their differential sensitivity to the inhibitor, saccharic acid-1,4-lactone, by their thermal stability and by different pH optima. Typically, modifying assay conditions by raising the pH to 7.5 as well as pre-incubating the tissue at 55°C for 1 h prior to conducting the assay at the same elevated temperature appears to remove any endogenous activity while allowing the UidA activity to be determined (49). Since the level of endogenous glucuronidase activity can be genotype dependant, it is important to stress that appropriate non-transformed control material is included in all experiments, particularly if UidA GUS levels are low, and appropriate changes to the basic protocol made if necessary.

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17. Materials can range from whole seedlings, leaves, or roots through to 205 mm hand sections cut with a sharp double-edged razor blade 2–5 mm3. Alternatively, 200–300 µm sections cut with a vibratome can be used. Unless otherwise targeted, UidA accumulates in the cytoplasm, and therefore staining of very small, unfixed samples can lead to poor localization. Cut sections should be used if the material is to be embedded and sectioned. 18. Penetration of the substrate can be aided by a light pre-fixation step. Tissues are immersed in 0.1–1.0% glutaraldehyde, 25 mM sodium phosphate pH 7.0, vacuum infiltrated, and incubated for 30 min on ice. Samples should then be washed three times for 5 min in cold water or phosphate buffer. Alternative pre-fixation steps include immersion in ice-cold 90% acetone and incubation for 20 min on ice, or incubation in buffered 2% paraformaldehyde, 100 mM Na-phosphate pH 7.1 mM EDTA for 30 min on ice. Note: glutaraldehyde and paraformaldehyde solutions should be freshly prepared and never used if milky or discoloured and disposed of in hazardous-waste containers. For any new material, an unfixed sample should also be included to gauge the effect of fixation on glucuronidase activity and staining. 19. Colour formation should be monitored during this time, and the reaction stopped once sufficiently dark. It should be noted, however, that if GUS activity is low, it could be obscured by the chlorophyll present in most samples. 20. This will require several changes of EtOH. To speed up the clearing, we often incubate in freshly made FAA: 5% formaldehyde, 5% acetic acid, 40% EtOH for 10 min after the 25% ETOH incubation, even if the intention is to view material without embedding and sectioning. Material such as seeds can be particularly difficult to clear for these refer to (63). 21. Fixation, embedding, and sectioning after staining can be performed with any common procedure most appropriate to the material being stained, including FAA or gluteraldehyde. For general plant microscopy methods, refer to (64). Once embedded, sections are best cut thick, at least 2–3 µm, to allow sufficient blue colour to be visible and viewed under interference contrast optics with polarized light, to enhance unstained cellular structure. 22. Measuring protein concentrations can be difficult in this lysis buffer because of the high levels of detergents (>0.1%) and mercaptoethanol. Selection of a proprietary protein assay that is tolerant of these will be necessary (65), or if the protein extract is sufficiently concentrated, then a suitable dilution should allow analysis by most methods.

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23. For manual luminometers the luciferase assay buffer is dispensed into tubes and the reaction initiated by addition of lysate; for machines with injectors, samples are placed in the tubes or assay plate and the reaction is started by addition of the assay buffer. 24. To use a scintillation counter, the coincidence circuit of the counter should be turned off but if this not possible a linear relationship between bioluminescence and counts detected can be generated by calculating the square root of the measured counts per minute (cpm) minus background cpm. Samples are placed either directly into the scintillation vial or within smaller tubes, although these need to all be placed in the same orientation as each other within the scintillation vial. For consistency, it is best to operate in manual mode mixing and reading samples individually with all channels open and collecting data from 10 s to 5 min after a suitable delay (10–20 s) to reduce the background. With scintillation counters, again it is important to determine the linear range of your instrument. 25. Light intensity is a measure of the rate of catalysis by luciferase and is therefore dependent upon temperature. The optimum temperature for luciferase activity is 20–25°C. It is important that the luciferase assay buffer, stored frozen, is fully equilibrated to room temperature for at least 30 min before use. 26. Serially dilute firefly luciferase (available from Promega) in the range of 1 fg to 1 ng into 20 µl of lysis buffer supplemented with 1 mg/ml BSA. Assay aliquots as above. 27. To aid uptake, 0.01% final concentration of non-ionic detergent such as Tween-80 can be added, but some detergents have been found to produce necrotic lesions on leaves and affect plant development (7). It is important to test this and include appropriate controls in your experiments. Alternatively, a 1 mM luciferin solution can be added to the growth media to be taken up by roots or detached plant parts. Particular plant structures, such as maturing seeds, may present more of a problem for luciferin access, requiring wounding to assist uptake before LUC can be detected (52). 28. As levels of CoA are low in plants cells, and oxyluciferin is released slowly from the enzyme complex, there will be a an initial high level of light produced from any accumulated LUC within cells, which slowly declines over a period of hours. Spraying of plants over a period of days is carried out to remove all accumulated LUC enzyme. The number of sprays necessary will depend on the strength of the promoter being used and the particular plant tissue being analysed.

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A time course should be established determining when there is no further increase in light production or change in tissue specific pattern. Note that high levels of luciferin have been reported to be toxic to some cells and tissues, and this too should be checked for your particular system. 29. For cooled cameras, it is therefore necessary to either employ a filter that can pass wavelengths at peak 560 nm and block those above 600 nm, or alternatively delay imaging plants for 5–10 min after exposure to light (53) to deal with the problem of chlorophyll auto-fluorescence, which has an emission maximum at 685 nm. Conversely, pH effects not only LUC activity but also shifts the bioluminescence towards the red end of the spectrum below pH 7.0. Hence, when using an intensified CCD camera, care is needed to show that apparent drops in LUC activity are not associated with changes in cellular pH (7). Hence, subcellular targeting of LUC may need to be considered carefully because of differing pH levels in sub-cellular compartments. 30. Note: it will take around 2 h for such cameras to reach the −120°C working temperature. Warning: the darkroom housing the CCD camera should be well ventilated to prevent O2 depletion by evaporated liquid nitrogen. Ideally, the O2 levels within the darkroom should be monitored. 31. To reduce background chlorophyll auto-fluorescence in photosynthetic tissues when using a cooled CCD camera, plants are placed in the dark prior to image capture and worked with, if necessary, under green safe light. 32. The length of exposure will depend on a number of parameters, the most important being the strength of the promoter being used to drive the reporter gene. The lens in use and the f-stop employed will also affect exposure times. A strong constitutive promoter such as the CaMV35s can be easily imaged by exposures in the range of 1–5 min; for other constructs, 30 min exposures are not uncommon (8). 33. Cosmic rays can also be detected by the sensor but they are usually in the form of a single high intensity pixel unlike the excitation pattern from LUC-expressing plants. Images from untransformed controls should indicate the likelihood of detecting cosmic rays, which varies depending on solar activity. 34. It is well to remember, though, that improved resolution can be at the expense of emission signal; hence, confocal imaging will not necessarily enhance a weak signal with low signal-tonoise ratio. 35. In fluorescence microscopes, light from the lamp/laser is first filtered through an excitation filter that permits selected

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wavelengths to pass through towards the specimen. The dichromatic mirror (or beam splitter) functions to direct the excitation wavelengths on to the specimen while at the same time allowing the returning emission wavelengths to pass on to the barrier filter which itself lets through only selected emission wavelengths. Filters can be termed as either shortor long pass (relating to wavelength) or band pass, which can be broad or narrow. Band-pass filters will be more specific but give rise to a weaker signal, while long-pass filters lead to stronger signals but more background. 36. Other approaches to dealing with the problem are using dual-wavelength excitation or emission correction, where the sample is either excited sequentially at two wavelengths, one to excite the FP and the other to excite the auto-fluorescence, or data is captured at two wavelengths. Image correction is then employed to separate out the fluorescence spectra detected. The use of chemicals to quench auto-fluorescence is also documented (55). 37. It should be kept in mind that the chromophore is fluorescent only when encapsulated; denatured FPs are poorly fluorescent, if at all, while truncations lead eventually to loss of fluorescence. Therefore, any fusions must allow for correct folding to take place, a parameter that can only really be established empirically. FP signal intensity can also be adversely affected by targeting to some locations, such as the vacuole, where proteolytic degradation can occur or the cell wall where low pH can be an issue, particularly for YFPs. 38. Warning: care needs to be taken when using a mercury lamp: refer to the manufacturer’s safety information before use. UV light can also damage your eyes and should never be viewed directly. The microscope should be fitted with a safety screen to cover specimens on the microscope stage. 39. For epi-fluorescence detection, samples can be either immersed in water or a simple medium, or viewed through air (taking care that the material does not become dehydrated (66). Note that some media exhibit significant levels of auto-fluorescence and this should be checked before use. For higher power microscopy and confocal studies, where it is necessary to bring the sample closer to the objective lens, samples should be mounted in water or the medium and covered with a cover slip of the correct thickness. 40. It is important to establish that your material is undergoing as little stress as possible; long periods spent under a cover slip, for example, can induce anerobic conditions, but conversely samples can speedily dry out when illuminated if not immersed. As general rule, dead, dying, or stressed cells will

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produce higher levels of auto-fluorescence, and developmentally your data will be questionable; so keeping the sample in the best possible conditions is of prime importance. Reviews detailing the use of imaging of live plant material should be consulted (18, 66), and some time should be spent carefully examining untransformed material under different conditions in brightfield. It appears rare for FPs to cause toxic effects in cells (the tetrametric forms may be an exception to this); but a consequence of requiring O2 for chromophore formation is the production of H2O2, equimolar with protein maturation, which may account for early reports suggesting thathigh expression of GFP was deleterious in plants (59). Subcellular targeting to the endoplasmic reticulum appeared to resolve this. Targeted FP may also help prevent fixation artefacts due to leakage of the small FP proteins from cells via symplastic connections. 41. It is possible to visualize GFP in fixed and embedded material, but auto-fluorescence is often enhanced and GFP fluorescence can be adversely affected by dehydration in ethanol (55). 42. If required, material can be counterstained to help show cellular structure. For example, imaging cell walls in root tips can be enhanced by staining for a few minutes in 10 µg/ml propidium iodide solution before mounting. 43. For GFPs, three common filters in use are GFP1 (Ex 425 nm (60), DB 470 nm, Em 480 nmLP); GFP2 or (GFP Plus) (Ex 480 nm (40), DB 505 LP, Em 510 nmLP); and GFP3 or (GFP plant) (Ex 470 nm (40), DB 495 nm, Em 525 (50)). Having a choice of filters can be invaluable in determining whether the fluorescence observed is from the FP or auto-fluorescence originating from the material. In some instances, a narrow band-pass emission filter may not discriminate between auto-fluorescence and the FP, whereas a long-pass emission filter can allow the colour of the FP and the auto-fluorescence to be distinguished. To aid your identification of the fluorescence emanating from the particular FP you are using, studying both untransformed material and a known positive transformant can be very useful. 44. UV light is damaging to the specimen and will, over time, photobleach the FP, so limit exposure to the mercury lamp to a minimum. 45. Image acquisition and manipulation are important skills to master (66). Quantification of GFP in images can be made using suitable software (see section on LUC analysis). 46. Excellent confocal and wide-field microscope primers can be obtained on the web (67).

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eliminating endogenous beta-glucuronidase background in barley. Plant Sci. 105, 63–69. Van der Eycken, E., Terryn, N., Goeman, J. L., Carlens, G., Nerinckx, W., Claeyssens, M., Van der Eycken, J., Van Montagu, M., Brito-Arias, M. and Engler, G. (2000) Sudan-beta-D-glucuronides and their use for the histochemical localization of betaglucuronidase activity in transgenic plants. Plant Cell Rep. 19, 966–970. Fleming, A. J., Manzara, T., Gruissem, W. and Kuhlemeier, C. (1996) Fluorescent imaging of GUS activity and RT-PCR analysis of gene expression in the shoot apical meristem. Plant J. 10, 745–754. van der Krol, A. R., van Poecke, R. M. P., Vorst, O. F. J., Voogt, C., van Leeuwen, W., Borst-Vrensen, T. W. M., Takatsuji, H. and van der Plas, L. H. W. (1999) Developmental and wound-, cold-, desiccation-, ultraviolet-B-stress-induced modulations in the expression of the petunia zinc finger transcription factor gene ZPT2–2. Plant Physiol. 121, 1153–1162. de Ruijter, N. C. A., Verhees, J., van Leeuwen, W. and van der Krol, A. R. (2003) Evaluation and comparison of the GUS, LUC and GFP reporter system for gene expression studies in plants. Plant Biol. 5, 103–115. van Leeuwen, W., Ruttink, T., Borst-Vrenssen, A. W. M., van der Plas, L. H. W. and van der Krol, A. R. (2001) Characterization of position-induced spatial and temporal regulation of transgene promoter activity in plants. J. Exp. Bot. 52, 949–959. Billinton, N. and Knight, A. W. (2001) Seeing the wood through the trees: a review of techniques for distinguishing green fluorescent protein from endogenous autofluorescence. Anal. Biochem. 291, 175–197. Abdel-Aal, E. S. M. and Hucl, P. (1999) A rapid method for quantifying total anthocyanins in blue aleurone and purple pericarp wheats. Cereal Chem. 76, 350–354. Doshi, K. M., Eudes, F., Laroche, A. and Gaudet, D. (2006) Transient embryo-specific

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expression of anthocyanin in wheat. In Vitro Cell. Dev. Biol.-Plant 42, 432–438. Abdel-Aal, E. S. M., Young, J. C. and Rabalski, I. (2006) Anthocyanin composition in black, blue, pink, purple, and red cereal grains. J. Agr. Food Chem. 54, 4696–4704. Haseloff, J., Siemering, K. R., Prasher, D. C. and Hodge, S. (1997) Removal of a cryptic intron and subcellular localization of green fluorescent protein are required to mark transgenic Arabidopsis plants brightly. P.N.A.S. USA 94, 2122–2127. Farrell, L. B. and Beachy, R. N. (1992) Review of the use of the GUS gene for analysis of secretory system, in GUS Protocols, (Gallagher, S. R., ed.), Academic, San Diego, pp. 127–133. Kavanagh, T. A., Jefferson, R. A. and Bevan, M. W. (1088) Targeting a foreign protein to chloroplasts using fusions to the transit peptide of a chlorophyll A/B protein. Mol. Gen. Genet. 215, 38–45. Huttly, A. K. and Baulcombe, D. C. (1989) A wheat alpha-AMY2 promoter is regulated by gibberellin in transformed oat aleurone protoplasts. EMBO J. 8, 1907–1913. Stangeland, B. and Salehian, Z. (2002) An improved clearing method for GUS assay in Arabidopsis endosperm and seeds. Plant Mol. Biol. Rep. 20, 107–114. Ruzin, S. E. (1999) Plant Microtechnique and Microscopy. OUP, New York. eNotes, P. (2007) Compatibility of the Pierce BCA protein assay with Promega lysis buffers and lytic assay reagents. Available online: http://www.promega.com/enotes/ applications/ap0047_tabs.htm Shaw, S. L. (2006) Imaging the live plant cell. Plant J. 45, 573–598. Microscope, Microscope Primers. Available online: http://micro.magnet.fsu.edu/primer/techniques/confocal/index.html, http://www. hi.helsinki.fi/amu/AMU%20Cf_tut/cf_tut_ part1.htm, http://www.olympusfluoview. com/theory/confocalintro.html.

Chapter 4 Biolistics Transformation of Wheat Caroline A. Sparks and Huw D. Jones Abstract We present a complete, step-by-step guide to the production of transformed wheat plants using a particle bombardment device to deliver plasmid DNA into immature embryos and the regeneration of transgenic plants via somatic embryogenesis. Currently, this is the most commonly used method for transforming wheat and it offers some advantages. However, it will be interesting to see whether this position is challenged as facile methods are developed for delivering DNA by Agrobacterium tumefaciens or by the production of transformants via a germ-line process (see other chapters in this book). Key words: Wheat, biolistics, particle bombardment, transgenic plant.

1. Introduction Wheat transformation using biolistics has become a robust platform technology that can routinely produce between 5 and 20 independent transgenic plants per 300 immature embryos bombarded. It is less genotype dependent than Agrobacteriumbased methods and is generally more efficient, although this may change as the relatively more recent Agrobacterium protocols become widely adopted and optimized. It is also less demanding with respect to vector requirements: the trait gene being co-bombarded with a separate, selectable marker plasmid. However, the DNA integration events are often more complex and contain more transgene copies. This last point is particularly important if regulatory approval is needed for field trails, or if known transgene–genomic DNA junctions are required, for example, to ‘sequence walk’ out of the transgene insertion into neighbouring plant DNA. Huw D. Jones and Peter R. Shewry (eds.), Methods in Molecular Biology, Transgenic Wheat, Barley and Oats, vol. 478 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-379-0_4

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Compared to some species, wheat offers only a few explant tissues suitable for tissue-culture regeneration. The most common target tissue used is the scutellum surface of the immature seed embryo which is amenable to DNA uptake via both biolistics and Agrobacterium and readily forms embryogenic callus. The protocol below focuses on this explant, although it could be readily adapted for immature inflorescence, an alternative tissue that has been used to successfully produce transgenic wheat plants (1–7). The protocol described here is based on the work of Paul Lazzeri, and others (4,8–14) who worked at Rothamsted Research during the 1990s, and it has evolved into its current form through adaptation and optimization. For the purposes of this chapter, the protocol has been divided into sections with a step-wise guide to the following: preparation of donor material, coating gold particles with DNA, particle bombardment, callus induction, regeneration, and selection. This method has been fine-tuned to transform the spring variety Cadenza (cpb-Twyford, UK) but with minor modifications, to take into account genotype-specific tissue culture responses, has also been used to transform over 35 wheat and associated genotypes including two T. turgidum ssp. durum varieties (6, 15) and the durum wheat/ barley hybrid Tritordeum (3, 5, 7, 10, 16). It is used routinely in association with the selectable marker genes bar and nptII and the respective selective agents phosphinothricin (PPT) and G418 (13), although other gene and selective agent combinations could also be used. Transformation, both for stable plants and transient expression studies, is widely used as a reverse genetics tool for candidate gene validation in wheat. A wide range of trait targets has been investigated using a transgenic approach in wheat including glutenin functionality (15, 17–32), starch (33–36), grain hardness (37, 38, 39), pre-harvest sprouting (40), etc. Transient expression and stably transformed plants have also been frequently used for promoter analysis studies in wheat (34, 41– 45).

2. Materials 2.1. Growth of Donor Plants

The condition of wheat donor plants (Triticum aestivum L.) is critical to successful transformation. In order to provide healthy plants with consistent quality, plants are grown as follows (see Note 1): 1. Soil composition (Petersfield Products, Leicestershire, UK): The following table gives the details.

Biolistics Transformation of Wheat

Component

Amount

Fine-grade peat

75%

Screened sterilized loam

12%

Screened lime-free grit

10%

Medium vermiculite

3%

Osmocote plus (slow release fertilizer, 15N/11P/13K plus micronutrients)

2 kg/m3

PG mix (14N/16P/18K granular fertilizer plus micronutrients)

0.5 kg/m3

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2. Planting: Five plants per 21 cm diameter plastic pot (Nursery Trades (Lea Valley) Ltd., Hertfordshire, UK). Plants are stripped to leave five tillers per plant, once the plants are 6–8 weeks old. 3. Vernalization: For winter wheat varieties, it is carried out at 4–5°C for 8 weeks from sowing. 4. Growth room conditions: 18–20°C day and 14–15°C night temperatures under a 16-h photoperiod provided by banks of HQI lamps 400 W (Osram Ltd., Berkshire, UK) to give an intensity of approximately 700 µmol/m/s photosynthetically active radiation (PAR) (see Note 2). 5. Watering: Initially all plants are top-watered in order to monitor water requirements and thereby provide sufficient water without water logging. An automated flooding system is used once the root system reaches the base of the pot. 6. Pests and disease: These are kept to a minimum by restricting access to growth rooms and following good housekeeping practices. Any diseased plants are discarded immediately. To avoid mildew, the fungicide Flexity (BASF, Cheshire, UK) is applied as a preventative. Amblyseius cucumeris (Fargro, West Sussex, UK) is used as a biological control agent to manage thrips. 2.2. Sterilization Materials for Wheat Caryopses

Aqueous ethanol 70% (v/v) Aqueous Domestos 10% (v/v) (Lever Fabergé Ltd., Surrey, UK) Sterile water (see Note 3)

2.3. Stock Solutions of Basal Culture Media Components

Below are tables detailing stock solutions of basal culture media components: macrosalts, microsalts, vitamins, amino acids, hormones and stress inducers, selection agents, and agargel, from which the final tissue culture media are prepared (see Notes 3 and 4).

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1. Macrosalts Component

MS macrosalts (×10) (g/l)

L7 macrosalts (×10) (g/l)

NH4NO3

16.5

2.5

KNO3

19.0

15.0

KH2PO4

1.7

2.0

MgSO4·7H2O

3.7

3.5

CaCl2·2H2O (see Note 5)

4.4

4.5

Sterilization

Autoclave at 121°C for 20 min

Autoclave at 121°C for 20 min

Storage (see Note 6)

4°C

4°C

2. Microsalts Component

L7 microsalts (×1,000) (g/l)

MnSO4 (see Note 7)

15.0

H3BO3

5.0

ZnSO4·7H2O

7.5

KI

0.75

Na2MoO4·2H2O

0.25

CuSO4·5H2O

0.025

CoCl2·6H2O

0.025

Sterilization (see Note 8)

Prepare 100 ml at a time. Filter sterilize

Storage (see Note 6)

4°C

3. Vitamins

Component

MS vitamins (-glycine) (×1,000) (g/l)

L7 vitamins + myoinositol (×200) (g/l)

Thiamine HCl

0.1

2.0

Pyridoxine HCl

0.5

0.2

Nicotinic acid

0.5

0.2

Myo-inositol

–

40.0

Ca-pantothenate

–

0.2

Ascorbic acid

–

0.2

Sterilization (see Note 8) Prepare 100 ml at a time.Filter sterilize

–

Storage (see Note 6)

−20°C in 10 ml aliquots

4°C

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75

4. Amino acids Component

3AA amino acids (g/l)

L-Glutamine

18.75

L-Proline

3.75

L-Asparagine

2.5

Storage (see Note 6)

−20°C in 40 ml aliquots

5. Hormones and stress inducers

Hormone/chemical

Conc. (mg/ml)

Solute

Sterilization (see Note 8)

Storage (see Note 6)

2,4-Dichlorophenoxyacetic 1 acid (2,4-D)

Dissolve powder in ethanol. Add water to volume

Filter sterilize

−20°C in 1 ml aliquots

Zeatin (mixed isomers)

10

Dissolve powder in small volume 1 M HCl and make up to volume with water

Filter sterilize

−20°C in 1 ml aliquots

Silver nitrate (AgNO3) (see Note 9)

20

Water

Filter sterilize

−20°C in 1 ml aliquots in the dark

Copper (II) sulfate (CuSO4·5H2O) (see Note 10)

25

Water

Filter sterilize

4°C in 1 ml aliquots

6. Selection agents

Selection agent

Conc. (mg/ml)

Solute

Sterilization (see Note 8)

Storage (see Note 6)

Glufosinate ammonium (see Note 11)

10

Water

Filter sterilize

−20°C in 1 ml aliquots

Geneticin disulfate (G418) (see Note 12)

50

Water

Filter sterilize

−20°C in 1 ml aliquots

7. Agargel Agargel (×2) Conc. (g/l) Solute

Sterilization

Storage

Agargel

Autoclave at 121°C for 20 min

Room temperature

10

Water

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2.4. Induction Media

1. MSS 3AA/2 9%S (×2) Component

MSS 3AA/2 9%S (×2)

MS macrosalts

200 ml/l

L7 microsalts

2 ml/l

Ferrous sulfate chelate solution (×100)

20 ml/l

MS (-glycine) vitamins

2 ml/l

Myo-inositol

200 mg/l

3AA amino acids (see Note 14)

40 ml/l

Sucrose (see Note 15)

180 g/l

pH

pH to 5.7 with 5 M NaOH or KOH

Osmolarity

800–1,100 mOsM

Sterilization (see Note 8)

Filter sterilize

Storage (see Note 6)

4°C

2. MS 9% 0.5DAg Mix an equal volume of MSS 3AA/2 9%S (×2) with sterilized, melted agargel (×2). Add 0.5 mg/l 2,4-D (see Note 16) and 10 mg/l AgNO3 and pour into 9-cm diameter Petri dishes (approximately 28 ml per dish). Store at 4°C in the dark (see Notes 9, 13 and 17). 2.5. Regeneration Media

1. R (×2) Component

R (×2)

L7 macrosalts

200 ml/l

L7 microsalts

2 ml/l

Ferrous sulfate chelate solution (×100)

20 ml/l

L7 vitamins/inositol

10 ml/l

Maltose

60 g/l

pH

pH to 5.7 with 5 M NaOH or KOH

Osmolarity

269–298 mOsM

Sterilization (see Note 8)

Filter sterilize

Storage (see Note 6)

4°C

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2. RZDCu Mix an equal volume R (×2) with sterilized, melted agargel (×2). Add 5 mg/l zeatin, 0.1 mg/l 2,4-D, and 25 mg/l CuSO4 (see Note 10) and pour into 9-cm Petri dishes (approximately 28 ml per dish). Store at 4°C (see Notes 13 and 17). 2.6. Selection Media

1. RZPPT4 or RZG50 – Mix an equal volume of R (×2) with sterilized, melted agargel (×2) and add 5 mg/l zeatin and 4 mg/l glufosinate ammonium (PPT4) or 50 mg/l G418 (G50) (see Note 18). Pour into 9-cm Petri dishes (approximately 28 ml per dish). Store at 4°C (see Notes 13 and 17). 2. RPPT4 or RG50 – Mix an equal volume of R (×2) with sterilized, melted agargel (×2) and add 4 mg/l glufosinate ammonium (PPT4) or 50 mg/l G418 (G50) (see Note 18). Pour into 9-cm Petri dishes (approximately 28 ml per dish) or GA-7 Magenta vessels (Sigma-Aldrich) (approximately 60 ml per vessel). Store at 4°C (see Notes 13 and 17).

2.7. Bombardment Materials/ Consumables

3. Gold particles: 0.6 µm (sub-micron) gold particles (BIO-RAD Laboratories, Hertfordshire, UK) (see Note 20) (for preparation, see step 1 – Section 3.2).

Solute

Sterilization Storage (see Note 8) (see Note 6)

Component

Conc.

1.

Calcium chloride (CaCl2·2H2O)

2.5 M

Dissolve 3.67 g in 10 ml water

Filter sterilize

−20°C in 50 µl aliquots

2.

Spermidine free base (see Note 19)

1M

Dissolve 1 g bottle spermidine powder in 6.89 ml sterile water

–

−80°C in 20 µl aliquots

0.1 M

Prepare 1:10 dilution of 1 M stock in sterile water under sterile conditions

–

Store immediately at −20°C in 10 µl volumes

4. Macrocarriers, stopping screens, 650 or 900 psi rupture discs (all BIO-RAD Laboratories) (see Note 21). 5. Plasmid DNA: Prepare using Qiagen Maxi-prep kit (Qiagen Ltd., West Sussex, UK) and resuspend at 1 mg/ml in sterile TE buffer (pH 8.0) or sterile water. Store in 20 µl aliquots at −20°C (see Note 22).

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3. Methods 3.1. Preparation of Donor Material

The method described below has been optimized for transformation of immature scutella but immature inflorescences can be used as alternative explants. These may be more responsive than immature scutella for certain varieties, e.g. T. aestivum vars. Baldus and Brigadier, and other Triticeae, e.g. T. turgidum ssp. durum and tritordeum (see Notes 23 and 24). ● Collection and sterilization of wheat caryopses 1. Collect spikes from growth-room-grown plants approximately 10–12 weeks after sowing (see Note 25). 2. Remove the caryopses from the panicles (see Note 26). 3. Sterilize the caryopses by rinsing in 70% (v/v) aqueous ethanol for 5 min; then soak in 10% (v/v) Domestos for 15–20 min with occasional shaking. 4. Rinse copiously with at least three changes of sterile water. Maintain the sterilized caryopses in moist conditions but do not keep immersed in water. ●

Isolation and pre-culture of immature scutella 1. In a sterile environment, isolate the embryos microscopically (see Fig. 1a) and remove the embryo axis to prevent precocious germination (see Note 27). 2. Place 25–30 scutella within a central target area of a 9-cm Petri dish containing induction medium (MS9%0.5DAg), orientating them with the cut embryo surface in contact with the medium, i.e. the uncut scutellum is bombarded (see Note 28, Fig. 1b). 3. Seal the plates with Nescofilm (Fisher Scientific UK) and pre-culture the prepared donor material in the dark at 22°C for 1–2 days prior to bombardment (see Note 29).

3.2. Coating Gold Particles with DNA

●

Preparation of gold particles 1. Place 20 mg BIO-RAD sub-micron gold particles (0.6 µm) in a 1.5-ml Eppendorf and add 1 ml 100% ethanol. Sonicate for 2 min, pulse-spin in a microfuge for 3 s, and remove the supernatant. Repeat this ethanol wash twice more. 2. Add 1 ml sterile water and sonicate for 2 min. Pulse-spin in a microfuge for 3 s and remove the supernatant. Repeat this step. 3. Add 1 ml sterile water and resuspend fully by vortexing. Aliquot 50 µl into sterile 1.5ml Eppendorf tubes, vortexing between taking each aliquot to ensure an equal distribution of particles. Store at −20°C.

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Fig. 1. (a) Isolation of immature embryo from wheat caryopsis. (b) Immature scutella plated for bombardment. (c) Formation of embryogenic callus. (d) Regeneration of plantlet on selective medium. (e) Regenerated transformed plants growing in GM containment glasshouse. (f) Gus expression in leaf section (top), control leaf (bottom). (See Color Plate 1)

Coating of gold particles with DNA for bombardment The following procedure should be carried out on ice, in a sterile environment. ●

1. Allow a 50 µl aliquot of prepared gold (see step 1 – Section 3.2) to thaw at room temperature, and then sonicate for 1– 2 min (see Note 30). The tubes can be vortexed following sonication to ensure total resuspension, particularly if the aliquots are to be sub-divided for smaller preparations (see Note 31). 2. Add 5 µl DNA (1 mg/ml in TE or water (see Note 32) or water (see Note 33) and vortex briefly to ensure good contact of DNA with the particles (see Note 34). 3. Place 50 µl 2.5 M CaCl2 and 20 µl 0.1 M spermidine into the lid of the Eppendorf and mix together (see Note 35). Briefly vortex into the gold/DNA solution. Centrifuge in a microfuge at top speed for 3–5 s pulse to pellet the DNA-coated particles. Discard the supernatant.

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4. Add 150 µl of 100% ethanol to wash the particles, resuspending them as fully as possible (see Notes 36 and 37). Centrifuge in a microfuge at top speed for 3–5 s pulse to pellet the particles, and discard the supernatant. 5. Resuspend fully in 85 µl 100% ethanol and maintain on ice (see Note 38). 3.3. Particle Bombardment Using the PDS-1000/He Particle Gun (BIORAD) (see Note 39)

Note: The delivery system involves the use of high pressure to accelerate particles to high velocity. Appropriate safety precautions should be taken and safety spectacles should be worn when operating the gun. Delivery of DNA-coated gold particles In any bombardment experiment, controls should be included to monitor regeneration and selection efficiencies (see Note 40). ●

1. DNA-coated gold particles (see step 2 – Section 3.2) are delivered using the PDS-1000/He particle gun (BIORAD) (see Fig. 2) according to the manufacturer’s instructions. The following settings are maintained as standard for this procedure (see Note 41): gap 2.5 cm (distance between rupture disc and macrocarrier), stopping plate aperture 0.8 cm (distance between macrocarrier and stopping screen), target distance 5.5 cm (distance between stopping screen and target plate), vacuum 91.4–94.8 kPa, vacuum flow rate 5.0, vent flow rate 4.5 (8). 2. Sterilize the gun’s chamber and component parts by spraying with 90% (v/v) ethanol, which should be allowed to evaporate completely. 3. Sterilize macrocarrier holders, macrocarriers, stopping screens, and rupture discs by dipping in 100% ethanol and allow the alcohol to evaporate completely on a mesh rack in

Fig. 2. The PDS-1000/He particle gun (BIO-RAD) (left) and diagram of component parts as described in step 1 – Section 3.3 (right).

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a flow hood (see Note 42). Place the macrocarrier holders into sterile 6-cm Petri dishes and introduce one macrocarrier into each holder. 4. Briefly vortex the coated gold particles (see step 2 – Section 3.2), take a 5 µl aliquot and drop centrally onto the macrocarrier membrane. Allow to dry naturally, not in the air flow (see Note 43). 5. Load a rupture disc (650 or 950 psi) (see Note 21) into the rupture disc retaining cap (see Fig. 2) and screw into place on the gas acceleration tube, tightening firmly using the mini torque wrench (see Note 44). 6. Place a stopping screen into the fixed nest. Invert the macrocarrier holder containing macrocarrier + gold particles/ DNA and place over the stopping screen in the nest and maintain its position using the retaining ring. Mount the fixed nest assembly onto the second shelf from the top to give a gap of 2.5 cm (see Fig. 2). 7. Place a sample on the target stage on a shelf to give the desired distance; fourth shelf from the top gives a target distance of 5.5 cm. 8. Draw a vacuum of 91.4–94.8 kPa and fire the gun (see Note 45). 9. After firing, release the vacuum, remove the sample and disassemble the component parts, discarding the ruptured disc and macrocarrier (see Note 46). 10. Place the macrocarrier holder and stopping screen in 100% ethanol to re-sterilize if they are to be reused for further shots; otherwise place in 1:10 dilution Savlon (Novartis Consumer Health, West Sussex, UK) to soak. Sonicate for 10 min prior to reuse (see Note 47). 3.4. Culture of Immature Scutella Following Bombardment

●

Induction of embryogenic callus 1. Following bombardment, spread the scutella evenly across the medium, dividing each replicate between two or three plates of induction medium (MS9%0.5DAg), i.e. approximately 10 scutella per plate (see Note 48). 2. Seal the plates with Nescofilm and incubate at 22°C in the dark for induction of embryogenic callus (see Notes 49 and 50, Fig. 1c).

●

Regeneration and selection 1. After 3–5 weeks on induction medium, transfer any calli bearing somatic embryos to regeneration medium (RZDCu) in 9-cm Petri dishes and incubate at 22°C in the light for 3–4 weeks (see Notes 49 and 51).

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2. After 3–4 weeks on regeneration medium, transfer calli to RZ + selection in 9-cm Petri dishes with high lids (see Notes 52 and 53) (the selection medium is determined by the selectable marker gene in the transforming plasmid: RZPPT4 for bar or RZG50 for nptII (see Notes 11 and 12)). Seal the plates with Nescofilm and incubate at 22°C in the light (see Notes 49 and 54). 3. After a further 3–4 weeks, transfer surviving calli to regeneration medium + selection but without hormones (RPPT4 or RG50) in 9-cm Petri dishes with high lids (see Notes 53 and 55, Fig. 1d). Seal the plates with Nescofilm and incubate at 22°C in the light (see Note 49). 4. Once regenerating shoots are clearly defined and can be separated easily from the callus, transfer these to regeneration medium + selection but without hormones (RPPT4 or RG50) in GA-7 Magenta vessels, placing no more than 4–6 plantlets per Magenta. Incubate at 22°C in the light (see Note 49). ●

Potting putative transgenic plants to soil 1. Plantlets can be transferred to soil once a reasonable root system has been established and leaves are approximately 10–15 cm. Carefully remove plantlets from the agargelsolidified medium (rinsing the roots with water if necessary to remove excess agargel) and pot into soil in 8 cm square plastic pots (Nursery Trades, (Lea Valley) Ltd.). Place plantlets within a propagator to provide a high humidity for 1–2 weeks to acclimatize them from tissue culture, and grow in a GM containment glasshouse (see Notes 56 and 57). Typically it takes at least 3 months from bombardment to potting into soil. 2. Once suitably established (3–4 leaves), a leaf sample can be taken from which to extract genomic DNA for analysis by PCR to establish whether the plant is transformed. Once confirmed PCR positive, plants are re-potted to 13cm diameter pots (Nursery Trades (Lea Valley) Ltd.) and grown under the same glasshouse conditions (see Note 57). Plants should reach maturity in 3–4 months (see Fig. 1e). 3. Transgenic plants can be analysed in a number of ways: reporter gene expression can be assessed using, for example, the histochemical GUS test for uidA (46) (see Fig. 1f), UV visualization of GFP, herbicide leaf paint assay (47) and/or the ammonium test (48) for bar . Gene integrations can be studied using Southern analysis and fluorescent in situ hybridization (FISH).

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4. Notes 1. The conditions described are suitable for growth of T. aestivum plants, but for T. turgidum ssp. durum different growing conditions are necessary. 2. Although glasshouse-grown plants can be used, these tend to be more variable due to seasonal variation. 3. Reverse-osmosis, polished water with a purity of 18.2 MΩ;/ cm should be used for all solutions. 4. Modifications to the media detailed here may be required for alternative varieties or wheat species. For example, the choice of basal salts (MS or L7), the concentration of sugars (sucrose or maltose), the level of hormones, etc. need to be empirically determined. 5. Before mixing with other components, dissolve CaCl2·2H2O in water. 6. Sterile stock solutions can be stored at 4°C for 1–2 months. Some settling of salts may occur during storage, so the medium should be shaken well prior to use. Stock solutions stored at −20°C should remain effective for at least a year, provided no freeze/thawing has occurred. 7. MnSO4 is available in various hydrated states which will alter the required weight: for MnSO4·H2O, add 17.05 g/l; for MnSO4·4H2O, add 23.22 g/l; or for MnSO4·7H2O, add 27.95 g/l. 8. Filter sterilization is carried out using 0.2 µm filter size. For large volumes use MediaKap (NBS Biologicals Ltd., Cambridgeshire, UK), and for smaller volumes use a Nalgene syringe filter (Fisher Scientific UK). 9. AgNO3 is used to promote embryogenesis; silver thiosulfate (a mix of silver nitrate and sodium thiosulphate) at 10 mg/l can be used as an alternative. Both are photosensitive, so the stock solutions and any media plates containing them should be kept in the dark. 10. Copper sulfate is a stress-inducing agent (similar to silver nitrate) used to promote shooting. The preferred copper sulfate concentration is 100 µM (25 mg/l), but if too much shooting occurs, 50 µM can be used. 11. Glufosinate ammonium is synthetically produced phosphinothricin (PPT) bound to ammonium (Greyhound Chromatography and Allied Chemicals, Cheshire, UK), and is the active component in herbicides such as Basta. Bialaphos (phosphinothricylanalylanaline, sodium; Melford Laboratories Ltd.) at 3–5 mg/l is a successful alternative selection agent.

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12. Kanamycin, paromomycin, and neomycin are alternative aminoglycoside antibiotics that can be used for selection with the nptII gene. However, although they may be successful for some plant species, they are not recommended for wheat, as untransformed tissues exhibit natural resistance. Geneticin disulfate (G418) is available from Melford Laboratories Ltd. 13. The agargel solution should be shaken well both before and after autoclaving to avoid non-uniform solidification which leads to difficulties when remelting. 14. Instead of using the 3AA stock solution, 0.75 g/l L-glutamine, 0.15 g/l L-proline, and 0.1 g/l L-asparagine can be added individually. 15. Sucrose (9%) partially plasmolyses the cells during pre-culture, which may increase their ability to withstand bombardment. However, this is variety and species dependent and 3% sucrose is often suitable, e.g., for T. turgidum ssp. durum scutella. The osmolarity for 3% sucrose medium should be within the range 355–398 mOsM. 16. Picloram (Sigma-Aldrich) at 2–6 mg/l can be used as an alternative auxin (8, 10). 17. Tissue culture media should be prepared as freshly as possible and not be stored for more than 2–3 weeks in Petri dishes and Magenta vessels. However, media should be prepared a few days in advance to allow any contamination to be detected before use. To minimize condensation in the plates, allow the agargel to cool once melted, and pour the final medium at approximately 50°C. 18. The selection agent should be used at a concentration which is known to fully inhibit the growth of non-transformed explants. However, the concentration should be gauged according to the development of the cultures at each transfer stage. Generally, use within the range 2–6 mg/l glufosinate ammonium (PPT) and 25–50 mg/l G418. 19. Spermidine is hygroscopic and oxidizable, and consequently difficult to weigh. A 1-g bottle should therefore be dissolved directly in sterile water. Spermidine also deaminates with time, therefore solutions should be maintained below −20°C, preferably at −80°C, and any unused aliquots once thawed, should be discarded. 20. Heraeus gold particles of 0.4–1.2 µm diameter (W.C. Heraeus GmbH & Co. KG, Hanau, Germany) have also been used successfully for transformation. However, the sub-micron BIO-RAD particles give more consistent results for wheat. The smaller, more uniform size of the latter particles is pref-

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erable for small wheat cells, but for other species larger particles may be suitable. 21. Rupture pressures of 650 or 900 psi have been found to be optimal for the wheat varieties reported here; 450 or 1,100 psi pressures will result in successful transformation but with lower efficiency. Rupture discs of 450, 650, 900, 1,100, 1,350, 1,550, 1,800, 2,000, 2,200 psi are available, and a range should be tested if attempting transformation of any new variety or species. 22. Plasmids tend to be pUC-based and contain one or more gene cassettes. To allow selection of transformed tissues, a selectable marker gene is required: for example, the bar or nptII gene under the control of a constitutive promoter (e.g. maize ubiquitin or CaMV35S) and a suitable terminator (e.g. nos). The bar gene confers resistance to the herbicides Basta (glufosinate ammonium/PPT) and Bialaphos, and the nptII gene confers resistance to the antibiotics geneticin disulfate (G418), kanamycin, neomycin, paromomycin, etc. (see Note 12). A reporter gene (e.g. uidA, luc, or gfp) is also a useful tool to monitor both transient and stable transformation (see Fig. 1f). These marker genes can be located in the same plasmid or on separate plasmids co-precipitated onto the gold particles. 23. The methods detailed in this chapter have been used for transformation of a number of commercial wheat varieties but with a range of efficiencies; Cadenza, Canon, and Florida have given the highest efficiencies (up to 13%) (8, 9). T. turgidum ssp. durum (e.g. cvs. Ofanto and Venusia) can also be transformed by this method (6, 49) but for these and alternative wheat varieties, modifications may be required (4, 5). 24. Immature inflorescences are an alternative explant for transformation, as they can have high regeneration potential, and for certain varieties these may be more responsive than immature scutella. References describing modifications necessary when using immature inflorescences are the following: for T. aestivum varieties (1, 8); for T. turgidum ssp. durum (6, 49); for barley (5); and for tritiordeum (a fertile cereal amphidiploid obtained from crosses between Hordeum chilense and durum wheat cultivars, and containing the genome HCHHCHAABB) (3, 4, 16). 25. Embryos at the correct stage are usually found approximately 12–16 days post anthesis. A few caryopses can be opened at the time of collection to determine the size of the embryos within. Although it is not encouraged, if the caryopses will not be used the same day, it is possible to store the spikes intact at 4°C, with stems in water.

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26. Avoid using the inner caryopses of the spikelet, as these generally contain smaller embryos because of asynchronous development. 27. The most responsive size range for the varieties reported here is 0.5–1.5 mm length; smaller and larger embryos may respond but with much lower efficiencies. Size is not quite as important for transient experiments. 28. Typically the gun shot fires most gold particles within a central approximately 2-cm diameter circular area of a Petri dish. Arranging scutella within this area maximizes particle delivery (as shown by transient expression studies (8)). 29. The pre-culture phase allows the tissues to recover from the isolation procedure before being subjected to bombardment and may also pre-plasmolyse the cells (see Note 15). However, it also allows any contamination to be detected prior to bombardment. Should it be difficult to sterilize donor material resulting in contaminated explants, Plant Preservative Mixture (PPM) (Plant Cell Technology Inc., Washington D.C., USA) can be included in tissue culture media at 1 ml/l. This is a non-toxic, broad-spectrum preservative and biocide that does not interfere with callus proliferation or regeneration. 30. The sonication has worked effectively if the gold particles have resuspended in the liquid rather than being present as a pellet in the base of the tube. However, there is evidence that over-sonication can cause aggregation, so the particles should not be sonicated longer than 1–2 min. 31. If fewer shots are required or a variety of DNAs are to be compared, the gold preparation can be sub-divided and volumes scaled down accordingly. 32. If plasmids are not at a concentration of 1 mg/ml, recalculate the volume to give 5 µg DNA and add to the gold. Too large a volume of DNA should be avoided, however; if the DNA is very dilute, reprecipitate the DNA and resuspend at a higher concentration. 33. In order to monitor regeneration and selection efficiencies of a bombardment experiment, control plates are required (see Note 40). Some particles should therefore be prepared without DNA, replacing the DNA solution with sterile water. 34. The standard amount of DNA is 5 µg/50 µl gold suspension. If using more than one plasmid, i.e. for co-bombardment, the amounts of DNA added should be calculated such that equimolar quantities are used, with a total of 5 µg DNA for the two plasmids (greater than 5 µg may cause clumping of particles). Alternatively, different ratios can be used to skew for the gene of interest, i.e. 1.5 plasmid of interest:1

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selectable marker construct; plants surviving selection will then have an increased probability of containing both selectable marker and the gene of interest. 35. CaCl2 and spermidine act to bind, stabilize, and precipitate the DNA. Precipitation onto the gold particles is very rapid so the CaCl2 and spermidine are mixed first to ensure that the coating is as even as possible. 36. The gold must be fully resuspended at this stage, as the remaining clumps cannot be removed during later resuspension steps. The particles should be resuspended as well as possible by scraping the side of the tube with the pipette tip to remove clumps, and drawing up and expelling the solution repeatedly. Vortexing will not aid resuspension. 37. Ideally, the coated particles should be used as soon as possible; however, they can be kept on ice at this point (but for no longer than an hour), completing the rest of the protocol just prior to use. 38. Avoid aspirating too much at this stage, as the ethanol will evaporate and increase the final concentration of particles. Some natural evaporation means there is generally enough for only 10–12 shots from the 85 µl final volume, even though there should be sufficient for 16–17 shots (5 µl/ shot). In order to reduce further evaporation of the ethanol before the resuspended particles are required, the Eppendorf lids can be sealed with Nescofilm. However, it is advisable to use coated gold particles as soon as possible. 39. Agrobacterium tumefaciens-mediated transformation of wheat is a viable, alternative DNA delivery system. Transformation protocols have been reported elsewhere (47, 50, 51). 40. Various control plates should be included within each experiment: unbombarded to monitor the development/ regeneration of donor tissue; bombarded with gold (no DNA) and unselected to monitor tissue culture response following bombardment; and bombarded with gold (no DNA) and selected to monitor the effects of the selection on regeneration. 41. Although these settings were found to be optimal for the wheat varieties routinely used, they may need to be altered for different varieties or species. 42. The rupture discs are composed of laminate layers, therefore they should not be sterilized for more than 10 min or the layers may become separated. 43. Once the coated particles have been dispensed onto the macrocarriers, the ethanol should be allowed to evaporate slowly. The flow hood may cause vibration, which could

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cause particle agglomeration, so in order to create an even spread of dried particles on the macrocarrier, place macrocarriers within their sterile Petri dishes outside of the flow hood on a non-vibrating surface. Macrocarriers should be used when recently dried, so only a few should be loaded with gold at any one time. Macrocarriers can be examined microscopically prior to bombardment to determine the uniformity and spread of particles, discarding any that have agglomerated clumps of gold, which will reduce transformation efficiency. 44. The helium pressure on the cylinder should be set to approximately 200 psi more than the intended rupture pressure. 45. The helium pressure accumulates until the rupture disc breaks, propelling the macrocarrier onto the stopping plate, thereby releasing and dispersing the gold particles. The actual pressure at which the rupture disc bursts should be monitored to ensure a successful shot, otherwise transformation efficiencies may be affected. 46. Following a shot, the macrocarrier can be observed microscopically to visualize the mesh pattern left by the stopping screen. This will demonstrate how much gold has been released/ retained. 47. The macrocarriers and stopping screens are sonicated to destroy any adhering DNA and prevent carry-over to future bombardments. 48. The scutella are spread more evenly in order to reduce the culture density and prevent competition for nutrients. 49. Incubation is carried out in a controlled environment room with a 12-h photoperiod provided by cool white fluorescent tubes emitting lighting levels approximately 250 µmol/m2/s PAR. The room temperature is set at 22°C at the level of the cultures under the lights. Trays are covered with foil to create darkness for the callus induction phase. 50. Transient assays, e.g. histochemical gus assay, can be carried out after 1–3 days depending on the construct, i.e. strength of the promoter. 51. At this stage, whole calli should be transferred without division, placing approximately 10 calli per plate. 52. The induction period for somatic embryogenesis is usually 3–5 weeks. However, the explants should be observed regularly to check for contamination. Judgement and experience is required to monitor development in order to determine the best time for transfer to regeneration medium; transfer is carried out when the embryogenic callus has mature somatic embryos some of which may just be forming small shoots.

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53. ‘High lids’ are created by using the upturned base of another Petri dish as the lid. This provides greater height for growth of shoots. 54. Selection is generally applied at the second and subsequent transfers, until all control plantlets have been killed (see Note 40). However, selection can be introduced earlier, i.e. at callus induction or at the first round of regeneration. This may serve to reduce the numbers of calli and/or plantlets surviving but may also result in loss of transformants if they are not strong enough to survive selection early on. 55. The number of calli per 9-cm Petri dish should be reduced to prevent overcrowding if the regenerating calli to be transferred are large. The callus can be divided if necessary but each of the callus pieces should be monitored in order to trace plants with possible clonal origin. 56. Tissue-cultured plantlets have little or no waxy cuticle, so are particularly prone to desiccation after transfer to soil; a propagator maintains humidity while the cuticle forms. 57. Glasshouse conditions are 18–20°C day and 14–16°C night temperatures with a 16-h photoperiod provided by natural light supplemented with banks of SonT 400 W sodium lamps (Osram Ltd.) giving 400–1,000 µmol/m2/ s PAR.

Acknowledgements Rothamsted receives grant-aided support from the Biotechnological and Biological Sciences Research Council, UK. We acknowledge other members of the Rothamsted Cereal Transformation Group, past and present, for their significant contribution to the protocols described here.

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15. He, G. Y., Rooke, L., Steele, S., Bekes, F., Gras, P., Tatham, A. S., Fido, R., Barcelo, P., Shewry, P. R. and Lazzeri, P. A. (1999) Transformation of pasta wheat (Triticum turgidum L. var. durum) with high-molecularweight glutenin subunit genes and modification of dough functionality. Mol. Breed. 5, 377–386. 16. Barcelo, P., Vazquez, A. and Martin, A. (1989) Somatic embryogenesis and plant regeneration from Tritordeum. Plant Breed. 103, 235–240. 17. Blechl, A. E. and Anderson, O. D. (1996) Expression of a novel high-molecular-weight glutenin subunit gene in transgenic wheat. Nat. Biotechnol.14, 875–879. 18. Vasil, I. K. and Anderson, O. D. (1997) Genetic engineering of wheat gluten. Trends Plant Sci. 2, 292–297. 19. He, G. Y., Jones, H. D., D’Ovidio, R., Masci, S., Chen, M. J., West, J., Butow, B., Anderson, O. D., Lazzeri, P., Fido, R. and Shewry, P. R. (2005) Expression of an extended HMW subunit in transgenic wheat and the effect on dough mixing properties. J. Cereal Sci. 42, 225–231. 20. Barro, F., Rooke, L., Bekes, F., Gras, P., Tatham, A. S., Fido, R., Lazzeri, P. A., Shewry, P. R. and Barcelo, P. (1997) Transformation of wheat with high molecular weight subunit genes results in improved functional properties. Nat. Biotechnol. 15, 1295–1299. 21. Rooke, L., Barro, F., Tatham, A. S., Fido, R., Steele, S., Bekes, F., Gras, P., Martin, A., Lazzeri, P. A., Shewry, P. R. and Barcelo, P. (1999) Altered functional properties of tritordeum by transformation with HMW glutenin subunit genes. Theor. Appl. Genet. 99, 851–858. 22. Rooke, L., Bekes, F., Fido, R., Barro, F., Gras, P., Tatham, A. S., Barcelo, P., Lazzeri, P. and Shewry, P. R. (1999) Overexpression of a gluten protein in transgenic wheat results in greatly increased dough strength. J. Cereal Sci. 30, 115–120. 23. Alvarez, M. L., Guelman, S., Halford, N. G., Lustig, S., Reggiardo, M. I., Ryabushkina, N., Shewry, P. R., Stein, J. and Vallejos, R. H. (2000) Silencing of HMW glutenins in transgenic wheat expressing extra HMW subunits. Theor. Appl. Genet. 100, 319– 327. 24. He, G. Y., D’Ovidio, R., Anderson, O. D., Fido, R., Tatham, A. S., Jones, H. D., Lazzeri, P. and Shewry, P. R. (2000) Modification of storage protein composition in transgenic bread wheat, in Wheat Gluten,

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ing for phosphinothricin (PPT) – resistant transgenic wheats. Mol. Breed. 5, 255–262. 49. Lamacchia, C., Shewry, P. R., Di Fonzo, N., Forsyth, J. L., Harris, N., Lazzeri, P. A., Napier, J. A., Halford, N. G. and Barcelo, P. (2001) Endosperm-specific activity of a storage protein gene promoter in transgenic wheat seed. J. Exp. Bot. 52, 243–250. 50. Jones, H. D., Doherty, A. and Wu, H. (2005) Review of methodologies and a protocol for the Agrobacterium-mediated transformation of wheat. Trends Plant Sci. 1, 5. 51. Amoah, B. K., Wu, H., Sparks, C. and Jones, H. D. (2001) Factors influencing Agrobacterium-mediated transient expression of uidA in wheat inflorescence tissue. J. Exp. Bot. 52, 1135–1142.

Chapter 5 Agrobacterium-Mediated Transformation of Bread and Durum Wheat Using Freshly Isolated Immature Embryos Huixia Wu, Angela Doherty, and Huw D. Jones Abstract Agrobacterium-mediated transformation of wheat is becoming a viable alternative to the more established biolistic protocols. It offers advantages in terms of simple, low-copy-number integrations and can be applied with similar efficiencies to specific durum wheat and spring and winter bread wheat types varieties. Key words: Agrobacterium tumefaciens, bread wheat, durum wheat, AGL1, inoculation, immature embryo.

1. Introduction Agrobacterium tumefaciens is the only known prokaryote genus whose members have the innate ability to transfer genes to plants, although it has been recently demonstrated that this ability can be acquired by other bacterial genera after modification with a Ti plasmid (1). During the late 1980s, several dicot species were transformed using A. tumefaciens, but at that time it was thought that cereals were somehow recalcitrant to infection by A. tumefaciens and that physical method of DNA transfer was the only route to genetic engineering (see Chapter 1). However, the perceived benefits of Agrobacterium-mediated transformation, particularly in terms of simple and low-copy-number T-DNA integrations, as seen in dicot transformants, were a major research driver, and

Huw D. Jones and Peter R. Shewry (eds.), Methods in Molecular Biology, Transgenic Wheat, Barley and Oats, vol. 478 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-379-0_5

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demonstrations of heritable transgene integrations mediated by Agrobacterium were eventually published for rice in 1994, for maize in 1996, and for spring wheat and barley in 1997. All the major cereals with the exception of oat and millet have now been reportedly transformed using Agrobacterium. Relatively few wheat genotypes have been transformed using Agrobacterium and almost all are spring types (reviewed in (2)). The most commonly used explants are immature embryos, although embryogenic callus has also been used successfully. The protocol that follows here uses freshly isolated, immature embryos of between 0.8 and 1.5 mm long which are co-cultivated with the Agrobacterium strain AGL1 (3) housing a pGreen-based binary and a pSoup-based helper plasmid containing additional vir genes (4,–6). The procedure takes approximately 13 weeks from inoculation to potting into soil and has been successfully used for spring and winter bread wheat varieties as well as one durum wheat variety (2, 4, 7). In our laboratory, transformation efficiencies for this protocol, calculated as the proportion of PCR positive transgenic plants to immature embryos used, ranged between 0.3 and 9%.

2. Materials 2.1. Plant Donor Material Supply

1. For winter bread wheat varieties, vernalization is carried out from seed at 4 5°C. Light is supplied by 70 W fluorescent lamps for 12 h every day giving approximately 150 µ mol/ m2/s down at the level of shelf surface. After 8 weeks, seedlings are transferred to a controlled growth room with conditions as detailed below. 2. For spring varieties, seeds are directly sown into 21 cm diameter pots with Rothamsted mix compost detailed in (8). All the pots are placed and maintained in a controlled environment. Day/night temperature is set at 18–20°C/14–15°C, and the room humidity is kept at 50–70%. Light is supplied by 400 W quartz iodine lamps for 16 h leaving 8 h of darkness each day, and the intensity is 700 µ mol/m2/s photosynthetically active radiation at the level of pot surface. To begin with, the plants are top-watered manually, and once the roots have established the pots are supplied by an automated flooding system. In order to protect plants from powdery mildew and other fungal diseases, preventative spray is applied once at around 4–6 weeks after sowing. 3. Growth condition for durum wheat is as follows (see Note 1). After 3 weeks of vernalization at 4°C from seed

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(vernalization conditions are the same as for bread wheat), seedlings are moved to glasshouse compartments (Envirocon) with automatic blinds, irrigation, and side and top vents. Day and night temperatures are maintained at 20–25°C day/16–18°C night with 16/8-h photoperiod. Manual watering is applied throughout, and preventative spray is carried out at 4 5 weeks after the seedlings have been moved from vernalization. 2.2. Standard Glycerol Setting for A. tumefaciens Cells

1. Use streak Agrobacterium cells on LB medium, made solid with 15 g/l Bacto agar (Table 1), and supplemented with the appropriate antibiotics. Grow at 28°C in the dark for 2–3 days until single colonies are easily visible (see Note 2). 2. Pick a single colony using a cocktail stick or pipette tip. Place into 10 ml of liquid LB medium (Table 1), with appropriate antibiotics, in a 50-ml disposable centrifuge tube. Grow at 28°C in the dark with vigorous shaking (250 rpm) until the suspension is turbid, reaching an optical density (OD) >1.0 (A600). 3. Spin the Agrobacterium culture at 4,500 × g for 5 min. Resuspend in 1 ml of sterilized 10 mM MgSO4, then mix in 3 ml of 80% sterilized glycerol. Aliquot in 400 µl volumes in sterile cryotubes for long-term storage at −80°C (see Note 3). 4. For short-term storage, grow a single colony as described in 3 above, but in 3.5 ml of liquid in a 15 ml disposable centrifuge. Aliquot in 400 µl volumes, but for each cryotube take 200 µl of the culture and mix with 200 µl of 30% sterilized glycerol. These can be stored at −20°C for 1–2 months or at −80°C for longer-term storage.

2.3. Media Used in the Protocol

LB medium (9): Bacto-tryptone 10 g/l, bacto-yeast extract 5 g/l, sodium chloride 10 g/l to volume with double distilled water. Adjust to pH 7.0 if necessary. Autoclave. MG/L medium (10): Mannitol 5 g/l, L-glutamic acid 1 g/l, potassium dihydrogen orthophosphate 250 mg/l, sodium chloride 100 mg/l, magnesium sulfate (7H2O) 100 mg/l, tryptone 5 g/l, yeast extract 2.5 g/l. Adjust to pH 7.0 if necessary. Autoclave. Add biotin at 1 µ g/l after autoclaving.

2.3.1. Plant Tissue Culture Media

All the plant tissue culture media used in this protocol are listed in Table 1. It includes inoculation medium/co-cultivation medium (liquid/solid), induction mediuma for bread wheat, induction mediumb for durum wheat, first selection1 and second selection2 media.

2.4. Preparation of Media

All the plant media are prepared from stock solutions at double strength to allow the addition of an equal volume of gelling agents

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Table 1 Composition of double-strength culture media (All concentrations are shown double strength except for the supplements added after pH adjustment and sterilization which are shown at their final concentrations.) Inoculation/co- Inductiona cultivation (l−1) (l−1)

Inductionb (l−1)

Regeneration (l−1)

Selection1 Selection2 (l−1) (l−1)

MS Macro salts (×10) (ml)

200

200

200

–

–

–

L7 Macro salts (×10) (ml)

–

–

–

200

200

200

L7 Micro salts (×1,000) (ml)

2

2

2

2

2

2

FeNaEDTA (×100) (ml)

20

20

20

20

20

20

MS vitamins (×1,000) (ml)

2

2

2

–

–

–

Vitamins/inositol – (×200) (ml)

–

–

10

10

10

Inositol (mg)

200

200

200

200

200

200

Glutamine (mg)

1,000

1,000

750

–

–

–

Proline (mg)

–

–

150

–

–

–

Asparagine (mg)

–

–

100

–

–

–

Casein hydrolysate (mg)

200

200

–

–

–

–

MES (g)

3.9

3.9

–

–

–

–

Glucose (g)

20

–

–

–

–

-

Maltose (g)

80

80

–

60

60

60

Sucrose (g)

–

–

90

–

–

–

Component

pH adjusted to 5.7 then filter sterilized 2,4-D (mg)

2

0.5

–

0.1

–

–

Picloram (mg)

2.0

2.0

-

-

-

-

Acetosyringone (µ M)

200

–

–

–

–

–

Timentin (mg)

–

160

160

160

160

160

Zeatin (mg)

–

–

–

5

–

–

PPT (mg)

–

–

–

–

2.5

3.5

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which are also at double concentration. The double strength of phytagel is 4 g/l for solid inoculation (co-cultivation) medium, and induction mediuma and mediumb. For the regeneration, selection1, and selection2 media, agargel (10 g/l) is used instead of phytagel. The liquid inoculation medium for re-suspending Agrobacterium cells in Section 3.1 is used as single strength. Therefore, add the same volume of sterile distilled water to the doublestrength inoculation medium. 2.5. Stock Basal Medium Composition (Adapted from (8))

MS macrosalts (×10): 16.5 g/l of NH4NO3, 19.0 g/l of KNO3, 1.7 g/l of KH2PO4, 3.7 g/l of MgSO4·7H2O, 4.4 g/l of CaCl2·2H2O. Dissolve each component in distilled water separately before mixing. Autoclave at 121°C for 20 min and store at 4°C. L7 macrosalts (×10): 2.5 g/l of NH4NO3, 15.0 g/l of KNO3, 2.0 g/l of KH2PO4, 3.5 g/l of MgSO4·7H2O, 4.5 g/l of CaCl2·2H2O. Dissolve each component in distilled water separately before mixing. Autoclave at 121°C for 20 min and store at 4°C. L7 microsalts (×1,000): 15.0 g/l of MnSO4, 5.0 g/l of H3BO3, 7.5 g/l of ZnSO4·7H2O, 0.75 g/l of KI, 0.25 g/l of Na2MoO4·2H2O, 0.025 g/l of CuSO4·5H2O, 0.025 g/l of CoCl2·6H2O. Prepare 100 ml of microsalt stock solution at a time. Filter sterilize and store at 4°C. MS vitamins (glycine) (×1,000): 0.1 g/l of thiamine HCl, 0.5 g/l of pyridoxine HCl, 0.5 g/l of nicotinic acid. Prepare 100 ml at a time. Filter sterilize and store at 4°C. Vitamins/inositol (×200): 40.0 g/l of myo-inositol, 2.0 g/l of thiamine HCl, 0.2 g/l of pyridoxine HCl, 0.2 g/l of nicotinic acid, 0.2 g/l of Ca-pantothenate, 0.2 g/l of ascorbic acid. Filter sterilize and store at −20°C in 10 ml aliquots.

2.6. Supplements

Acetosyringone (3′,5′-dimethoxy-4′-hydroxyacetophenone): Dissolve in 70% ethanol to give 10 mg/ml or 50 mM stock solution. Filter sterilize, aliquot, and store at −20°C. 2,4-Dichlorophenoxyacetic acid (2,4-D): 1 mg/ml in ethanol/ water (dissolve powder in ethanol then add water to volume). Filter sterilize and store at −20°C in 1 ml aliquots. Zeatin mixed isomers (10 mg/ml): Dissolve powder in a small volume of 1 M HCl and make up to volume with water, and mix well/vortex. Filter sterilize and store at −20°C in 1 ml aliquots. Picloram (1 mg/ml): Dissolve picloram in water, filter sterilize, and store at −20°C in 2 ml aliquots.

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Timentin (300 mg/ml): Dissolve timentin (Ticarcillin/Clavulanic (15:1)) in water. Filter sterilize and store at −20°C in 1 ml aliquots. Glufosinate ammonium (PPT) (10 mg/ml): Dissolve in water, and mix well/vortex. Filter sterilize and store at −20°C in 1 ml aliquots. Silwet L-77 (1% v/v): Dissolve in water, filter sterilize, and store at 4°C in 0.5 ml aliquots. Biotin (1 mg/100 ml): Dissolve in water, filter sterilize, and store at −20°C in 0.5 ml aliquots (add 100 µl to 1 l MG/L).

3. Methods 3.1. Preparation of A. tumefaciens Cells for Inoculation

1. Take a 400 µl of glycerol stock aliquot, one per 10 ml MG/L liquid medium, and grow overnight (17 20 h) in a 50-ml sterile disposal tube with the appropriate antibiotics at 28°C on an orbital shaker (250 rpm) (17 20 h), until an OD 1.0 1.5 approximately (A600) is reached. 2. Pellet Agrobacterium cells by centrifugation at 4,500 × g for 10 min, dispose of the supernatant (see Note 4), and resuspend the cells in 4 ml of 1× inoculation medium (Table 1). 3. Put the 50-ml tube with 4 ml of inoculation medium plus Agrobacterium cells back in the shaker for 1–3 h at 28°C, in the dark (see Note 5).

3.2. Preparation of Explants for Inoculation with A. tumefaciens (see Note 6)

1. Collect ears of both bread wheat and durum wheat at 12–16 days post anthesis and surface-sterilize immature seeds by soaking for 1 min in 70% (v/v) ethanol followed by 10 min in 10% (v/v) bleach with gentle shaking. At least three rinses should be given with sufficient sterilized distilled water (see Note 7). 2. Isolate immature embryos from the seeds under a stereomicroscope in a laminar flow hood (see Note 8). Use a sharp scalpel to remove the axis. 3. Place the embryos scutellum side up onto the inoculation medium (Table 1) in 55- or 60-mm Petri dishes. This is for the scutellum to have maximum contact with Agrobacterium cells, see Section 3.3, step 2. Isolate about 50 immature embryos per plate, making sure the embryos are in good contact with medium to prevent them from floating when the Agrobacterium is added.

3.3. Inoculation of the Immature Embryos

1. When the plate is ready, take a tube from the shaker (see Note 9) and add 60 µl of 1% (v/v) Silwet to make a final concentration of 0.015%.

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Fig. 1. (a) Pouring the Agrobacterium slowly from the edge of the Petri dish to prevent the embryos from floating. (b) Illustration of the removal of the Agrobacterium–tilt plate slightly and with care not to disturb the embryos and remove as much of the liquid as possible. (c) An indication of what the plate will look like after 3 days co-cultivation. (d) Closeup of the embryos to show the level of Agrobacterium growth. d1. One would expect good embryogenesis from these embryos. d2. Overgrowth of Agrobacterium will damage the embryos and prevent good embryogenesis.

2. Immediately after step 3 in Section 3.2, pour all the Agrobacterium cells (4 ml) from one tube onto the plate (see Fig. 1a, Note 10). 3. Leave the plates in the dark for 15 min to 3 h (see Note 11). Other plates can be prepared for inoculation with Agrobacterium cells during this time. 4. Pipette off Agrobacterium suspension (see Fig. 1b). Care should be taken not to pipette off any embryos (see Note 12). Transfer the embryos onto a fresh Petri dish with solid inoculation medium. Make sure the scutellum side is up and the axis side is in contact with medium. 5. Co-cultivate plates at 22–23°C in a culture room in the dark for 2–3 days.

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3.4. Induction of Embryogenic Calli and Regeneration

1. After 2–3 days of co-cultivation, transfer all embryos to induction medium (Table 1, inductiona for bread wheat and inductionb for durum wheat) for embryogenesis (see Fig. 1c and d). At this stage, a record of the number of embryos transferred should be kept, as it is this number that will be used for calculating transformation efficiency. After the transfer, make sure the embryos are still scutellum side up. Keep them in the dark at 22–23°C. 2. After 3–4 weeks, transfer the embryogenic calli to regeneration medium (Table 1) for a further 3–4 weeks and put in the light with a photoperiod of 12 h/12 h for day/night (see Note 13).

3.5. Selection (see Note 14)

1. Transfer the cultures to the first round of selection medium (Table 1, selection1) (see Note 15). Some of the calli break up naturally when handing: this is not a problem, but do place the pieces near each other and mark them in some way so that it is known that they originated from the same embryo. 2. After a further 3 or 4 weeks select calli with shoots and roots and transfer them to a second round of selection (Table 1, selection2). At this stage they are big enough to go into a Magenta box. Continue to keep those originating from the same callus together marking as above.

3.6. Identification of Transgenic Plants and Obtaining Transgenic Seeds

1. Choose plantlets with good roots and shoots (see Note 16) at the end of the second round of selection, and transfer them to soil. Initially, pot up individual plants into 8-cm square plant pots and place under a propagator for a week or two so that they can acclimatize to the green house conditions. When they are big enough, leaf material can be collected, and DNA can then be extracted for PCR and Southern blot analysis (see Note 17). 2. For the winter varieties, the Magenta boxes containing the putative transgenic plants should be placed in a vernalization room (for condition see Section 2.1) for 6–8 weeks. Then transfer them to soil as above. 3. Confirmed transgenic plants should each be transferred to 13cm diameter pots to grow to maturity and set seed.

3.7. Calculation of Transformation Efficiency

1. The transformation efficiency is usually given as a percentage. This is calculated by dividing 100 by the total number of embryos inoculated and multiplying by the total number of confirmed independent transgenic plants. 2. When more than one confirmed transgenic plant is produced from one embryo, they are treated as just one independent event. All the other plants from this event are treated as

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sister plants. There is a possibility that some of these plants have derived from different cells with in the same callus and are therefore independent events but this can only be confirmed by Southern analysis. Therefore it is possible that the calculated efficiency is an underestimation for some experiments.

4. Notes 1. Most durum wheat varieties can be treated as spring wheat without the need for vernalization, but the temperature range for optimal durum wheat growth in CE or glasshouse is different from bread wheat and should be applied accordingly. 2. The antibiotics used depend on the selectable makers in the Agrobacterium strain and binary vectors used. For the AGL1 strain used in this protocol, carbenicillin (200 mg/l) is used and pAL154/pAL156 or pAL155/pAL156 combinations are selected with kanamycin (100 mg/l), which is the selectable marker on pAL156 (2, 7). 3. Ideally, the glycerol stocks should be put into liquid nitrogen immediately prior to the storage at −80°C. 4. The supernatant should be reasonably clear and the pellet should look creamy; if not, your culture might be a bit too old. 5. The 3-h incubation time is not a critical maximum; it could be left up to 8 h. Transgenic plants have been produced with inoculums incubated for longer than 3 h but the Agrobacterium does get clumpy. 6. Always prepare two control plates. A dry control is a plate of embryos treated in exactly the same way but with no inoculation step. Also a wet control is treated in exactly the same way, but the 1× inoculation medium plus silwet without Agrobacterium cells is used as the inoculum. These two controls are used to monitor the quality of the immature embryos, and the liquid and solid media. 7. Immature seeds can be detached from the ears and stored at 4°C overnight. Care should be taken to prevent the seeds from drying out. After surface sterilization, the seeds should not be stored for more than a few hours. The embryos should be 0.8–1.5 mm in length and translucent to white in appearance. If they are of a creamy colour, they are too old.

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8. The tube should be in the shaker for at least 30 min after resuspension but before use. 9. Most of the embryos will stay attached to the medium while submerged in the Agrobacterium suspension. If some emb-ryos float, use a little force to submerge them, as the scutellum side needs to be in contact with Agrobacterium cells. 10. The plates can be covered with a blue roll or foil to keep them dark. The time range may also seem extreme. Fifteen minutes is, in theory, enough to get T-DNA transfer but this may not fit in with the way the experiment is being carried out. It is therefore possible to extend the inoculation stage to timings that are more amenable, if necessary, but be aware that embryos will be damaged by long-term wetness, and too much Agrobacterium growth is also damaging (see Note 12). 11. Do not to leave embryos too wet; take as much Agrobacterium suspension as possibly before transfer, but do not use filter paper to dry the embryos. This is a crucial step and it is worth taking time over, as too much Agrobacterium growth during the co-cultivation stage will damage the embryos. 12. Some severely damaged embryos without any sign of growth during the induction period can be removed at this stage, but a record should be taken. 13. PPT used as the binary vector in this protocol has the bar gene as the selectable marker. 14. As the majority of embryogenic calli give rise to shoots and some roots, it is important that the roots are pushed fully into the selection medium. This will lower the risk of escapes (i.e. plantlets forming that are not transgenic). Double-bottom Petri dishes (one bottom being used as the lid) can be used at this stage to allow more room for the shoots to form properly. 15. If the first leaf is not healthy, it is unlikely that the plantlet is tolerant to selection pressure. Usually, if a plantlet has developed strong roots deep in the selection medium, it is an indication that it may be transgenic. 16. If the plantlets are big and strong enough at this stage, leaf samples can be taken for DNA extraction and PCR reaction, enabling an early confirmation of transgenic plants. If the binary vector has gusA gene, a segment of leaf can be taken for a GUS activity assay (4, 11).

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Acknowledgments Rothamsted Research receives grant-aided support from the Biotechnology and Biological Sciences Research Council (BBSRC) of the UK. CIMMYT receives grant-aided support for wheat genetic transformation from the European Commission, the Japanese Government, and the Molecular Plant Breeding Cooperative Research Center.

References 1. Broothaerts, W., Mitchell, H. J., Weir, B., Kaines, S., Smith, L. M. A., Yang, W., Mayer, J. E., Roa-Rodriguez, C. and Jefferson, R. A.(2005) Gene transfer to plants by diverse species of bacteria. Nature 433, 629–633. 2. Jones, H. D., Doherty, A. and Wu, H. (2005) Review of methodologies and a protocol for the Agrobacterium-mediated transformation of wheat. Plant Methods 1, 5. 3. Lazo, G. R., Stein, P. A. and Ludwig, R. A. (1991) A DNA Transformation-competent Arabidopsis genomic library in Agrobacterium. Bio-Technology 9, 963967. 4. Wu, H., Sparks, C., Amoah, B. and Jones, H. D.(2003) Factors influencing successful Agrobacterium-mediated genetic transformation of wheat. Plant Cell Rep. 21, 659– 668. 5. Hellens, R. P., Edwards, E. A., Leyland, N. R., Bean, S. and Mullineaux, P. M.(2000)pGreen: a versatile and flexible binary Ti vector for Agrobacterium-mediated plant transformation. Plant Mol. Biol. 42, 819– 832. 6. Amoah, B. K., Wu, H., Sparks, C. and Jones, H. D.(2001)Factors influencing Agrobacterium-mediated transient expression of uidA

7.

8.

9.

10.

11.

in wheat inflorescence tissue. J. Exp. Bot. 52, 1135–1142. Wu, H., Doherty, A. and Jones, H. D. (2008) Efficient and rapid Agrobacteriummediated transformation of durum wheat (Triticum turgidum L. ssp durum) using additional virulence genes. Transgen. Res. 17, 425–436. Sparks, C. A. and Jones, H. D.(2004) Transformation of wheat by biolistics, in Transgenic Crops of the World Essential Protocols, (Curtis, I. P., ed.), Kluwer, Dordrecht, pp. 19–35. Sambrook, J., Fritsch, E. F. and Maniatis, T. (1989) Molecular Cloning: A Laboratory Manual, 2nd ed., Cold Spring Harbor Laboratory, New York. Tingay, S., McElroy, D., Kalla, R., Fieg, S., Wang, M. B., Thornton, S. and Brettell, R. (1997) Agrobacterium tumefaciens-mediated barley transformation. Plant J. 11, 1369–1376. Wu, H., Sparks, C. A. and Jones, H. D. (2006) Characterisation of T-DNA loci and vector backbone sequences in transgenic wheat produced by Agrobacterium-mediated transformation. Mol. Breed. 18, 195–208.

Chapter 6 Floral Transformation of Wheat Sujata Agarwal, Star Loar, Camille Steber, and Janice Zale Abstract A method is described for the floral transformation of wheat using a protocol similar to the floral dip of Arabidopsis. This method does not employ tissue culture of dissected embryos, but instead pre-anthesis spikes with clipped florets at the early, mid to late uninucleate microspore stage are dipped in Agrobacterium infiltration media harboring a vector carrying anthocyanin reporters and the NPTII selectable marker. T1 seeds are examined for color changes induced in the embryo by the anthocyanin reporters. Putatively transformed seeds are germinated and the seedlings are screened for the presence of the NPTII gene based on resistance to paromomycin spray and assayed with NPTII ELISAs. Genomic DNA of putative transformants is digested and analyzed on Southern blots for copy number to determine whether the T-DNA has integrated into the nucleus and to show the number of insertions. The nonoptimized transformation efficiencies range from 0.3 to 0.6% (number of transformants/number of florets dipped) but the efficiencies are higher in terms of the number of transformants produced/number of seeds set ranging from 0.9 to 10%. Research is underway to maximize seed set and optimize the protocol by testing different Agrobacterium strains, visual reporters, vectors, and surfactants. Key words: Pre-anthesis wheat spikes, Agrobacterium, nuclear transformation, floral dip.

1. Introduction Early reports of Agrobacterium-mediated gene transfer to the floral tissue of hexaploid wheat (Triticum aestivum L.) set the stage for the merry-go-round of hopeful and discouraging reports of germ-line transformation in wheat. Seventeen years ago floral transformation of wheat was reported, but the transformation events were apparently not stable and were rearranged or lost in subsequent generations (1). Later it was reported that Agrobacterium-mediated germ-line transformation experiments in wheat, barley, and maize produced artifacts on Southern Huw D. Jones and Peter R. Shewry (eds.), Methods in Molecular Biology, Transgenic Wheat, Barley and Oats, vol. 478 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-379-0_6

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blots in the T1 generation when the transgenes were analyzed and these ‘artifacts’ were not stably transmitted to the next generation (2). They were thought to be due to bacterial contamination and/or transformation of endophytic bacteria (2). The target for these early attempts was purportedly the pollen (1, 2), and rudimentary Agrobacterium infiltration medium was applied into cut florets at or near anthesis. Subsequently, particle bombardment protocols of dissected embryos or embryogenic calluses were devised (3–5) that did not risk bacterial contamination. Wheat transformation is both an art and a science, and developing a protocol that consistently generates transformants is challenging. An additional complication is that there are technical difficulties in working with a large genome species. The hexaploid wheat genome is 17,000 Mbp per haploid nucleus (6), and molecular manipulations in this crop are not trivial. The choice of selectable markers and reporters may also determine success, as many screens (e.g. herbicide resistance) can be influenced by the environment. In addition, selecting for a transformation event in monocotyledons is not as clear-cut as in dicotyledonous plants. We are currently optimizing floral transformation of wheat to generate a protocol that consistently produces transformants (see Note 1).

2. Materials 2.1. Plant Materials

1. Crocus or Chinese Spring wheat seeds are available from the USDA-Germplasm Resources Information Network (GRIN) facility (http://www.ars-grin.gov/npgs/). Crocus is a highquality hard red Canadian spring germplasm line (7) and Chinese Spring is commonly used in genetic studies (see Note 2).

2.2. Growth Facilities

1. A controlled temperature greenhouse or walk-in growth chamber. 2. Standard potting mix such as Fafard #2 (Knoxville Seed and Greenhouse). 3. Osmocote 14-14-14 (Hummert International). 4. Six inch plastic pots.

2.3. Agrobacterium Growth

1. A glycerol stock, derived from a single colony of AGL1 or C58C1 Agrobacterium transformed with pBECKSred (8, 9) by a standard electroporation protocol (see Note 3). 2. LB broth (Fisher Biotech) with 50 µg/ml of spectinomycin and 50 µg/ml of rifampicin.

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3. LB agar plates (LB broth with 12 g/l of bacterial agar (Fisher)) with 50 µg/ml of spectinomycin and 50 µg/ml of rifampicin. 4. YEP broth (20 g/l of peptone, 10 g/l of yeast) with 50 µg/ml of spectinomycin, 50 µg/ml of rifampicin, and 200 µM of acetosyringone (3′,5′-dimethoxy-4′-hydroxyacetophenone). The acetosyringone is dissolved in a couple of drops of dimethyl sulfoxide (DMSO). 2.4. Infiltration Medium and Agrobacterium Treatment

1. Half-strength Murashige and Skoog (MS) (10) medium (Fisher Biotech) pH 5.8, 5% w/v sucrose, 0.04% v/v Silwet L-77, 0.5 mM of 2-(N-morpholino)ethanesulfonic acid (MES), and 200 µM of acetosyringone (see Note 4) (11, 12). 2. Small plastic bottles that will hold 250 ml of liquid and can be autoclaved afterwards. 3. Small clear, plastic bags and glassine pollination bags. 4. Small scissors and tweezers.

2.5. Screening

1. Dissecting stereoscope. 2. Commercial bleach (30%); optiona l- cefotaxime 1,000 ppm (Fisher) (13). 3. A 2% (w/v) of paromomycin spray (Fisher) mixed with 0.2% (v/v) of Tween 20 to act as a surfactant (14). 4. NPTII ELISA (Enzyme Linked ImmunoSorbent Assay) kit (Agdia, Inc, Indiana) and a microplate reader at 540 nm wavelength.

2.6. Copy Number Southern Gel Blots to Show Integration of the T-DNA

1. See Chapter 13.

3. Methods 3.1. Growth of the Wheat Plants

1. The temperature of the growth facility should never exceed 25°C. Ideal daytime temperatures are 21–25°C with a nighttime temperature of 16°C. A light cycle of 16 h is desirable. 2. Six-inch pots are filled with Fafart #2 potting medium and mixed with a standard analysis fertilizer such as Osmocote 1414-14. The wheat seeds are planted one per pot. Additional nutrients are applied if required. 3. Light quality and/or quantity has not been monitored but never drops below 400–500 µE. 4. Water the plants well before floral transformation and then do not water until at least 2 days after the Agrobacterium treatment.

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3.2. Growth of Agrobacterium and Induction of the Vir Genes

1. The glycerol stock of Agrobacterium harboring the plasmid is used to inoculate 5 ml of LB broth with the appropriate antibiotics and grown overnight at 22 ± 4°C with shaking at 2 × g.

3.3. Infiltration Media

The YEP/Agrobacterium culture is centrifuged at 6,400 × g for 15 min at room temperature and the supernatant is discarded. All reagents except the Silwet L-77 are mixed and used to gently re-suspend the cells to a final cell density of OD600 = 1.0. The Silwet L-77 is added to the infiltration medium just prior to treating the plant.

3.4. Preparation of the Wheat Spikes and Agrobacterium Treatment

1. Pre-anthesis wheat spikes at the early, mid to late uninucleate microspore stage yield transformants. This is the early boot stage when the spike, still enclosed in the sheath, is 6–7 cm long (Fig. 1, see Note 5).

2. The next day, 2.5 ml of this culture is used to inoculate 250 ml of YEP broth containing the antibiotics and 200 µM of acetosyringone, and grown at 22 ± 4°C with shaking at 2 × g until the optical density (OD600) = 1.0.

2. The sheath is opened and the developing spike is carefully removed. The terminal florets and the inner florets can be removed, as they are frequently sterile, or they can be left

Fig. 1. The early boot stages of wheat. The early uninucleate stage (left) and the mid to late uninucleate stage (right) are the correct stages for floral transformation. The spike, still enclosed in the sheath, is gently removed from the sheath and the florets are clipped before treatment with Agrobacterium infiltration medium. Floral transformation at an earlier stage will produce fewer seeds.

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intact. The remaining florets are clipped to slightly below the end of the glumes. 3. Shake the Agrobacterium infiltration medium before immersing the spike in it, and keep the spike immersed for 1–2 min (see Note 6). 4. Cover the spike in small, clear plastic bags for 2 days to ensure high humidity. 5. Remove the plastic bags and allow the spikes to recover from the stress. Allow the spikes to dry after the Agrobacterium treatment, and as they near anthesis, cover them in glassine bags to prevent cross pollination and let the T1 seeds set naturally (see Note 7). 3.5. Screening

A series of facile screens are conducted consecutively before performing the time-consuming and labor-intensive Southern blots for determining copy number, and reverse transcriptase polymerase chain reaction (RT-PCR) or northern blots for studying gene expression.

3.6. Visual Inspection of the Seed

1. The TI seeds are examined for red embryos and compared with several wild-type seeds under a stereoscope (Fig. 2, see Note 8). 2. Putatively transformed seeds are washed for 30 min with shaking in 30% commercial bleach solution to kill residual Agrobacterium, and rinsed three times with water at 5 min per wash before planting in potting mix.

Fig. 2. Seeds of a putative transformant (left) versus wild type (right). This putative transformant was produced after treatment with Agrobacterium harbouring pBECKSred which carries Lc/C1, and the embryo/endosperm is a deeper red than wild type.

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3. Optionally, a cefotaxime treatment (1,000 ppm; in addition to step 1 – Section 3.6) can also be applied to germinating seedlings for 2 h at room temperature to kill residual Agrobacterium (13). 3.7. Screening for Resistance to 2% Paromomycin at the Whole Plant Level

1. Putative transformed seedlings and several wild-type control plants are sprayed at the 3–4 leaf stage with the 2% (w/v) paromomycin spray containing 0.2% (v/v) Tween 20 to act as a surfactant (14). 2. The plants are scored as resistant or sensitive 5–7 days later (see Note 9). Save the resistant plants as putative transformants.

3.8. Screening with NPTII ELISAs

1. Follow the protocol for the NPTII ELISA (Agdia Inc.). 2. It is important to solubilize the NPTII standards provided with the kit in the protein extracts of the wild-type control plants. 3. Include more than one wild-type control plant, as one may produce a false positive (see Note 10). 4. Samples of healthy plant tissue of the putative transformants are selected and extracted in microfuge tubes at 4°C to produce concentrated protein samples (1:5 tissue weight:buffer volume) using the Agdia extraction buffer. A 1-ml pipette tip works well to macerate the tissue in place of a pestle. The samples are centrifuged for 5–10 min at 7,500 × g and the supernatant is saved. 5. If possible, triplicate wells for each sample and standard should be assayed in the ELISA. 6. At the completion of the assay, stop the reaction with 3 M H2SO4 and use a microplate reader at 450 nm to quantify the results. Select the plants with ELISA readings higher than those of wild type as putative transformants.

3.9. Copy Number Southern Blots to Show Integration of the T-DNA

Genomic DNA of putative transformants, wild-type controls, and plasmid is extracted, digested to completion, fractionated according to size in agarose gel electrophoresis, blotted on to a membrane, and hybridized with a probe complementary to the gene of interest. See Chapter 13.

4. Notes 1. Patent Application “In Planta Transformation of Cereals” U.S. Serial No. 11/112,393, filed April 22, 2005 – Patent Pending. 2. Crocus is subject to post harvest grain dormancy. To speed up germination, incubate the seeds on moistened filter

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paper at 4°C. Crocus and Chinese Spring both carry the double recessive alleles for high crossability with rye (7, 15). It is not yet known if this transformation method is genotype dependent. 3. pBECKSred carries the NPTII selectable marker driven by the nos promoter, and the Lc/CI anthocyanin regulators can act as cell autonomous indicators of a transformation event by turning the embryo tissue red (8, 16). While several transformants have been produced with this vector, the pCambia vectors are preferred since they are small, stable in Agrobacterium, and have been sequenced (17). In addition to C1, B-Peru allele is another useful reporter derived from a maize transcription factor that regulates the anthocyanin pathway (18). The 35S:adhI intron:B-Peru and the 35S:adh1 intron:C1 reporters have recently been cloned into pCambia2200 and are available on request. pCambia2200 also contains the NPTII selectable marker with the double enhancer version of the 35S promoter for better gene expression. The NPTII genes in pBECKSred and pCambia do not have introns, therefore, it is possible that the 35S promoter and the nos promoter maintain a low level of functionality in Agrobacteria (19). The pCambia vectors also contain a multiple cloning site for a gene of interest. Both C58C1 and AGL1 Agrobacterium strains have been used to produce wheat transformants. 4. Transformants have been produced with acetosyringone concentrations ranging from 200 µM to 1 mM with no additional detriment to seed set. 5. Spike development varies in different environmental conditions. If in doubt about the stage of microspore development, anthers can be fixed in formaldehyde:acetic acid: alcohol (1:1:3) for 24 h and then stored in 70% ethanol. Before examination, stain for 1 h in 1% acetocarmine stain and examine under a light microscope (20). Florets in the middle of the spike mature sooner than the terminal florets. Agrobacterium treatment before the early uninucleate microspore stage decreases seed set considerably and after the late uninucleate stage decreases the chance of producing transformants. The spike is fragile at this stage and care must be taken not to break it off. 6. In some environments (e.g. increased heat stress), it may be necessary to decrease the dipping time and/or decrease the Silwet L-77 concentration in order to maximize seed set. Seed set at these recommendations is generally 22–23% of a fully fertile spike. 7. Out-crossing rates for some spring wheat cultivars can be relatively high (21), and it is necessary to prevent cross

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pollination. However, covering the spikes in cellophane or glassine bags before the spike has dried after the Agrobacterium treatment will increase humidity and fungal growth. 8. Putative T1 transformants will have red embryos if transformed with a construct containing Lc and/or C1. It should be kept in mind that severely stressed hard red wheat seeds may also turn a darker red; therefore, one must determine whether the color change in the embryo of the T1 seed is caused by the stress of the Agrobacterium treatment or a transformation event. Additional controls in these experiments could be included, in which wheat spikes are treated with infiltration media alone or with Agrobacterium infiltration media without the Silwet, although these controls have been less than helpful in distinguishing true transformation events. 9. Paromomycin is a geneticin G418 analog and useful as a spray at the whole plant level (14). Resistance to paromomycin indicates the presence of the NPTII gene. If the NPTII gene is present, minimal bleaching and flecking will occur in comparison to wild-type control plants. The strength of the promoter must be considered when scoring the plants; transformed plants carrying the NPTII gene driven by a weak promoter (e.g. nos) may bleach minimally in comparison to wild type. 10. False positives may be due to cellular debris in the extract that have not been removed by centrifugation. 11. The anthocyanin transcriptional regulators are toxic to white wheat (9). T1 seeds may be produced and germinate, but may die before anthesis (J. Zale, unpublished observations). 12. The female gametophyte appears to be the recipient of the T-DNA in Arabidopsis (22). At present, it is not known whether the target of T-DNA transfer is the female or male gametophyte in wheat. 13. PCR is never conducted on putative T1 transformants because it will produce false positives due to Agrobacterium contamination.

Acknowledgements This material is based upon work supported by the National Science Foundation under Grant No. 0638421 and the USDA NRI Grant No. 2001-01856.

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References 1. Hess, D., Dressler, K. and Nimmrichter, R. (1990) Transformation experiments by pipetting Agrobacterium into the spikelets of wheat (Triticum aestivum L), Plant Sci. 72, 233–244. 2. Langridge, P., Brettschneider, R., Lazzeri, P. and Lorz, H. (1992) Transformation of cereals via Agrobacterium and the pollen pathway: a critical assessment, Plant J. 2, 631–638. 3. Vasil, V., Castillo, A., Fromm, M. and Vasil, I. K. (1992) Herbicide resistant fertile transgenic wheat plants obtained by micro-projectile bombardment of regenerable embryogenic callus, Bio/Technology 1, 667–674. 4. Weeks, J. T., Anderson, O. D. and Blechl, A. E. (1993) Rapid production of multiple independent lines of fertile transgenic wheat (Triticum aestivum L.), Plant Physiol. 102, 1077–1084. 5. Nehra, N., Chibbar, R., Leung, N., Caswell, K., Mallard, C., Steinhauer, L., Baga, M. and Kartha, K. (1994) Self-fertile transgenic wheat plants regenerated from isolated scutellar tissues following microprojectile bombardment with two distinct gene constructs, Plant J. 5, 285–297. 6. Bennett, M. D. and Leitch, I. J. (1996) Nuclear DNA amounts in Angiosperms, Ann. Bot. 76, 113–176. 7. Zale, J. and Scoles, G. (1999) Registration of Crocus hard red spring wheat, Crop Sci. 39, 1539–1540. 8. McCormac, A. C., Elliott, M. C. and Chen, D. F. (1997) pBECKS. A flexible series of binary vectors for Agrobacterium-mediated plant transformation, Mol. Biotechnol. 8, 199–213. 9. McCormac, A. C., Wu, H., Bao, M., Wang, Y. Q., Xu, R., Elliot, M. C. and Chen, D. F. (1997) The use of visual marker genes as cell-specific reporters of Agrobacteriummediated T-DNA delivery to wheat and barley, Euphytica 99, 17–25. 10. Murashige, T. and Skoog, F. A. (1962) A revised medium for rapid growth and bioassays with tobacco tissue culture, Physiol. Plant. 15, 473–497. 11. Bechtold, N., Ellis, J. and Pelletier, G. (1993) In planta Agrobacterium mediated gene transfer by infiltration of adult Arabidopsis thaliana plants, C. R. Acad. Sci. 316, 1194–1199.

12. Clough, S. J. and Bent, A. F. (1998) Floral dip: a simplified method for Agrobacteriummediated transformation of Arabidopsis thaliana, Plant J. 16, 735–743. 13. Supartana, P., Shimizu, T., Nogawa, M., Shioiri, H., Nakajima, T., Haramoto, N., Nozue, M. and Kojima, M. (2006) Development of simple and efficient in planta transformation method for wheat (Triticum aestivum L.) using Agrobacterium tumefaciens, J. Biosci. Bioeng. 102, 162–170. 14. Cheng, M., Fry, J. E., Pang, S., Zhou, H., Hironaka, C. M., Duncan, D. R., Conner, T. W. and Wan, Y. (1997) Genetic transformation of wheat mediated by Agrobacterium tumefaciens, Plant Physiol. 115, 971–980. 15. Lamoureux, D., Boeuf, C., Regad, F., Garsmeur, O., Charmet, G., Sourdille, P., Lagoda, P. and Bernard, M. (2002) Comparative mapping of the wheat 5B short chromosome arm distal region with rice, relative to a crossability locus, Theor. Appl. Genet. 105, 759–765. 16. McCormac, A. C., Elliott, M. C. and Chen, D. F. (1999) pBECKS2000: a novel plasmid series for the facile creation of complex binary vectors, which incorporates “cleangene” facilities, Mol. Gen. Genet. 261, 226– 235. 17. Cambia (2007) Center for the Application of Molecular Biology to International Agriculture in Canberra, Australia, June 15, 2007 ( http://www.cambia.org/daisy/cambia/585). 18. Selinger, D. A. and Chandler, V. L. (2001) B-Bolivia, an allele of the maize b1 gene with variable expression, contains a high copy retrotransposon-related sequence immediately upstream, Plant Physiol. 125, 1363–1379. 19. Jacob, D., Lewin, A., Meister, B. and Appel, B. (2002) Plant-specific promoter sequences carry elements that are recognised by the eubacterial transcription machinery, Transgen. Res. 11, 291–303. 20. Rybczynski, J. J., Simonson, R. I. and Baenziger, P. S. (1991) Evidence for microspore embryogenesis in wheat anther culture, In Vitro Cell. Dev. Biol. 27, 168–174. 21. Hucl, P. and Matus-Cadiz, M. (2001) Isolation distances for minimizing out-crossing in spring wheat, Crop Sci. 41, 1348–1351. 22. Bent, A. F. (2000) Arabidopsis in planta transformation. Uses, mechanisms, and prospects for transformation of other species, Plant Physiol. 124, 1540–1547.

Chapter 7 Highly Efficient Agrobacterium-Mediated Transformation of Wheat Via In Planta Inoculation Thierry Risacher, Melanie Craze, Sarah Bowden, Wyatt Paul and Tina Barsby Abstract This chapter details a reproducible method for the transformation of spring wheat using Agrobacterium tumefaciens via the direct inoculation of bacteria into immature seeds in planta as described in patent WO 00/63398 (1). Transformation efficiencies from 1 to 30% have been obtained and average efficiencies of at least 5% are routinely achieved. Regenerated plants are phenotypically normal with 30–50% of transformation events carrying introduced genes at single insertion sites, a higher rate than is typically reported for transgenic plants produced using biolistic transformation methods. Key words: Triticum aestivum, Agrobacterium tumefaciens, transformation, immature embryos, in planta inoculation.

1. Introduction Agrobacterium-mediated transformation of wheat has been the goal of many research groups since routine systems were developed for other less readily transformed crop species. Direct DNA delivery methods have enabled the routine production of transgenic wheat (1– 4). However, the high insert number associated with this method is often seen as a disadvantage to breeders, regulatory agencies, and those using this technology to validate gene function. Several groups now have established methods for the transformation of wheat and other monocotyledons by Agrobacterium tumefaciens, with varying transformation efficiencies reported (5) (see also Chapters 5,6 and 9 in this volume). Huw D. Jones and Peter R. Shewry (eds.), Methods in Molecular Biology, Transgenic Wheat, Barley and Oats, vol. 478 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-379-0_7

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The method outlined below (6) differs from other published wheat transformation methods in that the Agrobacterium is delivered to the immature embryo while it is still in the developing seed. This circumvents the need for many of the complicated and technically exacting delivery techniques required by other methods, such as the use of wetting agents and osmotic conditioning (7–9). The method is quick, and easy to undertake practically; experiments are initiated rapidly without the need for in vitro co-cultivation steps. The example given is for a spring wheat line (referred to herein as NB1), transformed with the A. tumefaciens supervirulent strain EHA105 (10), containing a binary superclean vector (SCV) (11) with the rice actin promoter (12), driving β-glucuronidase (13), and the Sc4 subterranean clover stunt virus promoter (14) linked to the selectable marker NptII (neomycin phosphotransferase II) (15).

2. Materials 2.1. Stock Plant Growth

1. Stock plants are produced in 12,7F (13 cm diameter) pots, four plants per pot. 2. Plants are grown in Levingtons M2 compost with 5 g/l added Osmocote Exact slow release fertilizer (15:9:9 + 3MgO + TE; 5to 6-month release); a thin layer of medium vermiculite is scattered on top of the compost. 3. Plants are supported with Teku plant supporters (LBS Horticulture, Colne, Lancashire, UK). 4. Trays are lined with Aquamat (LBS Horticulture), six pots per tray (30 cm × 50 cm base, deep gravel tray). 5. Watering is regulated with Octamitters. 6. Growth chamber used is a Conviron PGV/PGW 36.

2.2. Agrobacterium Mini-Glycerol Production

The following materials are required: 1. LB medium (Sigma-Aldrich) 2. Universal tubes (25 ml, Sterilin) 3. LB medium/glycerol (Sigma-Aldrich) 85:15 solution

2.3. Agrobacterium Preparation and Inoculation

The following materials are used: 1. YEP medium: 5 g/l yeast extract, 10 g/l bacto-peptone, 5 g/l NaCl, 15 g/l bacto-agar, pH 6.8 (all from Sigma-Aldrich) 2. TSIM medium: 4.4 g/l MS salts and vitamins (MP Biomedicals), 100 mg/l myo-inositol, 10 g/l glucose, 500 mg/l MES buffer (all from Sigma-Aldrich), pH 5.2, filter sterilized

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Fig. 1. Syringe and dispenser.

3. Acetosyringone (3,5-diphenoxy-4-hydroxyacetophenone) (Sigma-Aldrich), 0.1 M aqueous solution 4. Hamilton repeat syringe dispenser 25 µl to 2.5 ml (see Fig. 1) 5. Hamilton syringes (50 µl) with needles 6. Measuring cylinder (500 ml) 7. Glass rod or similar support 8. Transparent plastic bag (455 × 600 mm) 9. Sanyo Versatile Environmental Test Chamber 2.4. Tissue Culture

The following materials are required: 1. Domestos bleach (Lever Bros) 2. Ethanol solution (70% v/v): made up with de-ionized water 3. Sterile de-ionized water 4. Scalpel with no. 15 blades (Swann-Morton, Sheffield, UK) 5. W4 medium: 4.4 g/l MS salts and vitamins (MP Biomedicals), 20 g/l sucrose (Merck KGaA, Germany), 2 mg/l 2,4 dichlorophenoxyacetic acid, sodium salt (Sigma-Aldrich), 500 mg/l glutamine (Sigma-Aldrich), 100 mg/l casein enzymatic hydrolysate (Sigma-Aldrich), 6 g/l Type 1-A agarose (SigmaAldrich), 150 mg/l Timentin (ticarcillin disodium/potassium clavulanate) (Melford Laboratories, Ipswich, UK), pH 5.8. Table 1 outlines preparation details for media additives 6. W425G medium: as for W4 but with the addition of 25 mg/l G418, disulfate salt (Sigma-Aldrich) 7. MRM medium: 4.4 g/l MS salts and vitamins (MP Biomedicals), 20 g/l sucrose (Merck KGaA, Germany), 6 g/l Type 1A agarose (Sigma-Aldrich), 2 mg/l kinetin (Sigma-Aldrich), pH 5.8 8. MRM25G medium: as for MRM but with the addition of 25 mg/l G418, disulfate salt (Sigma-Aldrich) 9. Beatson jars (250 ml; Richardsons, Leicester, UK) with Petri dish base as lid and sealed with 25 mm 3M MicroporeTM tape

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Table 1 Preparation of media additives Tissue culture additive

Solvent/diluent

Filter sterilize or co-autoclave

Stock concentration (mg/ml)

Final concentration (mg/l)

2,4-D

1 M NaOH/H2O

Co-autoclave

1

2

Kinetin

1 M NaOH/H2O

Filter sterilizea

1

2

Timentin

H2O

Filter sterilize

100

150

G418

H2O

Filter sterilize

10

25

Stocks of tissue culture additives should be made up regularly and stored at −20°C a Filter sterilized after required amount of kinetin is made up in 100 ml of water/litre media and the pH altered to 5.8, added to volume-adjusted autoclaved media

10. MS20 medium: 4.4 g/l MS salts and vitamins (as above), 20 g/l sucrose (Merck KGaA, Germany), 7 g/l plant cell culture tested agar (Sigma-Aldrich), pH 5.8 11. Sanyo Versatile Environmental Test Chamber 2.5. Growth of Transgenic Plantlets

The following materials are required: 1. Jiffy 7 peat pellets (Jiffy Products, Norway) 2. Seed trays and 24-well tray inserts (any horticultural supplier) 3. Propagators to fit seed trays (any horticultural supplier)

2.6. Growth of Transgenic Plants

The following materials are required: 1. 12F (12 cm diameter) plant pots, one plant per pot 2. Levingtons M2 compost plus slow release fertilizer (as above) 3. Pea sticks/split canes for support 4. Micro perforated bread bags (Focus Packaging) for bagging

3. Methods 3.1. Stock Plant Growth

1. Sow seeds (four per pot) at a depth of 2–3 cm in M2 compost with slow release fertilizer, as above. 2. Scatter a thin layer of vermiculite on top of the pot to maintain the quality of the compost surface, and attach a Teku plant supporter to each pot. 3. Place pots in a Conviron at 20°C day/15°C night with 16-h photoperiod, 400 µE/m2/s (see Note 1).

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1. Harvest tillers at approximately 16–18 days post anthesis (dpa) when embryos have reached approximately 1 mm in size. Cut tillers close to the base of the plant (see Note 2). 2. Remove and discard lower leaves, keeping the flag leaf attached. 3. Put the tillers in water as soon as possible in a 500-ml cylinder, trimming tiller length so that the flag leaf is positioned just above the top of the cylinder.

3.3. De-husking

1. Remove the lower two or three spikelets from each ear to be prepared. 2. Carefully remove the glumes and lemmas from florets 1 and 2 (see Fig. 2) all the way up the ear from bottom to top, to expose the immature seeds in those two florets only. 3. Cut the remaining tissue that interferes with inoculation, i.e. parts of lemma/glume and awns, if present, on untouched florets (3, 4, etc.) with scissors. Take care not to cut into any uncovered seed, as this can lead to fungal contamination during co-cultivation. 4. Brush the ear upwards with a soft brush to remove anthers, another potential source of fungal growth. 5. Wipe leaves with wet tissue. 6. Lightly spray the ears with 70% v/v ethanol. Allow to dry before inoculation (see Note 3).

3.4. Agrobacterium Plates

1. Prepare mini-glycerol stocks of Agrobacterium containing plasmid. 2. Take a single colony from the master plate and set up an overnight culture in LB medium (16) with appropriate selection at 28°C with vigorous shaking. Transfer 1 ml to an Eppendorf tube and centrifuge at 10,000 × g for 30 s. Remove supernatant and re-suspend cells in 1 ml LB with 15% (w/v) glycerol. Aliquot into smaller volumes and maintain at −80°C. Use one mini-glycerol stock to inoculate each plate and discard after use. 3. Streak YEP plate containing appropriate selective antibiotics with Agrobacterium from the mini-glycerol stock prior to the experiment to form a lawn of Agrobacterium over the entire surface of the plate. 4. Seal the plate and incubate for 1 day at 28°C or 3 days at room temperature.

3.5. Agrobacterium Suspension

1. When the ears are ready, prepare the Agrobacterium suspension. 2. Wipe and flame a large flat-ended spatula and let it cool down thoroughly.

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Fig. 2. Anatomy of wheat spikelet. Image reproduced from WHEAT: THE BIG PICTURE (http://www.wheatbp.net).

3. Collect the bacteria by lightly scraping the surface of the plate with the spatula until a pellet is amassed, which can then be transferred into a 25-ml Universal tube containing 10 ml of TSIM and 400 µM of acetosyringone. Avoid transferring any agar from the plate, as this will interfere with the inoculation process. 4. Re-suspend the bacteria by repeated pipetting with a 1-ml Gilson pipette until a smooth suspension is achieved. 3.6. Inoculation

1. Sterilize a Hamilton syringe four or five times with 70% (v/v) ethanol and rinse four or five times with sterile water. Ensure the needle aperture is facing towards you. 2. Fill the syringe with the Agrobacterium suspension. 3. Hold the ear so that the embryo region of the seed is accessible to the needle. Push the needle into the ear so that the tip is roughly positioned at the interface of the endosperm and

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scutellum (Fig. 3). Minor damage of the scutellum is not a problem and provides evidence of correct inoculation. 4. Inject 1 µl by slowly depressing the button once. When a 50µl syringe is inserted, one depression of the button will deliver 1 µl of suspension. 5. Repeat the procedure for all the exposed seeds. 6. After all ears are inoculated, place a support into the cylinder (e.g. glass rod) and cover with a transparent plastic bag sealed at the base. Incubate at 22°C, 16-h photoperiod, 40 µE/m2/s for 2 or 3 days (see Note 4). 7. Clean the syringe by washing four or five times with 70% (v/ v) ethanol and rinse four or five times with sterile water. Rinse one more time with 70% ethanol and leave to dry. 3.7. Isolation

1. Two or three days after inoculation remove the seeds from the ears and place them into a suitable lidded container. 2. Surface sterilize the seeds for 1 min in 70% (v/v) ethanol followed by 25 min in 20% (v/v) Domestos solution, with agitation. 3. Wash thoroughly in a sieve with sterile distilled water. 4. Isolate the embryos and place onto W4 media with scutellum uppermost, 25 per plate (see Note 5). 5. Seal plates with Parafilm and incubate for 5 days at 28°C, 80 µE/m2/s, 16-h photoperiod.

3.8. Tissue Culture

Fig. 3. Site of inoculation.

1. Five days after isolation, remove the embryonic axes (see Note 6) from the scutella and transfer the scutella to fresh W4 medium (see Note 7).

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2. Twelve days after isolation, transfer the calli to selective medium (W4 25G). Incubate for 2 weeks at 25°C, 80 µE/ m2/s, 16-h photoperiod, (all subsequent transfers to be maintained at this temperature and light level). 3. After 2 weeks on selection, cut the calli into smaller pieces to improve contact with the selective agent and place on fresh W4 25G medium. 4. After a further 2 weeks on selection, transfer any actively growing embryogenic callus to regeneration medium containing selection agent (MRM25G). Repeat the transfer after 2 weeks, transferring any callus that has not already developed into shoots onto fresh MRM25G. 5. Transfer regenerating shoots to Beatson jars containing MS20 and place at 25°C (see Note 8). 6. Leave shoots to grow for approximately 1 month during which time leaves and roots should become well developed. 3.9. Transfer to Soil and Acclimatization

1. Prepare Jiffy 7 pellets by rehydrating in tap water and placing in 24-well tray inserts covered with a propagator.

2. Remove shoots from the Beatson jar, carefully avoiding transfer of agar with the roots. 3. Divide the mass of shoots until it appears that no further subdivision is possible i.e. each shoot is distinct. 4. Transfer shoots to Jiffy 7 pellets, ensuring all roots are encompassed by the Jiffy and that they are in close contact with the peat. Remove dead/dying leaves. Trim growing leaves back to 5–10 cm. Spray lightly with water before replacing propagator (see Note 9). 5. After about 2 weeks at 20°C the plants are ready for polymerase chain reaction (PCR) analysis, if required, and for potting (see Note 10). 6. For maximum seed production, transfer plantlets to 12F pots with M2 compost (as above), one plant per pot. The Jiffy is left intact at transfer to soil, the mesh surrounding the pellet not being removed. 7. Bag flowering plants to reduce pollen movement.

4. Notes 1. The quality of the stock plant material is critical. Biological control is used to contain most insect pests where needed.

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2. Primary and secondary tillers give similar results in terms of transformation efficiency. 3. Once ears have been prepared so that the seed has been exposed, do not leave tillers for any length of time before inoculation without first covering loosely with a plastic bag to prevent shrivelling of the seed. 4. Take care to ensure flag leaves are evenly spaced within the bag so that they achieve maximum light input during cocultivation. 5. Any visible Agrobacterium growth around the embryo at isolation will be controlled by transfer to media containing timentin. 6. Embryonic axes are removed from all embryos whether they have developed or not. 7. Transfer to fresh medium following removal of axes after only 4 or 5 days has a beneficial effect on callus production. 8. Pieces of callus cut from one initial inoculated embryo will often regenerate into multiple lines with several independent integration patterns. As a result, only one shoot regenerated from a callus line should be transferred to soil, unless Southern analysis is used to identify independent transformation events. Obviously, this facet could be exploited to increase the transformation efficiency. 9. After transfer to Jiffy 7 pellets, shoots are sprayed with water before placing in incubators to promote a humid environment. After 3 or 4 days, hardening off can begin by opening incubator vents; incubator lids can be removed after 1 week. 10. Plants that have been hardened off grow extremely quickly in Jiffy 7 pellets and need to be potted within a week or two of removing the incubator. For short-term use, e.g. plants for PCR analysis only, plants can be kept in Jiffys without potting as long as they are fertilized regularly (MiracleGro, manufacturer’s recommended concentration).

References 1. Christou, P. (1992). Genetic transformation of crop plants using microprojectile bombardment. Plant J. 2, 275–281. 2. Taylor, N. J. and Fauquet, C. M. (2002) Microparticle bombardment as a tool in plant science and agricultural biotechnology. DNA Cell Biol. 21, 963–977. 3. Rasco-Gaunt, S., Riley, A., Cannell, M., Barcelo, P. and Lazzeri, P. A. (2001)Procedures allowing the transformation of a range of

European elite wheat (Triticum aestivum L.) varieties via particle bombardment. J. Exp. Bot.52, 865–874. 4. Barsby, T., Power, J. B., Freeman, J., Ingram, H. M., Livesey, N. L., Risacher, T. and Davey, M. R. (2001) Transformation of wheat, in The World Wheat Book – A History of Wheat Breeding. Lavoisier Publishing, France and USA, and Intercept Ltd., UK, pp. 1081–1103.

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5. Bhalla, P. L., Ottenhof, H. H. and Singh, M. B. (2006). Wheat transformation – an update of recent progress. Euphytica 149, 353–366. 6. Risacher, T. and Craze, M. (1992) WO 00/63398. Plant Transformation Method. 7. Cheng, M., Fry, J. E., Pang, S., Zhou, H., Hironaka, C. M., Duncan, D. R., Conner, T. W. and Wan, Y. (1997) Genetic transformation of wheat mediated by Agrobacterium tumefaciens. Plant Physiol. 115, 971–980. 8. Cheng, M., Hu, T. C., Layton, J., Liu, C. N. and Fry, J. E. (2003) Desiccation of plant tissues post-Agrobacterium infection enhances T-DNA delivery and increases stable transformation efficiency in wheat. In Vitro Cell Dev. Biol. 39, 595–604. 9. Uze, M., Wunn, J., Puonti-Kaerlas, J., Potrykus, I. and Sautter, C. (1997) Plasmolysis of precultured immature embryos improves Agrobacterium-mediated gene transfer to rice (Oryza sativa L.). Plant Sci. 130, 87–95. 10. Hood, E. E., Gelvin, S. B., Melchers, L. S. and Hoekema, A. (1993) New Agrobacterium vectors for plant transformation. Transgen. Res. 2, 208–218.

11. Firek, S., Ozcan, S., Warner, S. A. and Draper, J. (1993) A wound-induced promoter driving npt-II expression limited to dedifferentiated cells at wound sites is sufficient to allow selection of transgenic shoots. Plant Mol. Biol. 22, 129–42. 12. McElroy, D., Zhang, W., Cao, J. and Wu, R. (1990) Isolation of an efficient actin promoter for use in rice transformation. Plant Cell, 2, 163–171. 13. Jefferson, R. A. (1987) Assaying chimeric genes in plants: the GUS gene fusion system. Plant Mol. Biol. Rep. 5, 387–405. 14. Schünmann, P. H. D., Surin, B. and Waterhouse, P. M. (2003) A suite of novel promoters and terminators for plant biotechnology. II. The pPLEX series for use in monocots. Funct. Plant. Biol. 30, 453–460. 15. Herrera-Estrella, L., De Block, M., Messens, E., Hernalsteens, J.-P., Van Montagu, M. and Schell, J. (1983) Chimeric genes as dominant selectable markers in plant cells. EMBO J. 2, 987–995. 16. Sambrook, J., Fritsch, E. F. and Maniatis, T. (1989) Molecular Cloning: A Laboratory Manual. Cold Spring Harbor Laboratory, New York.

Chapter 8 Barley Transformation Using Biolistic Techniques Wendy A. Harwood and Mark A. Smedley Abstract Microprojectile bombardment or biolistic techniques have been widely used for cereal transformation. These methods rely on the acceleration of gold particles, coated with plasmid DNA, into plant cells as a method of directly introducing the DNA. The first report of the generation of fertile, transgenic barley plants used biolistic techniques. However, more recently Agrobacterium-mediated transformation has been adopted as the method of choice for most cereals including barley. Biolistic procedures are still important for some barley transformation applications and also provide transient test systems for the rapid checking of constructs. This chapter describes methods for the transformation of barley using biolistic procedures and also highlights the use of the technology in transient assays. Key words: Biolistics, particle gun, microprojectile bombardment, barley transformation, immature embryo, transgenic plants.

1. Introduction Fertile transformed barley plants were first reported in 1994 (1) and were derived using particle bombardment or biolistic methods. Since that time there have been many reports of the production of transgenic barley using this technology. The methodology uses a particle delivery system or ‘gene gun’ to deliver the DNA of interest that is coated onto metal particles, or microprojectiles, of approximately 1 µm in diameter. Gold is preferred for the microprojectiles and the most commonly used gene gun is the PDS 1000/He biolistic delivery system (BioRad). Different particle bombardment devices and methods for the preparation of DNA-coated gold particles have been examined for their effect

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on barley transformation (2). This study found the PDS 1000/ He device to be most effective in yielding transformed barley plants. In common with Agrobacterium-mediated techniques, the most popular barley target tissue for transformation using particle bombardment is immature embryos. However, other targets such as microspores (3) and shoot meristematic cultures (4) have also been successfully used. The spring genotype Golden Promise is the most responsive in tissue culture and therefore the most amenable to transformation. Other more recalcitrant barley cultivars have also been transformed using particle bombardment (5) although at lower frequencies. A comparison of particle bombardment and Agrobacterium-mediated techniques for barley transformation (6) found that particle bombardment yielded lower transformation efficiencies, transgenic lines with higher numbers of copies of the transgene(s), and greater incidences of transgene silencing. It is for these reasons that Agrobacterium-mediated methods are now preferred for most applications. However, for some applications, such as the delivery of multiple transgenes, particle bombardment might still be the method of choice (7). In this chapter we describe a method for the particle bombardment of immature embryos of the spring barley cultivar Golden Promise and the recovery of transgenic plants using the bar gene, conferring resistance to the glufosinate group of herbicides, as a selectable marker. The accompanying chapter 9 on Agrobacterium-mediated barley transformation describes an alternative selection system based on resistance to the antibiotic hygromycin.

2. Materials 2.1. Plant Material

Plants of the spring genotype Golden Promise are grown under controlled environment conditions to provide immature embryos for transformation. Plants are grown at 15°C day and 12°C night temperatures, 80% relative humidity, and with light levels of 500 µmol/m2/s at the mature plant canopy level provided by metal halide lamps (HQI) supplemented with tungsten bulbs. The compost mix contains a 2:2:1 mix of Levington M3 compost/ Perlite/Grit. The mix also contains the slow release fertilizer Osmocote at the manufacturer’s recommended concentration. For further details see Chapter 9 the chapter on Agrobacteriummediated barley transformation.

2.2. Plant Tissue Culture Media

Four different basic plant tissue culture media are used during the transformation and regeneration process: callus induction,

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osmotic treatment, transition, and regeneration media (see Note 1). During selection stages on callus induction medium, Bialaphos (Meiji Seika Ltd., Tokyo, Japan) is added at 5 mg/l. During selection on transition and regeneration media, Bialaphos is added at 1 mg/l. Unless otherwise stated, all media components are supplied by Sigma-Aldrich and all media and stocks made up using water from an Elga water purifier. 1. Callus induction: 4.3 g/l Murashige and Skoog plant salt base (Duchefa M0221), 30 g/l of maltose, 1.0 g/l of casein hydrolysate, 350 mg/l of myo-inositol, 690 mg/l of proline, 1.0 mg/l of thiamine HCl, 2.5 mg/l of dicamba (Sigma-Aldrich D5417), 3.5 g/l of phytagel. The medium is adjusted to pH 5.8 with NaOH. 2. Osmotic treatment: This medium is identical to the callus induction medium but contains, in addition, 0.4 M of mannitol (72 g/l). 3. Transition: 2.7 g/l Murashige and Skoog modified plant salt base (without NH4NO3) (Duchefa M0238), 20 g/l maltose, 165 mg/l NH4NO3, 750 mg/l glutamine, 100 mg/l myoinositol, 0.4 mg/l thiamine HCl, 1.25 mg/l CuSO4·5H2O, 2.5 mg/l 2,4-dichlorophenoxy acetic acid (2,4-D) (Duchefa), 0.1 mg/l 6-benzylaminopurine (BAP) (Duchefa), 3.5 g/l phytagel. The pH is adjusted to 5.8. 4. Regeneration: 2.7 g/l Murashige and Skoog modified plant salt base (without NH4NO3) (M0238 Duchefa), 20 g/l maltose, 165 mg/l NH4NO3, 750 mg/l glutamine, 100 mg/l myo-inositol, 0.4 mg/l thiamine HCl, 3.5 g/l phytagel. The pH is adjusted to 5.8. Media stocks: 5. Callus induction 100× vitamin stock: 100 mg/l thiamine HCl, 35 g/l myo-inositol and 69 g/l proline. This stock should be filter sterilized ready for use and stored at 4°C. 6. Transition and regeneration 100× vitamin stock: 40 mg/l thiamine HCl and 10 g/l myo-inositol. This stock should be filter sterilized ready for use and stored at 4°C. 7. Dicamba: 2.5 mg/ml stock made up in water, filter sterilized, divided into 1 ml aliquots, and stored frozen. 8. 2,4-D: 2.5 mg/ml stock made up in 100% ethanol and stored at −20°C. 9. BAP: 1 mg/ml stock made up in water with a few drops of 1 M NaOH and stored at −20°C. 10. CuSO4 stock: 125 mg of CuSO4·5H2O dissolved in a total volume of 100 ml of water, filter sterilized, and stored at 4°C. 11. Bialaphos stock: 5 mg/ml stock made up in water, filter sterilized, and stored at −20°C.

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2.3. Plasmids for Transformation

An example of a plasmid containing the bar gene under the control of a maize ubiquitin promoter that has been successfully used for particle-bombardment-mediated barley transformation is shown in Fig. 1 (8) This plasmid, pAL51 contains an additional maize ubiquitin promoter to drive a gene of interest, in this case a firefly luciferase gene. The pBract series of plasmids described in the chapter 9 onAgrobacterium-mediated barley transformation could also be introduced by particle bombardment if required.

2.4. Particle Gun

The most commonly used particle gun or biolistic device is the PDS 1000/He device supplied by Bio-Rad. In addition, the consumables listed below are required for the transformation of barley. All consumables are supplied by Bio-Rad. 1. Gold microcarriers (1.0 µm) (165-2263) 2. Macrocarriers (165-2335) 3. Stopping screens (165-2336) 4. Rupture discs (1,100 psi) (165-2329).

2.5. Coating Gold Particles with DNA

1. Spermidine solution (0.1 M) 2. CaCl2 (2.5 M) 3. Ethanol (100%)

Fig. 1 Plasmid pAL51 as an example of a plasmid suitable for particle bombardment of barley. The plasmid contains the bar gene for selection under the control of a maize ubiquitin (ubi1) promoter. A second ubi1 promoter drives the gene of interest, in this case a firefly luciferase gene that has been extensively used as a marker gene in barley transformation (8).

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3. Methods 3.1. Tissue Culture Media Preparation

1. Phytagel is prepared in advance at two times the required concentration and sterilized by autoclaving (see Note 2). 2. All tissue culture media are filter sterilized; therefore all media components, except those stored as sterile stocks, are added at two times the required concentration and dissolved, the pH adjusted, and the media filter sterilized (Steritop 0.22-µm filters, Millipore). 3. Both the 2× phytagel and the 2× media are warmed to 60°C in a water bath before use. 4. The required stocks are added under sterile conditions to the warm media, and the media and the phytagel mixed and poured into 9-cm Petri dishes (Sterilin). 5. For regeneration, media are poured into deeper dishes (tissue culture dish, 100 mm × 20 mm, Falcon). 6. For osmotic treatment, smaller 5-cm Petri dishes are used (Sterilin).

3.2. Isolation of Immature Embryos

1. Barley spikes are collected when the immature embryos are 1–2 mm in diameter (see Note 3).

3.2.1. Collection and Sterilization of Immature Seeds

2. Before cutting the spikes, a single immature seed from the middle of each spike is checked to make sure that the size of the immature embryos is correct (see Fig. 9.2a–c in chapter 9 on Agrobacterium-mediated barley transformation for further guidance). 3. The immature seeds are removed from the spike and the awns broken off without damaging the seed coat. 4. The immature seeds are then sterilized by first washing in 70% ethanol for 30 s followed by three washes in sterile distilled water. This is followed by sterilization in a solution of sodium hypochlorite (sodium hypochlorite solution, Fluka 71696) diluted 50:50 with water for 4 min. The sterilization is followed by four washes in sterile distilled water after which the immature seeds are drained but left wet in a screw-top sterile jar.

3.2.2. Isolation of Immature Embryos and Removal of Embryonic Axis

All operations are performed in a laminar flow hood under sterile conditions. 1. Sterile seeds, approximately 20 at a time, are tipped onto a sterile blue or black tile under a dissecting microscope. The seed is held firm with a pair of fine forceps, and a second pair of fine forceps is used to expose the immature embryo and remove the embryonic axis (see Note 4).

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2. Embryos are placed, 25–30 per plate, scutellum side up, on callus induction medium and cultured at 25°C in the dark. 3.3. Preparation of DNA-Coated Gold Particles 3.3.1. Preparation of Gold Stocks

The method used for the preparation of DNA-coated gold particles is similar to the method described in (9). 1. Gold (40 mg) is weighed into a 1.5-ml Eppendorf tube, and 1 ml of 100% ethanol added. 2. The gold is mixed well by vortexing and then pulsed in a microfuge to pellet. 3. The ethanol is removed and this procedure repeated another two times. This washes the gold prior to use. 4. After pelleting the gold for the last time and removing the ethanol, 1 ml of sterile, distilled water is added to the gold and it is mixed by vortexing to re-suspend. 5. The gold is then divided into 50 µl aliquots in sterile Eppendorf tubes. While dispensing the gold, care must be taken to ensure that the gold stays in suspension by frequent mixing. Gold stocks are stored at −20°C until required.

3.3.2. Preparation of Spermidine

1. Spermidine, purchased in 1 g amounts from Sigma-Aldrich, is prepared by placing the unopened bottle in a water bath at 65°C and allowing the powder to melt. 2. In a flow hood using sterile pipette tips, the spermidine is placed in 14 µl aliquots into sterile 1.5-ml Eppendorf tubes. The tubes are then stored at −20°C until required.

3.3.3. Coating of the Gold Particles

1. One Eppendorf tube containing 14 µl of spermidine is removed from the freezer and allowed to thaw. 2. Sterile water (986 µl) is added and the solution mixed by vortexing. 3. Using a 1-ml syringe and small syringe filter (Minisart 0.2 µm filter unit, Sartorius) the solution is filter sterilized into a sterile 1.5-ml Eppendorf tube. This procedure prepares a 0.1 M spermidine solution. 4. One Eppendorf tube containing 50 µl of gold stock is removed from the freezer and allowed to thaw. 5. To this gold stock, 5 µl of the required plasmid DNA, at a concentration of 1 µg/µl is added by pipetting the DNA onto the side of the Eppendorf and then vortexing immediately to mix the gold and DNA (see Note 5). 6. Then, 50 µl of 2.5 M CaCl2 and 20 µl of 0.1 M spermidine are mixed in the lid of the Eppendorf tube. The tube is then closed, upside down so that the CaCl2 and spermindine do not mix with the gold until the tube is turned, and immediately vortexed to mix all components thoroughly.

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7. The tube is then incubated on ice for 1 min before centrifugation in a microfuge at 1,957 × g for a maximum of 30 s to pellet the gold. 8. The supernatant is removed and 250 µl of 100% ethanol added to the gold pellet and mixed first by gentle pipetting up and down and then by vortexing. This procedure washes the coated gold particles before use. 9. The gold particles are then pelleted by centrifugation for a maximum of 30 s at 1,957 × g, the supernatant removed, and 60 µl of 100% ethanol added. 10. The gold is re-suspended by vortexing. The DNA-coated gold particles are now ready for use. 3.4. Particle Bombardment of Immature Embryos 3.4.1. Osmotic Treatment of Embryos

3.4.2. Particle Gun Preparation

1. One day after isolation and 4 h before particle bombardment, embryos are arranged on osmotic medium, 20–30 embryos per plate, scutellum side up, in a 1.6 cm2 area in the centre of each plate. The embryos can be arranged close together but not touching each other. Embryos are left on the osmotic treatment plates for 16 h after particle bombardment. 1. All components of the particle gun and the inside of the gun chamber are cleaned with 100% ethanol. Stopping screens and macrocarriers are dipped in ethanol to sterilize and left to dry. Rupture discs are dipped into isopropanol immediately before use and do not need additional sterilization. 2. Once macrocarriers are dry, they are placed into macrocarrier holders and 3.5 µl of the prepared DNA-coated gold particles are carefully pipetted into the centre of each macrocarrier. Care must be taken to mix the gold particles by vortexing frequently to ensure that they stay in suspension.

3.4.3. Particle Gun Parameters

1. The components of the particle gun are adjusted to give the following distances within the gun chamber: Rupture disc to macrocarrier: 2.2 cm Macrocarrier to stopping screen: 1.3 cm Stopping screen to sample: 5.8 cm (position 3 from the base of the chamber) Vacuum: 28 in.Hg.

3.4.4. Particle Bombardment

1. Before starting bombardment of the immature embryos, two blank bombardments are carried out with the particle gun (see Note 6). 2. Following this, a 1,100 psi rupture disc is dipped into isopropanol and inserted into the rupture disc holder, which is then tightened onto the base of the gas acceleration tube.

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3. A stopping screen is inserted into the microcarrier launch assembly together with a prepared macrocarrier with dried, DNA-coated gold particles. The macrocarrier is inserted with the gold particles facing down. The top of the microcarrier launch assembly is tightened, and the whole unit inserted into the chamber below the rupture disc holder. 4. The plate containing the immature embryos is placed on the target shelf that has been inserted into position 3 from the base of the chamber. The lid of the dish containing the embryos is removed and the door of the gun chamber closed. The vacuum within the chamber is taken down to 28 in.Hg and held at this level, and then the gun is fired. 5. Once the vacuum is released, the door is opened and the lid returned to the plate of embryos. The gun is then prepared for bombardment of the next plate. 6. Bombarded embryos are incubated at 25°C in the dark. 3.5. Selection of Transformed Material

1. Sixteen hours after bombardment, embryos are removed from the osmotic treatment plates and transferred to callus induction plates containing 5 mg/l Bialaphos for selection. Plates are incubated at 25°C in the dark. 2. Embryos are sub-cultured onto fresh callus induction plates with 5 mg/l Bialaphos every 2 weeks. 3. At the second sub-culture, after a total of 4 weeks on selection medium, the callus derived from each embryo is divided into 3–6 small pieces. The plate is marked so that all callus derived from one original immature embryo is kept together. 4. At the third sub-culture, after 6 weeks, all remaining healthy callus is transferred to transition medium containing 1 mg/l Bialaphos and placed at 25°C under low light. The low light is achieved by placing the plates under light conditions in a tissue culture room but covering the plates with a thin sheet of paper.

3.6. Regeneration of Transgenic Plants

1. After 2 weeks on transition medium, the embryo-derived material is transferred to regeneration medium, in deep Petri dishes, without any growth regulators but with 1 mg/l Bialaphos (Fig. 2). Sub-cultures are made every 2 weeks, onto the same medium, until no more regenerated plants are produced. 2. Plantlets with shoots of 2–3 cm in length and forming roots are carefully removed from the plates and transferred to glass culture tubes (Sigma C-5916) containing 12 ml of callus induction medium, without any growth regulators, but with 1 mg/l Bialaphos. 3. Once rooted plants reach the top of the tubes, they can be transferred to soil. Plants are gently removed from tubes using long forceps and all tissue culture medium washed from the roots under a running tap.

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Fig. 2. Selection of transformed barley on Bialaphos containing regeneration medium. The line at the top of the plate is transformed and regenerating on Bialaphos containing medium. The callus within each area outlined in black on the plate is derived from a single immature embryo.

4. They are then planted in the same barley growth mix described above in 5-cm diameter pots. Plants are covered with small individual plastic jars with holes at the top as individual propagators to maintain humidity until the plants are well established in soil. 5. Once the plants are established in soil, leaf samples can be collected for further analysis to confirm the presence of the introduced genes. Bialaphos selection does allow some nontransformed plants to survive unlike hygromycin selection, described in chapter 9 on Agrobacterium-mediated transformation. 6. A quick and easy test of the transformed status of the regenerated plants is the leaf test for herbicide resistance (8). 3.7. Using the Particle Gun for Transient Assays

One important use of particle bombardment is in transient assay systems, in particular to check constructs in advance of stable transformation experiments. In transient assays, plates of immature embryos are often bombarded twice to increase the amount of DNA delivered. It is advisable to turn the plate containing the immature embryos through 90° between the two bombardments (see Note 7). Examples of immature embryos bombarded with plasmids containing the gus gene and a gfp gene are shown in Fig. 3a and b, respectively.

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Fig. 3. Transient expression following particle bombardment. a) An immature embryo bombarded with a plasmid containing the gus gene under the control of a maize ubiquitin promoter. The embryo was stained to reveal GUS activity 2 days after bombardment. (b) An immature embryo bombarded with a plasmid containing a green florescent protein (gfp) gene under the control of a maize ubiquitin promoter. The embryo was examined 2 days after bombardment.

4. Notes 1. Early barley transformation using the particle gun was carried out using a tissue culture and regeneration protocol that did not us e a transition medium (2). This procedure yielded transgenic plants but at a lower frequency than that obtained with the introduction of the transition medium. Additional improvements to the tissue culture protocol have been included for Agrobacterium-mediated barley transformation (see chapter 9). 2. Phytagel can be prepared in advance at two times concentration in water, autoclaved and stored at room temperature because it will not solidify until the other culture medium components are added. 3. Slightly smaller immature embryos can be used for particle bombardment than for Agrobacterium-mediated transformation, as there is no problem of them being overgrown by Agrobacterium. However a size of 1.5 mm in diameter is still ideal. 4. There are several different techniques for removing the immature embryos and removing the embryonic axis. The most efficient way is to carry out this operation in one step holding the immature seed by puncturing it with one pair of fine forceps and then exposing the immature embryo by peeling back the seed coat from the awn end of the seed. Once the embryo is exposed, the embryonic axis can be removed by first pinching off the shoot axis with the fine forceps and then remov-

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ing the root axis with a second pinching action. This should leave only the undamaged scutellum. If preferred, it is possible to carry out this operation in two steps. First, the immature embryos can be removed and placed intact onto callus induction medium. Once the required numbers have been isolated, they can be picked up in groups of around ten together with a small amount of culture medium, placed on the sterile tile, and the axis removed from each using the fine forceps. They are then immediately plated scutellum side up on callus induction medium. 5. The DNA should not be allowed to mix with the gold before vortexing, as this can cause clumping of the gold particles. Similarly, if the spermidine and CaCl2 are allowed to mix with the gold without thorough vortexing, then clumping of the gold can result. If the concentration of DNA is too high, then again the gold particles will clump, so it is important to check the DNA concentration and to avoid adding too much. 6. The blank bombardments are carried out by inserting a rupture disc into the holder at the base of the gas acceleration tube and following the bombardment procedure without stopping screen, macrocarrier, or target tissue. This allows the system to be checked before starting bombardment of the immature embryos. 7. The area covered by a single bombardment can be increased by using an adaptor (Hepta adaptor, Bio-Rad). This adaptor allows seven macrocarriers, coated with gold microcarriers, to be used in a single shot, thereby covering a much larger area of the Petri dish. This is not really necessary for small targets such as immature embryos but can be useful for transient expression studies in larger target tissues.

References 1. Wan, Y. and Lemaux, P. G. (1994) Generation of large numbers of independent transformed fertile barley plants. Plant Physiol. 104, 37–48. 2. Harwood, W. A., Ross, S. M., Cilento, P. and Snape, J. W. (2000) The effect of DNA/gold particle preparation technique, and particle bombardment device, on the transformation of barley (Hordeum vulgare). Euphytica 111, 67–76. 3. Yao, Q. A., Simion, E., William, M., Krochko, J. and Kasha, K. J. (1997) Biolistic transformation of haploid isolated microspores of barley (Hordeum vulgare L.). Genome 40, 570–581.

4. Zhang, S., Cho, M.-J., Koprek, T., Yun, R., Bregitzer, P. and Lemaux, P. G. (1999) Genetic transformation of commercial cultivars of oat (Avena sativa L.) and barley (Hordeum vulgare L.) using in vitro shoot meristematic cultures derived from germinating seedlings. Plant Cell Rep. 18, 959–966. 5. Cho, M.-J., Jiang, W. and Lemaux, P. G. (1998) Transformation of recalcitrant barley cultivars through improvement of regenerability and decreased albinism. Plant Sci. 138, 229–244. 6. Travella, S., Ross, S. M., Harden, J., Everett, C., Snape, J. W. and Harwood, W. A. (2005) A comparison of transgenic barley

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lines produced by particle bombardment and Agrobacterium-mediated techniques. Plant Cell Rep. 23, 780–789. 7. Agrawal, P. K., Kohli, A., Twyman, R. M. and Christou, P. (2005) Transformation of plants with multiple cassettes generates simple transgene integration patterns and high expression levels. Mol. Breed. 16, 247–260.

8. Harwood, W. A., Ross, S. M., Bulley, S. M., Travella, S., Busch, B., Harden, J. and Snape, J. W. (2002) Use of the firefly luciferase gene in a barley (Hordeum vulgare) transformation system. Plant Cell Rep. 21, 320–326. 9. Becker, D., Brettschneider, R. and Lorz, H. (1994) Fertile transgenic wheat from microprojectile bombardment of scutellar tissue. Plant J. 5, 299–307.

Chapter 9 Barley Transformation Using Agrobacterium-Mediated Techniques Wendy A. Harwood, Joanne G. Bartlett, Silvia C. Alves, Matthew Perry, Mark A. Smedley, Nicola Leyland, and John W. Snape Abstract Methods for the transformation of barley using Agrobacterium-mediated techniques have been available for the past 10 years. Agrobacterium offers a number of advantages over biolistic-mediated techniques in terms of efficiency and the quality of the transformed plants produced. This chapter describes a simple system for the transformation of barley based on the infection of immature embryos with Agrobacterium tumefaciens followed by the selection of transgenic tissue on media containing the antibiotic hygromycin. The method can lead to the production of large numbers of fertile, independent transgenic lines. It is therefore ideal for studies of gene function in a cereal crop system. Key words: Barley transformation, Agrobacterium tumefaciens, transgenic plants, hygromycin, immature embryo.

1. Introduction The first reports of successful barley transformation (1) used biolistic-based techniques to introduce DNA to immature embryos. Immature embryos were also the target tissue used in the first reports of the generation of transgenic barley plants using Agrobacterium (2). Although alternative target tissues have been examined for use in barley transformation systems, immature embryos remain the target tissue of choice for obtaining high transformation efficiencies. An alternative Agrobacterium-mediated barley transformation system uses microspore cultures as the target tissue (3). A comparison of biolistic and Agrobacteriumbased methods for barley transformation highlighted some of Huw D. Jones and Peter R. Shewry (eds.), Methods in Molecular Biology, Transgenic Wheat, Barley and Oats, vol. 478 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-379-0_9

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the advantages of the Agrobacterium system (4). These included higher transformation efficiencies, lower transgene copy number, and more stable inheritance of the transgenes with less transgene silencing. Barley transformation is still very genotype dependent. The most responsive genotype is the spring cultivar Golden Promise. There have been reports of the transformation of other genotypes using Agrobacterium-mediated techniques but at lower frequencies (5). As well as the choice of target tissue and genotype, there are a number of other important variables in the Agrobacterium-mediated transformation system. These include Agrobacterium strain, co-cultivation conditions and timing, selection system, and plant regeneration system. For barley, Agrobacterium strains AGL1 or AGL0 are commonly used (6), but LBA 4404 has also been successfully used (7). In many reports of barley transformation, the bar gene conferring resistance to the glufosinate group of herbicides was used for selection together with Bialaphos or phosphinothricin (PPT) as the selective agent (4). This has now been largely replaced with the hpt gene conferring resistance to the antibiotic hygromycin as the selectable marker of choice. The hygromycin resistance gene can be driven by a CaMV 35s promoter for selection in barley, leaving stronger promoters such as the maize ubiquitin promoter (ubi1) available to drive a gene or cassette of interest. Many constructs have been developed that can be used for barley transformation. Here we describe the use of constructs from the pBract series (http://www.bract.org) which give high transformation frequencies and are simple to work with, as they are Gateway compatible for easy introduction of genes or sequences of interest. The method described below details the Agrobacteriummediated barley transformation procedure from growth of donor plants to provide immature embryos through to confirmation of the transgenic status of the regenerated plants.

2. Materials 2.1. Plant Material and Growth Conditions

1. Seed of the spring barley genotype Golden Promise is sown at two weekly intervals in a barley growth mix consisting of a 2:2:1 mix of Levington M3 compost/Perlite/Grit. The mix also contains the slow release fertilizer, Osmocote at the manufacturer’s recommended concentration. 2. Seed is initially sown in 5-cm diameter pots, and after approximately 30 days germinated plants are potted into 13-cm diameter round pots in the same growth mix for continued development.

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3. Plants are grown in a controlled environment room at 15°C day and 12°C night temperatures, 80% relative humidity, and with light levels of 500 µmol/m2/s at the mature plant canopy level provided by metal halide lamps (HQI) supplemented with tungsten bulbs (8) (see Note 1). 2.2. Agrobacterium Strains and Vectors

1. Agrobacterium strain AGL1 is used together with appropriate pBract vectors. In this example we use vector pBract 204 which contains the hpt gene conferring hygromycin resistance under a 35s promoter at the left border (LB) and a gus gene encoding β-glucuronidase under the control of the maize ubiquitin promoter at the right border (RB) (Fig. 1) (see Note 2). 2. The pBRACT vectors are based on pGreen, which is a small, versatile vector designed for easy manipulation in E. coli with a high copy number (9). To enable the small size of pGreen, the pSa origin of replication required for replication in Agrobacterium is separated into its two distinct functions. The replication origin (ori) is present on pGreen, and the trans-acting replicase gene (RepA) is present on an additional vector, named pSoup. Both vectors are required in Agrobacterium for pGreen to replicate.

2.3. Bacterial Culture Medium

The basic bacterial culture medium is MG/L (10), which contains 5.0 g/l tryptone, 5.0 g/l mannitol, 2.5 g/l yeast extract, 1.0 g/l L-glutamic acid, 250 mg/l KH2PO4, 100 mg/l NaCl,

Fig. 1. pBract 204, a pGreen-based plasmid containing the hpt (Hyg) gene conferring resistance to the antibiotic hygromycin under the control of a CaMV (cauliflower mosaic virus) 35s promoter at the left border and a β-glucuronidase (gus) gene under the control of a maize ubiquitin promoter (Ubi1) at the right border. (See Color Plates)

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100 mg/l MgSO4·7H2O, and 10 µl biotin (0.1 mg/l stock). The medium is adjusted to pH 7.2 with NaOH. For the preparation of plates, 15 g/l agar is added. 2.4. Plant Tissue Culture Media

Three different basic plant tissue culture media are used during the transformation and regeneration process: the callus induction, transition, and regeneration media. During all selection stages, hygromycin is added to the media at 50 mg/l (hygromycin B supplied as a sterile 50 mg/l stock from Roche). The antibiotic Ticarcillin with Clavulanic acid (Duchefa T0190) (otherwise known as Timentin) is also added at 160 mg/l to all selection stages. Additional copper (1.25 mg/l CuSO4·5H2O) is added during callus induction and transition stages. Unless otherwise stated, all media components are supplied by SigmaAldrich and all media and stocks made up using water from an Elga water purifier. 1. Callus induction: 4.3 g/l Murashige and Skoog plant salt base (Duchefa M0221), 30 g/l maltose, 1.0 g/l casein hydrolysate, 350 mg/l myo-inositol, 690 mg/l proline, 1.0 mg/l thiamine HCl, 2.5 mg/l Dicamba (Sigma-Aldrich D5417), 3.5 g/l Phytagel. The medium is adjusted to pH 5.8 with NaOH. 2. Transition: 2.7 g/l Murashige and Skoog modified plant salt base (without NH4NO3) (Duchefa M0238), 20 g/l maltose, 165 mg/l NH4NO3, 750 mg/l glutamine, 100 mg/l myo-inositol, 0.4 mg/l thiamine HCl, 2.5 mg/l 2,4-dichlorophenoxy acetic acid (2,4-D) (Duchefa), 0.1 mg/l 6-benzylaminopurine (BAP) (Duchefa), 3.5 g/l Phytagel. The pH is adjusted to 5.8. 3. Regeneration: 2.7 g/l Murashige and Skoog modified plant salt base (without NH4NO3) (M0238 Duchefa), 20 g/l maltose, 165 mg/l NH4NO3, 750 mg/l glutamine, 100 mg/l myo-inositol, 0.4 mg/l thiamine HCl, 3.5 g/l Phytagel. The pH is adjusted to 5.8. 4. Media stocks: (a) Callus induction 100 × vitamin stock: 100 mg/l thiamine HCl, 35 g/l myo-inositol, and 69 g/l proline. This stock should be filter sterilized ready for use and stored at 4°C. (b) Transition and regeneration 100× vitamin stock: 40 mg/l thiamine HCl and 10 g/l myo-inositol. This stock should be filter sterilized ready for use and stored at 4°C. (c) Dicamba: 2.5 mg/l stock made up in water, filter sterilized, divided into 1 ml aliquots, and stored frozen. (d) 2,4-D: 2.5 mg/ml stock made up in 100% ethanol and stored at −20°C.

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(e) BAP: 1 mg/ml stock made up in water with a few drops of 1 M NaOH and stored at −20°C. (f) CuSO4 stock: 125 mg of CuSO4·5H2O dissolved in a total volume of 100 ml water, filter sterilized, and stored at 4°C. (g) Hygromycin: purchased as a sterile 50 mg/ml stock, divided into 1 ml aliquots, and stored frozen. (h) Timentin: 160 mg/ml stock made up in water, divided into 1 ml aliquots, and stored frozen. 2.5. Other Equipment

1. Fine forceps No. 5 type (TAAB Laboratories Equipment Ltd. Ref. T083) 2. Binocular microscope (e.g. Leica MZ6).

3. Methods 3.1. Tissue Culture Media Preparation

Phytagel is prepared in advance at two times the required concentration and sterilized by autoclaving (see Note 3). All tissue culture media are filter sterilized, therefore all media components, except those stored as sterile stocks, are added at two times the required concentration, dissolved, the pH adjusted and the media filter sterilized (Steritop 0.22 µm filters, Millipore). Both the 2× phytagel and the 2× media are warmed to 60°C in a water bath before use. The required stocks are added under sterile conditions to the warm media, and the media and the phytagel mixed and poured into 9-cm Petri dishes (Sterilin). For regeneration, the media are poured into deeper dishes (tissue culture dish, 100 × 20 mm, Falcon).

3.2. Preparation of Agrobacterium Standard Inoculum

1. Standard inoculums are prepared using a slightly modified version of the method described in (2). 2. A single colony of Agrobacterium AGL1, containing the appropriate pBract vector together with pSoup, is used to inoculate 10 ml of MG/L medium with 25 µg/ml Rifampicin and 50 µg/ml Kanamycin. This is incubated at 28°C and shaken at 180 rpm for 40 h. 3. Ten millilitres of sterile 30% aqueous glycerol is added to the bacterial culture and mixed by inverting several times. 4. Aliquots of 400 µl of the standard inoculum are placed into 0.5-ml Eppendoff tubes and maintained at room temperature for 2 h mixing by inversion every 30 min. 5. Standard inoculums are then stored at −80°C ready for use.

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3.3. Isolation of Barley Immature Embryos 3.3.1. Collection and Sterilization of Immature Seeds

1. Barley spikes are collected when the immature embryos are 1.5–2 mm in diameter (see Note 4). 2. Before cutting the spikes, a single immature seed from the middle of each spike is checked to make sure that the size of the immature embryos is correct (Fig. 2a–c). 3. The immature seeds are removed from the spike and the awns broken off without damaging the seed coat. 4. The immature seeds are then sterilized by first washing in 70% ethanol for 30 s followed by three washes in sterile distilled water. 5. This is followed by sterilization in a solution of sodium hypochlorite (sodium hypochlorite solution, Fluka 71696) diluted 50:50 with water for 4 min. 6. The sterilization is followed by four washes in sterile distilled water after which the immature seeds are drained but left wet in a screw top sterile jar.

3.3.2. Isolation of Immature Embryos and Removal of Embryonic Axis

1. All operations are performed in a laminar flow hood under sterile conditions. 2. Sterile seeds, approximately 20 at a time, are tipped onto a sterile blue or black tile under a dissecting microscope. 3. The seed is held firm with a pair of fine forceps, and a second pair of fine forceps is used to expose the immature embryo and remove the embryonic axis (see Note 5) (Fig. 2c and d). 4. The embryo is then plated scutellem side up on callus induction medium. 5. Twenty-five embryos are placed on each 9-cm plate ready for Agrobacterium inoculation and stored at 23–24°C in the dark.

Fig. 2. Selection of barley spikes and immature embryos at the correct stage. (a) A barley spike containing immature embryos at the correct stage. (b) Isolated immature seed with immature embryo at the correct stage. (c) Intact immature embryo isolated from immature seed. (d) Immature embryo with the embryonic axis removed ready for Agrobacterium inoculation. (See Color Plate 2)

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1. An Agrobacterium culture is prepared overnight by adding a standard inoculum to 10 ml of liquid MG/L medium without any antibiotics. This is incubated on a shaker at 180 rpm at 28°C overnight (approximately 20 h). 2. The full-strength Agrobacterium culture is used to inoculate the embryos. 3. Using a 200-µl pipette, a small amount of Agrobacterium is dropped onto each embryo so that the surface is just covered. 4. Once all 25 embryos on a plate have been treated, the plate is tilted to allow any excess Agrobacterium culture to run off the embryos. 5. A maximum of two plates are treated with Agrobacterium before the embryos from the first plate are gently removed and transferred to a fresh callus induction plate, scutellum side down. 6. Care is taken not to transfer any excess culture medium or Agrobacterium culture with the embryos, and if necessary the embryos are gently dragged across the surface of the medium to remove excess Agrobacterium before transferring to a fresh plate. 7. Plate are sealed with Micropore surgical tape and incubated at 23–24°C for 3 days. It is convenient to isolate immature embryos one day and to inoculate with Agrobacterium the following day. However, it is also possible to inoculate the embryos on the same day as they are isolated (see Note 6).

3.5. Selection of Transformed Material

1. After 3 days co-cultivation, the embryos are transferred to fresh callus induction plates containing hygromycin as the selective agent and Timentin to remove Agrobacterium from the cultures (see Note 7). 2. Embryos are cultured scutellum side down at 23–24°C in the dark. This transfer is referred to as selection 1. 3. After 2 weeks, embryos are transferred to fresh selection plates as above (selection 2). The entire embryo and callus derived from it is transferred as a single unit and not split up. 4. After a further 2 weeks, each embryo is transferred to a third selection plate (selection 3) (see Note 8). At this stage the callus from each embryo may break up, and if so, it should be placed in a marked area of the plate so that all material derived from a single embryo can be tracked (Fig. 3b). 5. It is necessary to reduce the number of embryos per plate as the transformed embryo-derived callus starts to grow rapidly. 6. After 6 weeks’ selection on callus induction medium, the embryo-derived callus is transferred to transition medium,

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Fig. 3. Selection of transformed barley on hygromycin-containing media. (a) Regenerating transformed barley lines around the edge of a plate of regeneration medium with non-transformed lines in the middle. (b) Plates of regenerating transformed barley. Plate in the foreground shows four transformed lines together with non-transformed lines and also the method of marking the plate to distinguish material from individual embryos. (c) Transgenic barley plant transferred to culture tube showing good root system in hygromycin-containing medium. (d) Transgenic barley plantlets in the culture room just before transfer to soil.(See Color Plate 3)

again containing hygromycin and Timentin, for 2 weeks at 24°C under low light. The low light is achieved by placing the plates under light conditions in a tissue-culture room but covering the plates with a thin sheet of paper. During this 2-week culture period, transformed lines should become obvious and start to produce green areas and small shoots. Non-transformed callus rarely shows signs of regeneration on hygromycin-containing medium. 3.6. Regeneration of Transgenic Plants

1. After the 2 weeks on transition medium, the embryo-derived material is transferred one final time to regeneration medium, in deep Petri dishes, without any growth regulators but still with the same levels of hygromycin and Timentin. 2. As transformed lines will grow very vigorously, the number of lines per plate must be further reduced. 3. The appearance of typical regeneration plates can be seen in Fig. 3a and b. It is important to keep all regenerating calluses derived from a single embryo together. 4. Once shoots are 2–3 cm in length and roots have formed, carefully remove the small plantlets from the plates and transfer to glass culture tubes (Sigma C-5916) containing 12 ml of callus induction medium but without any dicamba or other growth regulators. This medium should still contain hygromycin and Timentin at the same concentrations. 5. The appearance of the plantlets in tubes can be seen in Fig. 3c and d. The plantlets should quickly form a strong root system in the hygromycin-containing medium.

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6. Plants that form strong roots in hygromycin are always found to be transformed, and this selection system does not give any ‘escapes’ or non-transformed plants that survive selection (see Note 9). 7. Once rooted plants reach the top of the tubes, they can be transferred to soil. 8. Plants are gently removed from tubes using long forceps and all tissue culture medium washed from the roots under a running tap. They are then planted in the same barley growth mix described above in 5-cm diameter pots. 9. Plants are covered either with small individual plastic jars with holes at the top as individual propagators, or entire trays are covered with propagators for a few days to maintain humidity until the plants are well established in soil. 10. Once plants are established in soil, leaf samples can be collected for further analysis to confirm the presence of the introduced genes. Figure 4 shows the expression of the gus gene in leaf samples from plants transformed with pBract 204.

Fig. 4. The results of staining leaf samples for GUS activity following transformation with pBract 204. Wells A and F do not show any GUS activity. The other wells show varying levels of GUS expression. (See Color Plate 4)

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4. Notes 1. The plants used to provide the immature embryos for transformation should not be sprayed with any pesticides. Spraying the plants as they approach flowering is a particular problem, as it can reduce the response of the immature embryos in tissue culture. 2. T-DNA transfer starts at the right border and proceeds to the left border. Therefore it is advisable to have the selectable marker at the left border and the gene of interest at the right border so that the gene of interest is transferred first and the selectable marker second. This means that selected transformed plants have a greater chance of containing the gene of interest. 3. Phytagel can be prepared in advance at 2× concentration in water, autoclaved, and stored at room temperature because it will not solidify until the other culture medium components are added. 4. The optimum size of immature embryos is 1.5 mm in diameter. Smaller embryos of 1 mm diameter are difficult to handle and more likely to suffer from overgrowth of Agrobacterium. Embryos of 2 mm diameter are at the upper limit of those that will respond well in culture. At the correct stage, the endosperm of the immature seeds should still be soft and quite liquid in appearance. If the endosperm is starting to look floury, then the embryos will be too old. 5. There are several different techniques for removing the immature embryos and removing the embryonic axis. The most efficient way is to carry out this operation in one step holding the immature seed by puncturing it with one pair of fine forceps and then exposing the immature embryo by peeling back the seed coat from the awn end of the seed. Once the embryo is exposed, the embryonic axis can be removed by first pinching off the shoot axis with the fine forceps and then removing the root axis with a second pinching action. This should leave only the undamaged scutellum. If preferred, it is possible to carry out this operation in two steps. First, the immature embryos can be removed and placed intact onto callus induction medium. Once the required numbers have been isolated, they can be picked up in groups of around ten together with a small amount of culture medium, placed on the sterile tile, and the axis removed from each using the fine forceps. They are then immediately plated scutellum side up on callus induction medium. 6. Transformation efficiencies of 25% have been obtained by infecting immature embryos either on the same day as isolation or on the day following isolation. Transformation efficiency is defined as the number of independent transformed plants

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as a percentage of the original number of immature embryos treated. 7. Using the procedure described, excessive overgrowth of the immature embryos with Agrobacterium should be avoided. However, if any embryo is completely overgrown with Agrobacterium following the co-cultivation then it should be discarded. This can be a problem if very small embryos are used. 8. If callus growth looks good after 4 weeks of selection, then it is possible to omit the final 2 weeks selection on callus induction medium (selection 3) and transfer to transition medium at this stage. If in any doubt, continue with selection 3, as this is likely to yield better results. 9. Although non-transformed plantlets do not normally survive and root on hygromycin-containing medium, it is occasionally possible to have two shoots close together, one transformed and one non-transformed. These shoots may be transferred together to a culture tube. This is not normally obvious until the plants are removed from the tube and the roots rinsed and ready for planting in soil. In this case, it is possible for a transformed shoot to maintain the growth of a non-transformed shoot.

References 1. Wan, Y. and Lemaux, P. G. (1994) Generation of large numbers of independently transformed fertile barley plants. Plant Physiol. 104, 37–48. 2. Tingay, S., McElroy, D., Kalla, R., Fieg, S., Wang, M., Thornton, S. and Brettell, R. (1997) Agrobacterium tumefaciens-mediated barley transformation. Plant J. 11, 1369–1376. 3. Kumlehn, J., Serazetdinova, L., Hansel, G., Becker, D. and Lorz, H. (2006) Genetic transformation of barley (Hordeum vulgare L.) via infection of androgenetic pollen cultures with Agrobacterium tumefaciens. Plant Biotechnol. J. 4, 251–261. 4. Travella, S., Ross, S. M., Harden, J., Everett, C., Snape, J. W. and Harwood, W. A. (2005) A comparison of transgenic barley lines produced by particle bombardment and Agrobacterium-mediated techniques. Plant Cell Rep. 23, 780–789. 5. Wang,M.-B.,Abbott,D.C.,Upadhyaya,N.M., Jacobsen, J. V. and Waterhouse, P. M. (2001) Agrobacterium tumefaciens-mediated transformation of an elite Australian barley cultivar with virus resistance and reporter genes. Aust. J. Plant Physiol. 28, 149–156.

6. Matthews, P. R., Wang, M. B., Waterhouse, P. M., Thornton, S., Fieg, S. J., Gubler, F. and Jacobsen, J. V. (2001) Marker gene elimination from transgenic barley, using co-transformation with adjacent “twin TDNAs” on a standard Agrobacterium transformation vector. Mol. Breed. 7, 195–202. 7. Shrawat, A. K., Becker, D. and Lorz, H. (2006) Agrobacterium tumefaciens-mediated genetic transformation of barley (Hordeum vulgare L.) Plant Sci. 172, 281–290. 8. Harwood, W. A., Ross, S. M., Cilento, P. and Snape, J. W. (2000) The effect of DNA/gold particle preparation technique, and particle bombardment device, on the transformation of barley (Hordeum vulgare). Euphytica 111, 67–76 9. Hellens, R. P., Edwards, E. A., Leyland, N. R., Bean, S. and Mullineaux, P. M. (2000) pGreen: a versatile and flexible binary Ti vector for Agrobacterium-mediated plant transformation Plant Mol. Biol. 42, 819–832. 10. Garfinkel, M. and Nester, E. W. (1980) Agrobacterium tumefaciens mutants affected in crown gall tumorigenesis and octopine catabolism. J. Bacteriol. 144, 732–743.

Chapter 10 Transformation of Oats and Its Application to Improving Osmotic Stress Tolerance Shahina B. Maqbool, Heng Zhong, Hesham F. Oraby, and Mariam B. Sticklen Abstract Oat (Avena sativa L.), a worldwide temperate cereal crop, is deficient in tolerance to osmotic stress due to drought and/or salinity. To genetically transform the available commercial oat cultivars, a genotypeindependent and efficient regeneration system from shoot apical meristems was developed using four oat cultivars: Prairie, Porter, Ogle, and Pacer. All these oat cultivars generated a genotype-independent in vitro differentiated multiple shoots from shoot apical meristems at a high frequency. Using this system, three oat cultivars were genetically co-transformed with pBY520 (containing hva1 and bar) and pAct1-D (containing gus) using biolistic™ bombardment. Transgenic plants were selected and regenerated using herbicide resistance and GUS as a marker. Molecular and biochemical analyses of putative transgenic plants confirmed the co-integration of hva1 and bar genes with a frequency of 100%, and 61.6% of the transgenic plants carried all three genes (hva1, bar and gus). Further analyses of R0, R1, and R2 progenies confirmed stable integration, expression, and Mendalian inheritance for all transgenes. Histochemical analysis of GUS protein in transgenic plants showed a high level of GUS expression in vascular tissues and in the pollen grains of mature flowers. Immunochemical analysis of transgenic plants indicated a constitutive expression of hva1 at all developmental stages. However, the level of HVA1 was higher during the early seedling stages. The characteristic of HVA1 expression for osmotic tolerance in transgenic oat progeny was analyzed in vitro as well as in vivo. Transgenic plants exhibited significantly (P <0.05) increased tolerance to stress conditions than non-transgenic control plants. The symptoms of wilting or death of leaves as observed in 80% of non-transgenic plants due to osmotic stress was delayed and detected only in less than 10% of transgenic plants. These observations confirmed the characteristic of HVA1 protein as providing or enhancing the osmotic tolerance in transgenic plants against salinity and possible water-deficiency stress conditions. Key words: Oat, (Avena sativa L.), hva1, bar, gus, shoot apical meristem, transgenic, progeny, osmotic stress.

1. Introduction Oat (Avena sativa L.) is a cool-season annual cereal crop, ranking seventh in the world cereal production after wheat, maize, rice, barley, sorghum, and millet. In contrast to other grains, most oats Huw D. Jones and Peter R. Shewry (eds.), Methods in Molecular Biology, Transgenic Wheat, Barley and Oats, vol. 478 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-379-0_10

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are consumed domestically and there is little export trade (1, 2). Oats thrive in cool, moist climates, but adapt to various soil types (3, 4). It has a wider pH adaptability than wheat or barley, ranging from 5.5 to 7.0, and even as low as 4.5 for some varieties. Oat also has a low lime requirement (5). However, it requires sufficient water for growth and grain production. Therefore, Russia, Canada, the United States, Finland, and Poland are the major oat-producing countries (6). The highest commercial yields for oats tend to come from the Netherlands, Switzerland, West Germany, the United Kingdom, Ireland, Sweden, and France (7). Because of the high levels of protein and essential minerals, oats have been an excellent livestock feed for centuries (8). In addition, the high fiber content of oats aid in digestion (6). Oats are also used for human food, in the form of oatmeal, oat starch, and cookies. Oats and oat bran have recently gained much popularity as health food products because of its characteristic to reduce blood cholesterol and regulate gastro-intestinal function (6). Moreover, most recently, oat is also used in cosmetics because of its two active compounds, avenanthramides and betaglucan. These compounds have been found beneficial for the skin care and are being used as anti-irritants and as aids in skin regeneration after damage by sunburns, etc. Oats are also used in paper and brewing industries, as well as in the production of plastics, pesticides, and preservatives (7). Oat is also used for hay, pasture, green manure, or as a cover crop. As a cover crop it enhances soil life, suppresses weeds, provides erosion control, and increases organic component (4, 8). All this has resulted in more demand for high-quality oats. Environmental stresses, such as drought and salinity, limit agricultural productivity worldwide (9, 10). In particular, about one-third of the world’s irrigated agricultural land is affected by salinity (11, 12). Arable land area is decreasing day by day because of bad farming practices (13). This has left no choice for the farmers but to cultivate in salinity-prone areas, and harvest salinity-stressed crops (14). Oat is sensitive to hot, dry weather and the production of oat grains depends upon the availability of sufficient water during the growth season (15). The adaptability of oat towards salinity is not much known. However, it is considered relatively less salt tolerant than other cereal or forage crops (16). Some reports describe the reduction in seed germination and subsequent development in different oat cultivars due to salinity (16, 17). Lodging is another threat to oat production. Oat is also susceptible to barley yellow dwarf viruses and insect pests (18, 19). Oat varieties have been modified to generate resistance towards its pathogens, but only at a limited scale (20).

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This problem cannot be solved by traditional breeding practices because of the conserved genomic background in oat varieties and inefficient selection methods (21). However, modern crop improvement strategies, such as biotechnology, can be used in conjunction with traditional breeding efforts to enhance agricultural productivity (22– 24). Biotechnology offers a promising way to not only eliminate the existing problems limiting crop production in developing countries but also to promote sustainable agriculture (25, 26). Therefore, a wise selection of varieties with genetic potential or the development of resistant cultivars via genetic engineering may help improve oat production under stress conditions. Recent improvements in the production of cereal crops depend upon the tissue culture and genetic engineering of useful trait genes (27– 29). However, the information about the genotype independency for in vitro regeneration and genetic engineering system of oat is limited (29– 34). In 1996, Zhang et al. developed a genotype-independent efficient regeneration system from shoot apical meristems for the genetic transformation of commercial oat cultivars (35). Later studies describe the use of BiolistcTM mediated-transformation system for genetic transformation of oat, using mature embryo-derived callus (36, 37), leaf base segments of young seedlings (38), and embryogenic calli derived from immature embryos (39). However, embryo-derived callus is not considered feasible in routine oat transformations because of the risks for somaclonal variations that may appear in the callus cultures due to the prolonged tissue culture (40, 41). Therefore, multiple shoot meristem culture derived from mature seeds was adopted as an alternative genotype-independent regenerable target tissue for genetic transformation of commercial cultivars of oat (35–47). The advantage of using this system includes the production of target tissues with highest frequency of regeneration and reproducibility. It also provides the maximum level of fertility and genomic stability in regenerated transgenic plants. Here, we describe the method of producing shoot apical meristems and biolistic-mediated genetic transformation of three oat cultivars and its application to improve osmotic stress tolerance with the use of an osmotic stress resistant gene hva1 (47– 51) as an example. Further, we describe the molecular and biochemical analysis of transgenic oat plants to confirm transgene integration, expression stability, and inheritance as well as determination of salinity and water-deficit tolerance in hva1 expressing transgenic plants.

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2. Materials

2.1. Oat Cultivars

1. Ogle (Brave/2/Tyler/Egdoion 23) 2. Pacer (Coachman × CI 1382) 3. Prairie (IL73-5743 × Ogle)

2.2. Plasmids

1. pBY520 containing two genes: a herbicide-resistant phosphinothricin acetyl transferase (bar) gene from Streptomyces hygroscopicus, under the control of cauliflower mosaic virus (CaMV35S) promoter and the 3′ non-coding region (nos) of Agrobacterium nopaline synthase gene, as selection marker; and the late embryogenesis abundant protein (hva1) gene from barley (Hordeum vulgare L.), under the control of rice actin 1 promoter (Act1) and 3′ non-coding region (pin II) of potato protease inhibitor II gene, for stress-resistance (Fig. 1). 2. pAct1-D containing the Escherichia coli β-glucuronidase (gus) gene under the control of the Act1 promoter and the nos terminator (Fig. 1).

2.3. Culture Media

1. MS1 (germination medium): 4.3 g/l Murashige and Skoog (52) basal salts (MS) and vitamins (GIBCO BRL Rockville, Maryland), 30 g/l sucrose (Sigma, St. Louis, MO, USA), and 3 g/l phytagel (Sigma), pH 5.6. 2. MS2 (shoot multiplication medium): MS medium, 30 g/l sucrose, 500 mg/l casein enzymatic hydrolysate (Sigma), 0.5 mg/l 2,4-dichlorophenoxyacetic acid (Sigma), 2 mg/l N6benzyladenine (Sigma), and 3 g/l phytagel, pH 5.6. 3. MS3: (particle bombardment medium): MS medium, 30 g/l sucrose, 500 mg/l casein enzymatic hydrolysate (Sigma), 0.5

Fig. 1. Schematic representation of the plasmids BY520 and Act1-D.

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mg/l 2,4-dichlorophenoxyacetic acid (Sigma), 2 mg/l N6benzyladenine (Sigma), and 5 g/l phytagel. 4. MS4 (shoot multiplication medium with less selection pressure): MS2 medium, 5 mg/l of glufosinate ammonium (Sigma) or 2 mg/l of Bialaphos (Meiji Seika Kaisha, Japan). 5. MS5 (shoot multiplication medium with high selection pressure): MS2 medium, 10 mg/l of glufosinate ammonium or 3 mg/l of Bialaphos. 6. MS6 (shoot regeneration medium): MS medium, 20 g/l sucrose, 0.5 mg/l N6-benzyladenine, 0.5 mg/l indole butyric acid (Sigma), 15 mg/l of glufosinate ammonium or 5 mg/l of Bialaphos, and 3–5 g/l phytagel, pH 5.6. 7. MS7 (root formation medium): 2.15 g/l MS medium, 10 g/l sucrose, and 15 mg/l of glufosinate ammonium or 5 mg/l of Bialaphos, and 3–5 g/l phytagel, pH 5.8. 8. MS8 (salt stress medium): 2.15 g/l MS medium, 10 g/ l sucrose, 100 mM NaCl (Sigma), and 3–5 g/l phytagel, pH 5.8. 9. MS9 (osmotic stress medium): 2.15 g/l MS medium, 10 g/l sucrose, 200 mM manifold (Sigma), and 3–5 g/l phytagel, pH 5.8. All media should be sterilized by autoclaving at 121°C for 20 min. Hormones, herbicides, or antibiotics should be added to the medium after autoclaving (50–40°C). 2.4. Other Materials

1. Clorox bleach (Clorox Professional Products Company, Oakland, CA 94612) 2. Plastic plant pots (8 cm square, Griffin Green House and Nursery Supplies, 1619 Main Street, Tewksbury, MA 01876). Peat and perlite soil mix (Griffin Green House and Nursery Supplies) 3. Soil mix (Metro Mix 360 Soil; Griffin Green House and Nursery Supplies) 4. Ignite® herbicide containing 200 g/l (16.222%) of glufosinate as the active ingredient (1% Ignite Hoechst-Roussel Agri-Vet Company, NJ) 5. Protein extraction buffer: 50 mM sodium phosphate (pH 7.0), 10 mM EDTA (ethylenediaminetetraacetate), 0.1% (v/ v) Triton X-100, 0.1% (w/v) Sarkosyl, 10 mM mercaptoethanol, and 10 mM PMSF (phenylmethylsulfonyl fluoride) 6. Alkaline phosphatase buffer: 100 mM Tris-base, 100 mM NaCl, 5 mM MgCl2, pH 9.8) containing 0.33 mg/ml of NBT (Nitro blue tetrazolium), and 0.16 mg/ml of BCIP (5bromo-4-chloro-3-indolyl phosphate)

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7. GUS substrate: 10 mM EDTA (pH 7.0), 0.1 M phosphate buffer (pH 7.0) and 1–5 mM X-Gluc (X-Gluc can be dissolved in dimethyl sulfo oxide; Sigma)

3. Methods 3.1. Shoot Tip Cultures

1. Use mature seeds of oat cultivars (Ogle, Pacer and Prairie) for shoot tip cultures. 2. Remove the lemmas and the paleas from the caryopses (seeds) by hand. 3. Surface sterilize the seeds using 70% ethanol (Sigma) for 5 min, wash with autoclaved distilled water once, treat with 20% for 30 min with constant shaking at 0.5 × g in a rotary shaker (VWR International, Bristol, CT 06011) (see Note 1), and wash three times with autoclaved distilled water. 4. For acclimatization, place the seeds (under sterile conditions) into a Petri dish containing moist filter paper and cover with a wet filter paper and incubate at 3–5°C for 2–3 days. Supplement water regularly to keep the moisture. 5. After acclimatization, germinate 10–12 surface-sterilized seeds aseptically in a Petri dish (100 × 15 mm; diameter × height; VWR) containing MS1 (25 ml/dish) and incubate in the growth room or incubator set at 25°C under darkness for a week (see Note 2). 6.

After germination, cut the shoot apices into small sections (3–5 mm in length) aseptically using sterilized scalper blades (Sigma) (see Note 3).

7. Culture five to seven excised shoot apices on MS2 (25 ml/ dish) horizontally in a Petri dish (100 × 15 mm) and incubate under constant light (60 µm/m2/s from cool-white 40 W Econ-o-watt fluorescent lamp; Philips Westinghouse, USA) at 25°C for 4 weeks (see Note 4). 8. Subculture the shoot apices after every week by removing the elongating leaves, coleoptiles, and stems of cultured shoot apices physically and cut again to 3–5 mm in length. 9. Maintain multiple shoot cultures by subculturing at 2-week intervals on MS2 medium (Fig. 2a). 10. Calculate the relative frequency of differentiation of multiple shoots after 8 weeks of culturing as the percentage of multiplied shoot apices that produce multiple shoots in total cultured shoot apices for each cultivar (see Note 5).

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Fig. 2. Transformation of oat (Avena sativa L.). (a) Four-week old cultured shoot apical meristems; (b) proliferated shoot cultures 2 weeks after bombardment without selection; (c ) regenerated transgenic shoots on selection medium; (d) matured transgenic oat plants in greenhouse; (e) GUS expression in multiple shoots; (f) localized GUS expression in the stem cross-section; p pith; v vascular bundles; c cortex; (g) mature pollen showing GUS expression; (h) GUS expression in seeds; (i) oat seedlings grown on salt-stress medium (100 mM); (j) oat plants watered with 150 mM saline solution. (k) Roots of oat plants watered with 100 mM saline solution; T: transgenic; C: non-transgenic. (See Color Plate 5)

3.2. Microprojectile Bombardment

1. For bombardment, sterilize 30 mg of gold particles (1 µm in diameter; Bio-Rad, Richmond, CA, USA) or tungsten particles (0.9 µm in diameter; GTE Sylvania, Towanda, PA, USA) in a 1.5-ml microcentrifuge tube (VWR) using 1 ml of 100% ethanol for 30 min with continuous vortexing at high speed. 2. While vortexing, take an aliquot of 50 µl of the particle–ethanol suspension into a new microcentrifuge tube and centrifuge for 30 s at 10,000 × g using a microcentrifuge (Brinkman, Westbury, NY 11590), wash with 1 ml autoclaved distilled water, vortex, and centrifuge again. After washing twice, resuspend the particles in 332 µl of autoclaved distilled water. 3. To the re-suspended particles add 15 µl of DNA sample containing 15 µg of pBY520 and pAct1-D at a 1:1 molar ratio, 225 µl of 2.5 M CaCl2 (Sigma), and 50 µl of 0.1 M spermidine (Sigma), and vortex for 5 min at room temperature (see Note 6).

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4. Incubate the particle–DNA suspension on ice for 10 min and centrifuge at 10,000 × g for 1 min. 5. Wash the particle–DNA pellet with 500 µl of 100% ethanol, vortex for 30 s, centrifuge for 1 min, and re-suspend in 100 µl of 100% ethanol. 6. Position the shoot tip cultures (2–3 shoot clumps depending upon the size) in the area of 1.5 cm diameter in a Petri dish containing MS3 (25 ml/dish) below the microprojectilestopping screen (see Note 7). 7. For bombardment, pipette 10 µl of the particle–DNA suspension onto the center of the macrocarriers and use for bombardment as soon as the ethanol evaporates (see Note 8). 8. Repeat the bombardment for each shoot tip cultures once with an interval of 2 h (see Note 9). 9. Transfer the bombarded shoot tip cultures to fresh MS2 (25 ml/dish) to proliferate for 4 weeks in continuous light (60 µm/m2/s) at 25°C with one subculture (Fig. 2b). 3.3. Selection of Transformed Tissues

1. After 4 weeks, carefully (avoid injury to shoot meristems) divide the bombarded shoot clumps into small clumps (5–10 mm in diameter) and transfer 6–8 clumps/plate to MS4 (25 ml/dish; 100 × 15). Calculate the relative frequency of transformation events (Table 1). 2. After 2 weeks, subculture the green shoot clumps (transgenic) on the fresh MS4 and discard the yellow or brown shoot tip cultures (non-transgenic).

Table 1 Relative frequency of differentiation of multiple shoot clumps after micro projectile bombardment on selection medium Cultivars

No. of bombarded shoot clumps

No. of independent eventsa

Transformation frequency (%)b

Ogle

15

11

73.3

16

13

81.2

15

11

73.3

18

15

83.3

17

12

70.5

12

10

83.3

Pacer

Prairie

a

Total number of proliferated independent shoot clumps after the first month of selection Proliferated shoot clumps after the first month of selection divided by total number of bombarded shoot tip clumps, ×100 b

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3. After 4 weeks of selection on MS4, further divide the green shoot clumps into 5–10 mm in diameter and subculture on MS5 (25 ml/dish; 100 × 20) for 4–6 weeks. 4. After a total of 4 months of selection and multiplication, transfer the fast growing multiple shoot clumps to Magenta boxes (99 × 68 mm; length × diameter; Sigma; 2–3 clumps/ Magenta) containing MS6 (50–70 ml/Magenta) for vegetative growth and then to MS7 (50–70 ml/Magenta) for root development (Fig. 2c). 5. When putative transgenic plants reach 5–10 cm in length with 2–3 leaves, transplant, one plant/pot, to plastic 8cm pots containing soil mix composed of 1:1 (v:v) peat and perlite. Grow the plants in a greenhouse under 16-h photoperiod and 70–80% humidity until maturity (Fig. 2d; see Note 10). 6. Let self-pollination of the transgenic plants to produce seeds, harvest the seeds, and store individually at 4°C and 70% humidity. 3.4. Analysis of Transgene Inheritance

1. After acclimatization, germinate seeds (10–12 seeds/dish) for 1 week on MS7 (25 ml/dish) to analyze transgene inheritance, e.g. bar in the R0, R1, and R2 plants (see Note 11). 2. Or, germinate seeds in soil (Metro Mix 360 Soil) using multicell plastic trays (Griffin Green House and Nursery Supplies) in the greenhouse for 2 weeks. Spray the growing plants with herbicide (see Note 12). 3. After 1 week, take the count of germinated and the non-germinated seeds on selection medium, or surviving and the dead seedlings after herbicide spray in the greenhouse. 4. Apply statistic χ2 test (53) for segregation analysis of transgene in R0, R1, and R2 progenies (Table 2).

3.5. Molecular Analyses of Transgenic Plants 3.5.1. Polymerase Chain Reaction (PCR) Analysis

1. For the detection of transgenes (e.g. bar and hva1) in the transgenic plants, use leaf discs to extract genomic DNA and REDExtract-N-Amp™ Plant PCR Kit (Sigma-Aldrich, St. Louis, MO, Cat # XNA-P) as per the manufacturer’s instruction. 2. Use forward (F) and reverse (R) PCR primers: For example: bar-F, 5′-ATG AGC CCA GAA CGA CG-3′; bar-R, 5′-TCA GAT CTC GGT GAC GG-3′ and hva1-F, 5′-TGG CCT CCA ACC AGA ACC AG-3′; hva1-R, 5′-ACG ACT AAA GGA ACG GAA AT-3′. 3. Perform DNA amplifications in a thermocycler (such as Perkin Elmer/Applied Biosystem, Foster City, CA) using initial denaturation at 94°C for 4 min, followed by 35 cycles of 1 min at 94°C, 1 min at 55°C, 2 min at 72°C, and a final 10 min extension at 72°C.

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Table 2 Analysis of transgene inheritance in R1 transgenic oat plants Transformants

Germinated seeds

Non-germinated seeds

χ2

P-value*

Ogle BRA-17

70

20

0.37

0.54

Ogle BRA-19

60

15

1.00

0.32

Ogle BRA-41

89

26

0.33

0.56

Pc.BA-4

96

30

0.09

0.75

Pc.BA-10

159

61

0.87

0.35

*No significant difference between the observed versus the 3:1 expected segregation (P > 0.05)

Fig. 3. Molecular analyses of hva1 in transgenic oat plants. (a) PCR amplification; (b) Southern blot analysis; (c) Northern blot analysis; (d) Western blot analysis. Lanes 1–5: R2 transgenic oat plants, Ogle BRA-82, Ogle BRA-17, Ogle BRA-8, Ogle BRA-19, and Ogle BRA-41, respectively; c non-transgenic oat plant; p positive control, plasmid DNA (pBY520) or purified HVA1 protein.

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4. Separate and analyze PCR products onto a 0.8% (w/v) agarose gel, stain with ethidium bromide, and visualize under UV light. The transgene product size is about 0.59 kb for bar gene and 0.70 kb for hva1 gene (Fig. 3a). 3.5.2. Southern Blot Analysis

1. For Southern blot hybridizations (54) use the transgene-coding sequence (e.g. hva1 or bar1) as a probe. 2. Isolate the genomic DNA from the transgenic and nontransgenic plants using Phytopure Plant DNA extraction kit (Amersham-Pharmacia Biotech). 3. Use 10 µg of genomic DNA from R0, R1, or R2 progenies to prepare Southern blots. 4. Digest the genomic DNA with HindIII or HindIII-BamHI restriction enzymes and fractionate on a 0.8% agarose gel. 5. Denature the gels, neutralize, and blot onto Hybond-N + membranes (Amersham-Pharmacia Biotech) according to Sambrook et al. (55). 6. Digest pBY520 with HindIII–BamHI restriction enzymes to isolate a fragment containing the hva1-coding sequence. Separate the restricted fragment on 0.8% low-melting agarose, elute the 1.0-kb fragment, and purify using the QIAquick PCR purification kit (QIAGEN). 7. Radio-label the purified, gene specific, DNA fragment with [32P]-dCTP using the Rad Prime labeling kit (GIBCO BRL) according to the manufacturer’s instructions. 8. Hybridize the prepared membrane blot with the gene-specific probe (hva1) according to standard procedures (55). 9. Subsequently, analyze the hybridize membranes by autoradiography using X-ray film (Kodak) at −80°C (Fig. 3b).

3.5.3. Northern Blot Analysis

1. Isolate the total RNA from young leaves of transgenic and non-transgenic plants using the TRIZOL Reagent (GIBCO BRL) according to the manufacturer’s instructions. 2. Fractionate 10 µg of total RNA using 1.2% agarose–formaldehyde denaturing gel according to Sambrook et al. (55). 3. Blot the gel onto Hybond-N nylon membrane (AmershamPharmacia Biotech) according to Sambrook et al. (55). 4. Analyze the transgene transcripts (hva1) with a standard northern-blotting procedure (55) using 32P labeled gene-specific probe (Fig. 3c).

3.5.4. Western Blot Analysis

1. Protein isolation, western blotting, and immunodetection protocols were as per Xu et al. (48). 2. Pulverize about 0.1 g of fresh leaf tissue from transgenic and non-transgenic plants each in liquid nitrogen and extract in 0.2 ml of protein extraction buffer.

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3. Measure total protein concentration in each sample using the Bio-Rad protein assay reagent (Bio-Rad) according to Bradford (56). 4. Fractionate 100 µg of total soluble protein from transgenic and non-transgenic samples using 12% SDS polyacrylamide gels. 5. After electrophoresis, electroblott the proteins onto nitrocellulose membrane (Amersham-Pharmacia Biotech) using a Semi-Dry Transfer Cell (Bio-Rad) following the manufacturer’s instructions. 6. Incubate the western blots with primary antiserum raised (in rabbit e.g.) against transgene-specific protein (e.g. HVA1) and secondary antiserum (alkaline phosphatase conjugated anti-rabbit IgG, Sigma-Aldrich). 7. Develop the membrane in alkaline phosphatase buffer (Fig. 3d). 3.6. Biochemical Analysis of Transgenes

1. For biochemical analysis of transgenes e.g. GUS, use the whole plant tissues or different organs from the transgenic and nontransgenic plants (Fig. 2e–h; (48)). 2. To remove chlorophyll, incubate the green tissues first in 70% ethanol for 2 h and then in 100% ethanol overnight. 3. Provide some vacuum to the tissues to remove the trapped air. 4. Immerse the samples in GUS substrate (10 mM EDTA (pH 7.0), 0.1 M NaPO4 (pH 7.0), and 1–5 mM X-Gluc; X-Gluc can be dissolved in dimethylsulfoxide; Sigma) immediately after vacuum treatment and incubate at 37°C as described in (57). 5. For localization of GUS expression, prepare the tissues using free-hand cross-sections and examine under a Zeiss SV8 stereomicroscope or a Zeiss Axioskop routine microscope.

3.7. Evaluating Transgenic Plants Expressing hva1 Under Stress Conditions 3.7.1. Salt Stress In Vitro

1. Surface sterilize 50 or 60 seeds (R0, R1, or R2 seeds; see Note 12) from each transgenic and non-transgenic line. After acclimatization, germinate 10–12 seeds/dish in the dark at 25°C on MS7 medium (25 ml/dish) for 4 days (see Note 13). Use MS7 medium without glufosinate ammonium or Bialaphos for non-transgenic seeds. 2. Divide the germinated seedlings into two sets: one set to grow without salt stress and the other to grow under salt stress conditions. 3. Transfer four or five growing seedlings from each independent transgenic and non-transgenic line to Magenta boxes containing MS8 (50–70 ml/Magenta) with and without NaCl and let them grow under light (60 µm/m2/s) at 25°C with a minimum of four replicates. 4. After 6 days analyze the growth of transgenic and non-transgenic plants growing with and without salt (Fig. 2i).

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5. Take the measurements such as plant fresh weight (see Note 14), plant height, root length, and plant dry weight (see Note 15; Table 3). 6. Apply analysis of variance (58) to statistically analyze the data. 7. Separate means by using Tukey’s Studentized range test at 95% confidence level. 8. Transfer the plants to soil mixture composed of 1:1 (v/v) peat:perlite for further growth and development in the greenhouse. 1. Germinate seeds (R0, R1, or R2 seeds) on MS7 medium as mentioned in Section 3.7.1.

3.7.2. Salt Stress In Vivo

2. Transfer 1-week old growing seedlings with (transgenic) and without selection (non-transgenic control plants) into small pots (8 × 4 × 6 cm) containing soil mix composed of 1:1 (v:v) peat and perlite (one plant per pot). 3. Keep the pots in water-filled, flat-bottom trays and let them grow in the greenhouse for an additional week before starting salt stress treatments. 4. Divide the growing plants into five sets: use eight or ten plants for each transgenic line and eight or ten non-transgenic control plants for each set. Before the start of stress treatments, take measurements such as the initial plant height and leaf numbers of each plant.

Table 3 In vitro analyses of stress tolerance using young oat seedlings Shoot length Non-stressed (cm)

Salt stressed (cm)

Non-transformed

10.67 ± 0.45

6.26 ± 1.1

Ogle BRA-82

12.23 ± 0.52* 9.05 ± 0.5*

Lines

Ogle BRA-17

11 ± 1

Ogle BRA-8

10.8 ± 0.7

7 ± 0.64

Root length Osmotic stressed (cm) 5 ± 1.2

Nonstressed (cm) 4.02 ± 0.45

Salt stressed (cm)

Osmotic stressed (cm)

2 ± 0.6

2.4 ± 0.6

9 ± 0.56*

5.8 ± 0.88* 3.3 ± 0.48* 3.9 ± 0.52*

9 ± 0.7*

5.5 ± 0.7*

2.5 ± 0.4

3.4 ± 0.3

9.3 ± 0.4*

8±1

3.64 ± 0.6

2.9 ± 0.3

3.3 ± 0.4

Ogle BRA-19

11 ± 0.8

8.5 ± 0.9

9.4 ± 0.4*

3.9 ± 0.4

3.8 ± 0.6*

3.8 ± 0.5*

Ogle BRA-41

12.3 ± 0.9*

8.5 ± 0.8

9.1 ± 0.63*

4.3 ± 0.9

2.7 ± 0.4

3.6 ± 0.25*

*P < 0.05

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5. Water the plants with different saline solutions (0, 50, 100, 150, and 200 mM NaCl) once per day for 14 days (see Note 16). 6. Follow by 1 week of irrigation with regular water to allow plants to recover. 7. Afterwards, resume the salt treatments uninterrupted for 5 weeks. 8. Perform this experiment for a minimum of four or five replicates. 9. At the end, take the measurements of plant height, root length, number of tillers, and kernel yield of each plant, and compare the growth difference between stressed and nonstressed transgenic and non-transgenic plants (see Note 17; Fig. 2j and k). 10. Apply analysis of variance (58) to statistically analyze the data. 11. Separate means by using Tukey’s Studentized range test at 95% confidence level (Table 4). 1. Germinate seeds (R0, R1, or R2 seeds) on MS7 medium as mentioned in Section 3.7.1. (see Note 18).

3.7.3. Water Deficit or Osmotic Stress In Vitro

2. Divide the germinated seedlings into two sets: one set to grow without water deficit or osmotic stress, and the other with water deficit or osmotic stress (see Note 19). 3. Transfer four or five growing seedlings from each independent transgenic and non-transgenic line to Magenta boxes containing MS9 (50–70 ml/Magenta) with and without

Table 4 In vivo analysis of stress tolerance using oat plants Kernel yield/plant (g) Line

0 mM NaCl

Non-transformed

16.93

7.26

3.75

2.01

1.11

Ogle BRA-82

12.10

10.29

9.19*

7.64

5.96

Ogle BRA-17

15.44

14.13

13.66*

10.04*

5.90

Ogle BRA-8

16.05

14.45

13.52*

9.82*

5.14

Ogle BRA-19

11.75

11.74

10.59*

8.72*

6.85*

Ogle BRA-41

14.31

13.95

13.73*

9.23*

8.37*

*P < 0.05

50 mM NaCl

100 mM NaCl

150 mM NaCl

200 mM NaCl

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mannitol and grow under light at 25°C with a minimum of four or five replicates. 4. After 6 days, analyze the growth of transgenic and nontransgenic plants growing with and without water deficit or osmotic stress. 5. Take the measurements of the plants as described for saltstress analysis and compare the growth difference between stressed and non-stressed transgenic and non-transgenic plants (Table 3). 6. Transfer the plants to soil mixture composed of 1:1 (v/v) peat/perlite for further growth and development in the green house. 3.7.4. Water Deficit Stress In Vivo

1. Germinate 30–40 seeds (R0, R1, or R2 seeds) on MS7 as described in Section 3.7.1. (see Note 20). 2. Transfer 1-week old growing seedlings with (transgenic) and without selection (non-transgenic control plants) into small pots of 8 cm square containing soil mix composed of 1:1 (v: v) peat and perlite (one plant per pot). 3. Keep the pots in water-filled trays and let them grow in the green house for an additional 2 weeks before starting waterdeficit treatment. 4. Divide the growing plants into two sets (a) watered (nonstressed) and (b) water deficit (stressed); use eight or ten plants for each transgenic line and eight or ten non-transgenic plants for each set. Before starting the stress treatment, take the measurements such as the initial plant height and leaf numbers of each plant. 5. Water the set (a) of plants (non-stressed) continuously from the trays for the whole week. 6. Water the second set (b) of plants (stressed) only for 2 days. 7. After 2 days of watering, completely remove the water from the trays of set (b) to create water-deficit condition for the rest of the week. 8. Repeat this treatment for 5 weeks. 9. At the end, take the measurements of plant height and number of tillers of each plant and compare the growth difference between stressed and non-stressed transgenic and non-transgenic plants. 10. Apply analysis of variance (58) to statistically analyze the data. 11. Separate means by using Tukey’s Studentized range test at 95% confidence level.

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4. Notes 1. Seeds can also be sterilized using 70% ethanol for 2 min; wash with autoclaved distilled water once, and then soak in 50% Clorox bleach for 15 min with constant shaking at 0.5 × g. 2. Before sowing the sterilized seeds on germination medium MS1, remove the excess water from seeds using sterilized filter paper sheets (3 mm; VWR). 3. Shoot apices contain apical meristems, two to three leaf primordial, and leaf bases and can be distinguished as a swollen part close to the leaf base. 4. Use 7–8 dishes for each cultivar. For optimum results working with different cultivars, try different combinations of 2,4-dichlorophenoxyacetic acid (0 and 0.5 mg/l) and N6benzyladenine (0, 0.5, 1.0, 2.0, 4.0, and 8.0 mg/l). 5. To calculate the relative frequency of differentiation of multiple shoots (see (47)). 6. For optimum results using more than one plasmid, try different molar ratios such as 3:1 or 2:1 (for two plasmids) and 1:1:1 or 3:2:1 (for three plasmids) where the plasmid that contains selection marker is the last component of each ratio (i.e. at the lowest concentration). 7. Use 1-month old multiple shoot cultures differentiated from the shoot meristems of multiple shoots for microprojectile bombardment (Fig. 2a); prior to bombardment, physically expose multiple shoot cultures by removal of the coleoptiles and leaves, if necessary. 8. Use a biolistic particle acceleration device (PDS 1000/He, Bio-Rad) for bombardment under 26 mmHg chamber pressure at a distance of 1.5, 2, and 6.5 cm from the rupture disc to the macrocarriers, to the stopping screen, and to the target, respectively, with 1,550 psi helium pressure. 9. After first bombardment, incubate the tissues in the dark at 25°C till second bombardment. 10. Transfer the plants to bigger pots (12 cm square) after 1 month of growth in the green house to enhance the vegetative growth and fertilize the plants weekly if necessary with peters 20:20:20 fertilizer (Griffin Green House and Nursery supplies). 11. Herbicide application: First locally paint the herbicide on the youngest leaf of the three-leaf-stage plants and when it reaches the six-leaf stage, spray the whole plant with 1% Ignite herbicide containing 200 g/l (16.222%) of glufosinate as the active ingredient. 12. Use the progeny (R0, R1, or R2) seeds from different independent transgenic lines. We used the R2 progeny from five

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independent transgenic lines Ogle BRA-82, Ogle BRA-17, Ogle BRA-8, Ogle BRA-19, and Ogle BRA-41 in this experiment. 13. Grow seeds from transgenic R0, R1, and R2 lines on MS7 medium with selection to screen transgenic and non-transgenic plants. Use only those seedlings that grow on selection to further test for stress resistance. 14. Plant fresh weight: take the whole plant out from Magenta carefully without damaging the roots in the medium, wash the medium off from roots, dry the roots using paper towels, and weigh using a weighing balance (Precision Weighing Balances, Bradford, MA 01835). 15. Plant dry weight: Place the whole plant on weighed 3mm filter papers (already dried to constant weight), wrap them inside aluminum foils individually, and place in an oven (VWR) set at 110°C for 24 h. Remove the plants from the oven and keep in a desiccator (VWR) to bring to room temperature and to avoid moisture absorption before weighing. Weigh each plant with the filter paper and calculate individual plant weight after subtracting the filter paper weight. 16. Use mannitol in the culture medium to create water-deficit or osmotic-stress conditions for in vitro grown plants. 17. This is the modified protocol to evaluate green house grown hva1 expressing transgenic lines for water-deficit stress tolerance (48). 18. The R3 progenies of five independently transformed oat lines of the Ogle cultivar (BRA-8, BRA-17, BRA-19, BRA41 and BRA-82) and control plants were grown under green house conditions. These were obtained from R2 plants generated in a previous study (47). 19. Use one set of plants for each saline solution. Water the plants with saline solution once every day to ensure adequate leaching and to prevent salinity excess. 20.Additional measurements can also be taken such as number of tillers/plant, number of days to heading, flag leaf area, 1,000 kernel weight, number of kernels/panicle, panicle length, and number of spikelets/panicle for each plant (48, 51).

Acknowledgements The MSU Plant Breeding and Genetics Program, the Northwest Plant Biotechnology Consortium, and Al-Azhar University, Assiut, Egypt, supported this research.

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Mragowa, Poland. 13–19 June. Kluwer, Amsterdam, The Netherlands. Martin, R. J., Jamieson, P. D., Gillespie, R. N. and Maley, S. (2001) Effect of timing and intensity of drought on the yield of oats (Avena sativa L.). Proceeding of the 10th Australian Agronomy Conference, Hobart. Murty, A. S., Misra, P. N. and Haider, M. M. (1984) Effect of different salt concentrations on seed germination and seedling development in few oat cultivars. Indian Journal of Agricultural Research 18, 129– 132. Verma, O. P. S. and Yadava, R. B. R. (1986) Salt tolerance of some oats (Avena sativa L.) varieties at germination and seedling stage. Journal of Agronomy and Crop Science 156, 123–127. Schonbeck, M. W. (1988) Cover Cropping and Green Manuring on Small Farms in New England and New York: An Informal Survey. Research Report 10, New Alchemy Institute, East Falmouth, MA 02536. Koev, G., Mohan, B. R., Dinesh-Kumar, S. P., Torbert, K. A., Somers, D. A. and Miller, W. A. (1998) Extreme reduction of disease in oats transformed with the 5’ half of the barley yellow dwarf virus PAV genome. Phytopathology 88, 1013–1019. Stoskopf, N. C. (1985) Barley and Oat, in Cereal Grain Crops, (Stoskopf, N. C., ed.), Reston Publishing, Reston, Virginia, pp. 444–458. Cushman, J. C. and Bohnert, H. J. (2000) Genomic approach to plant stress tolerance. Current Opinions in Plant Biology 3, 117–124. Abebe, T., Guenzi, A. C., Martin, B. and Cushman, J. C. (2003) Tolerance of mannitol-accumulating transgenic wheat to water stress and salinity. Plant Physiology 131, 1748–1755. Epstein, E., Norlyn, J., Rush, D., Kingsbury, R., Kelley, D., Cunningham, G. and Wrona, A. (1980) Saline culture of crops: a genetic approach. Science 210, 399–404. Ribaut, J. M. and Hoisington, D. A. (1998) Marker assisted selection: new tools and strategies. Trends in Plant Science 3, 236–239. FAO. (1999). Biotechnology in food and agriculture. http://www.fao.org/unfao/ bodies/COAG/COAG15/X0074E.htm. Sharma, H. C., Crouch, J. H., Sharma, K. K., Seetharama, N. and Hash, C. T. (2002) Application of biotechnology for crop

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improvement: prospects and constraints. Plant Science 163, 381–395. Mazur, B., Krebbers, E. and Tingey, S. (1999) Gene discovery and product development for grain quality traits. Science 285, 372–375. Rines, H. W., Phillips, R. L. and Somers, D. A. (1992) Application of tissue culture to oat improvement, in Oat Science and Technology, (Marshall, H. G., and Sorrels, M. E., eds.), American Society of Agronomy and Crop Science Society, Madison WI, pp. 777–791. Somers, D. A., Torbert, K. A., Pawlowski, W. P. and Rines, H. W. (1994) Genetic engineering of oat, in Improvement of Cereal Quality by Genetic Engineering, (Henry, R. J. and Ronalds, J. A., eds.), Plenum, New York, pp. 37–46. Cummings, D. P., Green, C. E. and Stuthman, D. D. (1976) Callus induction and plant regeneration in oats. Crop Science 16, 465–470. Rines, H. W. and McCoy, T. J. (1981) Tissue culture initiation and plant regeneration in hexaploid species of oats. Crop Science 21, 837–842. Bregitzer, P., Bushnell, W. R., Somers, D. A. and Rines, H. W. (1989) Development and characterization of friable, embryogenic oat callus. Crop Science 29, 798–803. Rines, H. W. and Luke, H. H. (1985) Selection and regeneration of toxin insensitive plants from tissue cultures of oat (Avena sativa) susceptible to Helminthosporium victoriae. Theoretical and Applied Genetics 71, 16–21. Somers, D. A., Rines, H. W., Gu, W., Kaeppler, H. F. and Bush-Nell, W. R. (1992) Fertile transgenic oat plants. Bio/Technology 10, 1589–1594. Zhang, S., Zhong, H. and Sticklen, M. B. (1996) Production of multiple shoots from apical meristems of oat (Avena sativa L.). Journal of Plant Physiology 148, 667–671. Torbert, K. A., Rines, H. W. and Somers, D. A. (1998) Transformation of oat using mature embryo-derived tissue cultures. Crop Science 38, 226–231. Cho, M. J., Jiang, W. and Lemaux, P. G. (1999) High frequency transformation of oat via microprojectile bombardment of seed-derived highly regenerative cultures. Plant Science 148, 9–17. Gless, C., Lorz, H. and Jahne-Gartner, A. (1998) Establishment of a highly efficient regeneration system from leaf base segments of oat (Avena sativa L.). Plant Cell Reports 17, 441–445.

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49. Patnaik, D. and Khurana, P. (2003) Genetic transformation of Indian bread (T. aestivum L.) and pasta (T. durum L.) wheat by particle bombardment of mature embryoderived calli. BMC Plant Biology 3, 1–11. 50. Maqbool, S. B., Zhong, H. and Sticklen, M. B. (2004) Genetic engineering of oat (Avena sativa L.) via the biolistic bombardment of shoot apical meristems, in Transgenic Crops of the World – Essential Protocols, Chap. 5, (Curtis, I. S., ed.), Kluwer, Dordrecht, The Netherlands, pp. 63–78. 51. Oraby, H. F., Ransom, C. B., Kravchenko, A. N. and Sticklen, M. B. (2005) Barley HVA1 gene confers salt tolerance in R3 transgenic oat. Crop Science 45, 2218– 2227. 52. Murashige, T. and Skoog, F. (1962) A revised medium for rapid growth and bioassays with tobacco tissue cultures. Physiology Plant 15, 473–497. 53. Strickberger, M. W. (1985) Genetics, 3rd ed. Macmillan, New York, pp. 126–146.

54. Southern, E. M. (1975) Detection of specific sequences among DNA fragments separated by gel electrophoresis. Journal of Molecular Biology 98, 503–517. 55. Sambrook, J., Fritsch, E. F. and Maniatis, T. (1989) Molecular cloning: A Laboratory Manual, 2nd ed. Cold Spring Harbor Lab, New York. 56. Bradford, M. (1976) A rapid and sensitive method for quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Annals of Biochemistry 72, 248–254. 57. Jafferson, R. A., Kavanagh, T. A. and Bevan, M. W. (1987) GUS fusions: β-glucuronidase as a sensitive and versatile gene fusion marker in higher plants. EMBO Journal 6, 3901–3907. 58. Lamb, C. R. C., Milach, S. C. K., Pasquali, G. and Barro, R. S. (2002) Somatic embryogenesis and plant regeneration derived from mature embryos of oat. Pesquisa Agropecuaria Brasileira 37, 123–130.vv

Chapter 11 Promoter Sequences for Defining Transgene Expression Huw D. Jones and Caroline A. Sparks Abstract The design of reverse genetic experiments that utilize transgenic approaches often requires transgenes to be expressed in a predefined pattern and there is limited information regarding the gene expression profile for specific promoters. It is important that expression patterns are predetermined in the specific genotype targeted for transformation because the same promoter–transgene construct can produce different expression patterns in different host species. This chapter compares constitutive, targeted, or inducible promoters that have been characterized in specific cereal species. Key words: Promoter, regulatory elements, transgene expression, constitutive, tissue specific, inducible, reporter genes.

1. Introduction The classical view of how eukaryote gene expression is regulated states that sequences located on the same DNA molecule (in cis) interact with proteins or RNA encoded by different DNA molecules (trans-acting signals) to control gene expression. These cis elements include the CAAT and TATA boxes, and the response elements of the promoter itself, enhancer elements that may be located far up- or downstream from the coding region, 5′ and 3′ untranslated regions (UTRs), introns, polyadenylation signals, etc. However, this rather simplistic view of regulated gene expression is being constantly challenged and updated, especially from insights into mammalian biology (see for example (1, 2)). Although the full repertoire of signals that regulate gene expression are yet to be fully understood, defined patterns of recombinant gene expression can be achieved using relatively short Huw D. Jones and Peter R. Shewry (eds.), Methods in Molecular Biology, Transgenic Wheat, Barley and Oats, vol. 478 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-379-0_11

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(up to 2 kb) promoter sequence, which can be further enhanced with an intron inserted within the transcriptional unit downstream of the promoter (3– 6). When planning a plant transformation experiment, one of the first challenges is to construct an expression cassette containing the gene of interest and appropriate regulatory sequences to give the desired patterns of gene expression. If the promoter sequence chosen has not previously been tested in the species to be transformed, it is recommended that this be done first, using a scorable reporter gene such as gfp or uidA (GUS). This chapter reviews the various promoter sequences that have been validated in cereals for constitutive, tissue-specific, or inducible expression. Table 1 lists all the promoters known to have been used for reporter gene expression studies in stably transformed wheat, barley, or oats with exemplar references so the reader can seek further details. Some information on promoter function can be inferred from expression of ‘trait’ (non-reporter) or selectable marker genes but these studies were not included in Table 1. In addition, many of the promoter sequences listed have also been used in rice and maize but these species have been covered elsewhere.

2. Constitutive Gene Expression Transformation projects, involving both stable transgenic lines and transient overexpression strategies, often demand high levels of gene expression in all tissues and at all developmental stages. To achieve a generally constitutive pattern of transgene expression in dicot plants, the cauliflower mosaic virus 35S (CaMV 35S) promoter (49, 50) with or without the addition of various upstream activator regions (UAR) is commonly used. The CaMV 35S promoter has also been used in cereals to drive selectable marker and trait genes and there has been some characterization of its expression using reporter genes in rice (51 –53). However, there are conflicting views regarding the merits of CaMV 35S in wheat, barely, or oats, with some authors claiming lower levels of marker gene expression and more frequent inactivation compared to monocot promoters (12, 54, 55). Other viral promoter sequences that have been tested in cereals include those obtained from the rice tungro bacilliform and cestrum yellow leaf curling viruses (RTBV and CmYLCV, respectively). Deletions of the RTBV promoter have been described as vascular- or phloem specific when used in transgenic rice (56–61). In transgenic wheat, it drives strong GUS expression in a range of tissues but

Act1

pAHC25 (10, 11) Maize Ubi1 plus Ubi1 intron:: uidA (Gus)::nos 3′

Ubi1

pRC-62 (16, 23) Rice Actin1D plus first in/ex:::uidA(Gus)/nptII::nos 3′

pAct1sGFP-1 (19) Rice Actin 1 plus first in/ex::sgfp(S65T)::nos 3′

pACT1-F (16) Rice Actin plus first in/ ex::uidA (Gus)::nos 3′ pDB1 (17) Rice Actin plus first in/ ex::uidA (Gus)::nos 3′

pAHC15 (11) Maize Ubi1 plus Ubi1 intron:: uidA (Gus)::nos 3′

pBargus (7) Maize Adh1 plus Adh1 intron::uidA (Gus)::nos 3′

Adh1

Promoters with generally constitutive expression patterns

Plasmid name (ref.) details of promoter/reporter gene/ Promoter terminator cassette Expression reported

GUS expression reported in leaves

(12)

(9)

(8)

Ref.

Oat (var. Melys); stable transgenics using bombardment of regenerable callus Oat (var. Jumbo); stable transgenics from bombardment of leaf bases Wheat (var. Florida) stable transgenics using bombardment of immature scutella Oat (var. Garry); stable transgenics using bombardment of regenerable callus Wheat (cv. Fielder) stable transgenics using bombardment of immature scutella Barley (cv. Golden Promise); stable transgenics using bombardment of immature scutella Wheat (cv. Fielder) stable transgenics using bombardment of immature scutella)

(24)

(22)

(21)

(20)

(17)

(12) (18)

(continued)

GUS expression reported in leaves Histochemical GUS activity reported in pollen, floral organs and leaves Histochemical GUS activity reported in leaves GFP expression reported in SMCs, anthers, ovary, stigma and seeds GFP expression reported in regenerable callus, leaves, developing shoots and embryos GFP expression reported in callus, pollen, ovary, stigma, root, immature embryo and endosperm Histochemical GUS activity reported in leaves, ovary, stigma, anthers and pollen

(13) Silencing of the uidA (Gus) and bar genes was reported due to methylation and chromatin condensation (14) Constitutive histochemical GUS expression Wheat (NILs L88-6 and L8831); stable reported, generally strong in young, metatransgenics using bombardment of immature bolically active tissues and in pollen grains scutella Oat (lines derived from GAF-30/Park); stable trans- Histochemical GUS activity reported in regen- (15) erative callus, anther, stigma and leaves genics using bombardment of regenerable callus

Oat (var. Melys); stable transgenics using bombardment of regenerable callus Barley (cv. Golden Promise); stable transgenics using bombardment of immature scutella

Oat (lines derived from GAF-30/Park); stable trans- GUS expression reported in PPT-resistant calli, and seeds genics using bombardment of regenerable callus Wheat (cv. Parvon and RH770019); stable trans- GUS expression reported in PPT-resistant calli, root tips and seeds genics using bombardment of regenerable callus

Species and mode of transformation

Table 1 Promoter expression validated using reporter genes in stably transformed wheat, barley or oats with exemplar references

Promoter Sequences for Defining Transgene Expression 173

pHP12679 Maize histone H2B plus Ubi1 intron 1:: uidA(Gus)::pinII 3′

H2B

(28) Histochemical GUS activity reported in endosperm and absent in leaves, roots and flowers Highly expressed in endosperm at early to mid-maturation stages

Tissue-specific GFP expression reported in endosperm only

Wheat (cv. Bobwhite); stable transgenics using bombardment of immature scutella

Barley (cv. Golden Promise); stable transgenics using bombardment of immature scutella

B- and pHorB-Gus and pHorD-Gus D-hor- Barley B- and D-type hordeins (1,043 and dein 834 bp, respectively:: uidA (Gus)::nos 3′ pD11-Hor3 (29) p16 Barley B1-hordein (550 bp):: uidA(Gus):: nos 3′ and Barley D-hordein (434 bp):: uidA(Gus)::nos 3′ pDhsGFP-1 Barley D-hordein:: sgfp(S65T)::nos 3′

(22)

(30)

(27)

Endosperm specific, on ~14 dpa, longer promoter stronger

Wheat (cv. Bobwhite); stable transgenics using bombardment of immature scutella

pLMWG1D1-326 pLMWG1D1-938 Wheat low molecular weight Glutenin subunit LMWG1D1::nos 3′ (two lengths −326 to +30, and −938 to +30)

LMWG

(25)

(12)

Ref.

(26)

Constitutive. Gus expression reported in all tissues tested

No expression reported from CaMV 35S promoter

Expression reported

Wheat (durum var. Ofanto); stable transgenics Endosperm specific 10–12 dpa using bombardment of immature inflorescences

pHMW-GUS Wheat high molecular weight glutenin subunit Glu-1D-1:: uidA (Gus)::nos 3′ (−1,191 to +58)

HMWG

Promoters with seed-specific expression patterns

Oat (var. Melys); stable transgenics using bombardment of regenerable callus

pJIT102 CaMV 35S:: uidA (Gus)::CaMV 35S 3′

CaMV 35S Wheat (var. Cadenza); stable transgenics using bombardment of immature scutella

Species and mode of transformation

Plasmid name (ref.) details of promoter/reporter gene/ Promoter terminator cassette

Table 1 (continued)

174 Jones and Sparks

pRSHOG and deletions Oat globulin::GFP::rbcS 3′

AsGlo1

pBSD5sGFP Barley Lem1:: sgfp(S65T)::nos 3′

p1414 and various deletions (39) barley Lem2:: sgfp(S65T)::nos 3′

Lem1

Lem2

Promoters with other expression patterns

pEvec202Em.gfpnos Wheat Em (646 bp Early methionine):: sgfp(S65T)::nos 3′

Em

Barley (cv. Golden Promise); stable transgenics using bombardment of immature scutella

Wheat (cv. Bobwhite); stable transgenics using bombardment of immature scutella

Barley (cv. Golden Promise); stable transgenics using Agrobacterium co-cultivation of embryogenic callus

(38)

(37)

(continued)

Strong cell- and tissue-specific GFP expression (40) reported in lemma/palea, glumes, coleoptile, auricle, ligule and, unexpectedly, epicarp

Tissue-specific expression of GFP reported in outer floret organs at anthesis

Endosperm-specific GFP expression reported

(36)

(35)

Barley (cv. Golden Promise); stable transgenics Tissue-specific GFP expression reported using Agrobacterium co-cultivation of embryoin various tissues of the developing and genic callus mature grain but not roots or leaves. Some regulation by GA

pA57 Barley bi-functional alpha-amylase/subtilisin inhibitor (1033-asi promoter):: sgfp(S65T)::nos 3′

asi

Barley (cv. Golden Promise); stable transgenics Tissue-specific GFP expression reported in using Agrobacterium co-cultivation of embryoembryo, aleurone and junction between genic callus endosperm transfer cells and aleurone layer. Differences in temporal and spatial GUS expression between barley and rice

(33)

(31)

Histochemical GUS reported in scutellum and embryonic axis of seeds at 4 days post germination

Wheat (var. Cadenza and durum var. Ofanto); GUS expression reported in starchy stable transgenics using bombardment of immaendosperm cells only ture inflorescences and scutella Wheat (var. Cadenza); stable transgenics using bombardment of immature scutella

Pina::uidA Pinb::uidA(32) Wheat puroindoline a and b::uidA (Gus):: nos 3′

αamy1 and pα1GT αamy2 pα2GT (34) Wheat alpha amylase 1 or 2:: uidA (Gus):: nos 3′

Pin A and Pin B

Promoter Sequences for Defining Transgene Expression 175

pPsEND1gusA (47) pea END1(2,731 bp):: Wheat (cv. Bobwhite); stable transgenics using uidA (Gus)::nos 3′ bombardment of immature scutella

End1

GUS expression localized to pollen; microspores binucleate and pollen tube stages

pCoYMV-GUS (45) Commelina yellow Oat (lines derived from GAF-30/Park); stable Gus localized in vascular tissue of shoots, mottle virus (−1,026 to +12) plus Adh1 transgenics using bombardment of regenerable leaves and floral bracts and in roots. Ovaintron:: uidA (Gus)::nos 3′ callus ries stained intensely and scutellum had expression but no staining in anthers or endosperm

CoYMV

(48)

(46)

(44)

Oat (cv. GAF-30/Park), barley (cv. Golden Prom- GUS expression reported in various tissues with differences between species ise) Wheat (cv. Bobwhite) stable transgenics using bombardment of embryo-derived calli or immature embryos

pSCBV-3m (43) Sugarcane bacilliform virus plus maize Adh1 leader and first intron:: uidA (Gus)::nos 3′

(41)

Tissue-specific expression of GUS reported in endosperm and aleurone

SCBV

(42)

Ref.

Tissue-specific expression of GUS reported in endosperm, pollen and carpel

Wheat (cv. Chinese Spring) stable transgenics using bombardment of embryo-derived calli

Expression reported

pAGP2::GUS (41) wheat ADP-glucose pyrophosphorylase large subunit:: uidA (Gus)::nos 3′

Species and mode of transformation

AGPL1

Plasmid name (ref.) details of promoter/reporter gene/ Promoter terminator cassette

Table 1 (continued)

176 Jones and Sparks

Promoter Sequences for Defining Transgene Expression

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not pollen nor roots (authors’ unpublished work). The CmYLCV promoter demonstrated strong, constitutive GUS or GFP expression in Arabidopsis, tobacco, maize, and rice (62). Two monocot promoters that are often used in preference to viral sequences for cereal transformation experiments are the maize ubiquitin promoter plus first exon and first intron (Ubi1) (10, 11) and the rice actin promoter and intron (Act1) (63). The Maize Ubi1 promoter driving the uidA (GUS) reporter gene has been examined in rice (64, 65), maize (66), and wheat (3, 14) and gave strong GUS expression in most tissues tested. Expression was found to be generally stronger in young, metabolically active tissues and pollen but low or absent in older leaves and roots, and somatic tissue of anthers (see Fig. 1a and b). It has also been reported to be induced by heat shock and stress (64), although there is also a high basal level of expression and most reports of maize Ubi1 in transgenic experiments do not actively utilize this inducibility. The heat shock elements in maize Ubi1 have been identified and, when replaced with a basic domain/leucine zipper factor binding site from a pea lectin promoter, shifts the

Fig. 1. Gus expression driven by various promoters in tissues of stably transformed wheat plants: (a) Expression from maize Ubiquitin1 promoter + intron in Cadenza leaf, (b) Cadenza anther showing gus stained pollen with maize Ubi1 promoter + intron, (c) expression driven by rice Actin 1 promoter in Cadenza stem and immature inflorescence, (d) Ofanto mature half seed showing endosperm specific expression from wheat high molecular weight (HMW) glutenin Glu-1D-1 promoter, (e) Ofanto mature half seed showing wheat puroindoline b (pinb) expression in starchy endosperm, and (f) Cadenza half seed showing localized expression from maize Glb1 promoter in transfer aleurone cells. (See Color Plate 6 )

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balance of seed expression away from the embryo and towards the starchy endosperm (66). Other ubiquitin promoters have also been characterized in monocot transformations including RUBQ2 and Rubi3 from rice (67, 68) and ubi4 and ubi9 from sugarcane (69), all of which give comparable or higher expression levels than maize Ubi1 or CaMV 35S. Maize Ubi1::GUS expression has also been shown to produce good expression in roots of transgenic Arabidopsis, tomato, tobacco, and pine (70). A good alternative to ubiquitin is the promoters from actin genes. A rice actin promoter (Act1-D) comprising 2.1 kb upstream of the Act1 translation initiation codon contains all the necessary 5′ regulatory elements for high-level GUS expression in transiently transformed rice and maize (16, 63) and in stably transformed rice (71). In addition to the promoter region itself, the Act1-D sequence also possessed a 5′ intron and a portion of the first exon of the Act1 gene. Act1-D::GUS expression was reported to be constitutive throughout the sporophytic and gametophytic tissues of the transgenic rice lines (71). Quantitative analysis showed that GUS protein represented as much as 3% of the total soluble protein (71). In wheat, this promoter drives strong constitutive expression in all major tissues at all stages of development (authors’ unpublished data). Rice Act1-D and a truncated version of the Act1 promoter (Act1F) have both been used to drive uidA in transgenic oat (12, 18), but detailed analysis of these lines has not been reported. Other broadly constitutive promoters that have been used and characterized in cereals include the maize H2B promoter with the Ubi first intron (25), the maize Adh1 promoter (8, 9), the rice cytochrome-c promoter (OsCc1) (72), and a range of regulatory sequences from the subterranean clover stunt virus (SCSV) (73).

3. TissueSpecific Gene Expression

The natural expression of most plant genes shows some organ- or tissue-specific pattern depending on the role of the gene product. The broad spatial pattern of gene expression is often preserved when a 1- or 2-kb promoter is used to construct a heterologous expression cassette. For example, a 1.2-kb promoter sequence from the wheat high-molecular-weight glutenin subunit gene, Glu-1D-1, gives starchy endosperm-specific expression of the uidA reporter gene in transgenic durum wheat (26) (see Fig . 1d). This and other glutenin promoters have been used to drive heterologous gene expression in wheat endosperm (reviewed in (74)). Other seed-storage promoters that have been validated include

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the low-molecular-weight glutenin subunit promoter (27) and the B- and D-hordeins (22, 28, 30). Seed-specific expression patterns using transgenic reporter genes have also been reported for Em, asi, αamy1 and 2 promoters, and puroindoline a and b (see Fig . 1e; (31, 33, 35, 36) ). In addition, unpublished work of the authors has demonstrated aleurone and embryo expression patterns for globulin promoter sequences (see Fig . 1f).

4. Inducible Gene Expression The ability to switch candidate gene expression on and off by applying an external stimulus would be invaluable for many research applications (reviewed in (75–77)). Inducible promoters also provide pseudo tissue specificity when the activating stimulus can be applied to particular tissues only. Many inducible promoters have been investigated in transgenic plants including alcR (alcohol inducible (78–81)), glutathione S-transferase/In2-2 (safener inducible (82, 83) ), various heat shock inducible polygalacturonase-inhibiting proteins (84–87) (HSPs, wounding inducible (88)), TetR and tRA (tetracycline-induced and repressed, respectively, (89–93)), EcR (ecdysone indicible (94–97)). However, most of this work was done in dicot plants, and while some information may be transferable to monocots, there is a real need to validate the best of these inducible systems in transgenic cereals.

Acknowledgments Rothamsted Research receives grant-aided support from the Biotechnology and Biological Sciences Research Council (BBSRC) of the UK.

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and olfactory receptor choice. Cell 126, 403–413. 3. Salgueiro, S., Pignocchi, C. and Parry, M. A. J. (2000) Intron-mediated gusA expression in tritordeum and wheat resulting from particle bombardment. Plant Mol. Biol. 42, 615–622.

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Chapter 12 Down-Regulation of Gene Expression by RNA-Induced Gene Silencing Silvia Travella and Beat Keller Abstract Down-regulation of endogenous genes via post-transcriptional gene silencing (PTGS) is a key to the characterization of gene function in plants. Many RNA-based silencing mechanisms such as post-transcriptional gene silencing, co-suppression, quelling, and RNA interference (RNAi) have been discovered among species of different kingdoms (plants, fungi, and animals). One of the most interesting discoveries was RNAi, a sequence-specific gene-silencing mechanism initiated by the introduction of doublestranded RNA (dsRNA), homologous in sequence to the silenced gene, which triggers degradation of mRNA. Infection of plants with modified viruses can also induce RNA silencing and is referred to as virus-induced gene silencing (VIGS). In contrast to insertional mutagenesis, these emerging new reverse genetic approaches represent a powerful tool for exploring gene function and for manipulating gene expression experimentally in cereal species such as barley and wheat. We examined how RNAi and VIGS have been used to assess gene function in barley and wheat, including molecular mechanisms involved in the process and available methodological elements, such as vectors, inoculation procedures, and analysis of silenced phenotypes. Key words: Gene silencing, dsRNA, RNA interference, virus-induced gene silencing, hairpinRNA.

1. Introduction Post-transcriptional gene silencing (PTGS), generally termed RNA silencing, is the plant-based down-regulation of an endogenous gene caused by the introduction of a homologous double-stranded RNA (dsRNA), transgene, or virus. In PTGS, the transcript of the silenced gene is synthesized but does not accumulate because it is rapidly degraded (1, 2). RNA interference (RNAi) is the process whereby dsRNA directly induces the

Huw D. Jones and Peter R. Shewry (eds.), Methods in Molecular Biology, Transgenic Wheat, Barley and Oats, vol. 478 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-379-0_12

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homology-dependent degradation of specific mRNA (3–5). The discovery and molecular understanding of RNAi is one of the most important technological breakthroughs in modern scientific history. This phenomenon was first unknowingly observed over a decade ago when RNA was shown to inhibit protein expression in plants and fungi by processes then known respectively as co-suppression and quelling (6, 7). Co-suppression is the simultaneous reduction in expression of a transgene and homologous endogenous genes. These initial observations suggested that the underlying mechanism was post-transcriptional and sequence specific. The real breakthrough came in 1998 when Fire, Mello, and colleagues first discovered that double-stranded RNA was the source of sequence-specific RNA degradation in Caenorhabditis elegans (3). For this landmark discovery, Fire and Mello were honoured with the 2006 Nobel Prize in Physiology or Medicine. It was later confirmed that dsRNA is also the effective trigger of PTGS in plants and of quelling in fungi (8, 9). The independent parallel discoveries of dsRNA as a trigger of RNA silencing in worms, plants, and fungi suggested a common underlying mechanism. Within a few years, it was demonstrated that they all require several similar genes for dsRNA-mediated silencing (for a review, see (10)). PTGS can be induced in plants by viral vectors harbouring specific genes, through the virus-induced gene silencing (VIGS) system (11), by inverted repeat transgenes producing hairpin transcripts (hairpin RNAs, hpRNAs) (12), by antisense RNA technology (13), or by gene over-expression (co-suppression) (6). Further studies in plants and invertebrate animals demonstrated that the actual molecules that lead to RNAi are small double-stranded RNAs (small interfering RNAs, siRNAs), 21–22 nucleotides in length with characteristic 3′ dinucleotide overhangs (4, 14, 15), which are processed in the cytoplasm by an enzyme called Dicer (15). Dicer recognizes dsRNA as aberrant and cleaves it from both ends into siRNAs (4). There are differences in siRNA-mediated RNA silencing pathways between plants and animals. For instance, two distinct short (21–22nts) and long classes (24–26nts) of siRNAs were shown to accumulate in plants (16), while only the short class of siRNAs accumulated in animals (4). Through sequence complementarity, siRNAs in association with an RNA-induced silencing complex (RISC) directs the cleavage of endogenous RNA transcripts, resulting in gene silencing. Small interfering RNAs are also responsible for amplifying the silencing signal by priming endogenous RNA which can be converted to dsRNA by the action of RNA-directed RNA polymerase (RdRP) encoded in the plant genome (17, 18). Thus, a few trigger dsRNA molecules are sufficient to inactivate a continuously transcribed target mRNA for long periods of time. The inactivation persists through cell division, spreads to untreated

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cells and tissues of plants, and is inherited to subsequent generations. The mechanism of RNAi is schematically summarized in Fig.1. RNAi has advantages over antisense-mediated gene silencing and co-suppression in terms of its efficiency, stability, and the shorter time needed to screen the targeted plants (12). The resulting silencing is almost as complete as that achieved in a gene knockout approach. dsRNA-triggered RNAi directly bypasses the requirement for dsRNA synthesis via a plant-encoded RNA-directed RNA polymerase (RdRP), which is probably the rate limiting step in antisense suppression and co-suppression. In co-suppression, plant transgenes that have integrated as inverted repeats (complex integration patterns) and are exhibiting RNA silencing can also produce dsRNA by read-through transcription from one T-DNA copy into another. Instead, engineered transgenes with transcribed inverted repeats homologous to the target RNA produce dsRNA by intramolecular base pairing and they have been shown to be much more potent at silencing gene expression than either a sense or an antisense transgene alone (8). Such constructs producing self-complementary transcripts are now widely used for functional genomics in plants (12, 19, 20). Since RNAi silencing is often genetically stable, its effects can be studied in progeny (21). Stoutjesdijk et al. (20) in Arabidopsis and Travella et al. (22) in wheat have clearly demonstrated that RNAi constructs

Fig.1. Schematic model of the RNAi mechanism. The cellular enzyme Dicer cleaves dsRNA into 21–25 nucleotide siRNAs. The siRNAs are incorporated into the RNA-induced silencing complex (RISC), which uses the antisense strand of the siRNA to find and destroy the target mRNA. siRNAs can also be used as primers for the generation of new dsRNA by RNA-dependent RNA polymerase (RdRP), thereby amplifying the silencing signal.

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are able to generate phenotypic changes that are inherited stably over several generations, making this approach a reliable tool not only for functional genomics but also for the genetic modification of agronomically interesting traits. For example, RNAi was used to demonstrate that reduction of VRN2 and VRN1 transcript levels, respectively, accelerated and delayed flowering initiation in winter wheat (23, 24). RNAi also offers advantages over mutation-based reverse genetics in its ability to silence one, several, or all members of a multigene family or homoeologous gene copies in polyploids by targeting sequences that are unique or shared by several genes (22, 25, 26). In addition, in contrast to insertional mutagenesis, RNAi offers the possibility of directing silencing in specific tissues depending on the promoter used to direct expression of the hpRNA. Similarly, systems to deliver inducible RNAi offer the advantage of silencing gene expression at specific developmental stages, because they provide flexibility for the timing and the degree of gene inactivation and have the potential for reversal of silencing by withdrawal of the inducer (27, 28). RNAi can be delivered into cereals by transient or stable methods via particle bombardment, Agrobacterium infiltration, or viral infection. In wheat and barley, it was shown that delivery by particle bombardment of specific dsRNA into single epidermal cells transiently interfered with gene function (1, 29, 30). The other approach to efficiently silence genes in cereals is VIGS (31). In this approach, target genes can be transiently inactivated by infecting the plants with a recombinant virus that expresses fragments of the endogenous plant gene transcripts (32–34). Viral RNA produces dsRNA during its replication. VIGS is particularly useful in plant species that are time consuming to transform or when analysing genes that are essential, either for housekeeping functions or for embryonic development (which cause embryonic lethality when knocked out), as the infectious transcripts can be applied to mature plants. VIGS is also well suited to the highthroughput analysis of genes, as VIGS vectors can be delivered by simply rubbing an infectious transcript onto a plant, although the cost of transcribing each construct in vitro might be prohibitively expensive on a large scale. Many viruses have been used as vectors for VIGS in dicotyledonous plants (32, 35, 36). However, VIGS in monocotyledons has been limited to only one virus, the Barley stripe mosaic virus (BSMV), in two hosts, barley and wheat (33, 34, 37). The VIGS system was optimized in these studies by silencing phytoene desaturase (PDS) expression, which provides a convenient visual reporter for silencing with photobleached leaves. Recently, VIGS was used in barley to demonstrate that HvWRKY1 and HvWRKY2 transcription factors act as repressors of basal defense to virulent Blumeria graminis (38). Recently, Ding et al. (39) reported the cloning and modification for VIGS of a virus (Brome mosaic virus, BMV) from Festuca arundinacea

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Schreb. (tall fescue) that causes systemic mosaic symptoms on barley, rice, and maize under greenhouse conditions. Additionally, this virus has no known insect vector, nor is it transmitted through seeds, increasing the safety of this system. Despite the described advantages of single-cell RNAi and VIGS, it is clear that the analysis of whole-organism gene function or, in case of a need of stable genetic changes in plants, RNAi-induced gene silencing system is preferable. Both RNAi- and VIGS-containing transformation cassettes are increasingly being used for reverse genetics as part of an integrated approach to determining gene function. They have proven to be very efficient in interfering with gene expression in various plant systems including cereals such as rice (26), barley (29, 30, 33), and wheat (22, 34, 40). There are no reports yet on RNA silencing in oat. The first large-scale efforts to use RNAi as a tool for plant functional genomics were specifically in Arabidopsis and maize, with over 100 genes targeted for silencing (41). Central to the effort has been the design and the construction of publicly available plasmid vectors that facilitate the stable chromosomal integration of transgenes that trigger RNAi. In this chapter, we will describe the different methodologies currently used with RNAi and VIGS to knock down gene expression in barley and wheat. Although RNA gene silencing is a widely used technology, relatively little is known about the optimal application of this technique in plants. There are many potential factors that can contribute to a given sequence’s ability to cause RNA-induced silencing of an endogenous target. The “21-bp rule” (5, 42) is commonly used to predict whether a given inverted repeat (IR) will target a given gene, meaning that 21bp of 100% identity should be enough to trigger silencing. Travella et al. (22) demonstrated in hexaploid wheat that RNAi silencing had the same quantitative effect on all three homoologous genes, which are known to share up to 99% identity at the nucleotide sequence level in the coding regions (43). However, McGinnis et al. (41), who worked in maize with families of chromatin-related genes sharing sequence similarity, recently showed that the steady-state RNA abundance of the genes homologous to the IR sequence was not reduced, even though these targets shared more than 90% identity with their respective IR sequences and had at least three 21-bp regions of perfect identity with the IR. This observation suggests that sequence identity is not the only determinant of whether an IR sequence can induce degradation of target gene mRNA and that other factors play a role. Additional analysis will be required to determine what characteristics can be used to model and predict the silencing effectiveness of a given IR sequence in plants.

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2. Materials 2.1. RNAi 2.1.1. Generation of Hairpin-RNA Vectors (hp-RNA)

1. Polymerase chain reaction (PCR) primers to amplify approximately 300–600bp of the target sequence with BamHI and BglII restriction sites (when using pAHC17 vector from (44)) appended to the 5′ end of the forward and reverse primers, respectively 2. PCR primers to amplify approximately 300–600bp of the intron sequence cloned between the antisense and sense sequences of the target gene in the hpRNA construct. In this case, BglII and BamHI restriction sites are appended to the 5′ end of the forward and reverse primers, respectively, for directional cloning (see Note 1) 3. pAHC17 vector (44) containing the maize ubi-1 promoter and the nopaline synthase (nos) terminator (modify restriction sites on the primers if you are using other vectors such as pStarling and pStargate, http://www.pi.csiro.au/RNAi/vectors.htm) 4. pGEM-T vector (Promega) to clone the PCR product 5. PCR purification kit or columns (Promega or others) 6. DNA template (20ng) 7. Primers (10µM each) 8. dNTP mixture (10mM) 9. Taq DNA polymerase (Sigma) and its own buffer 10. Appropriate restriction enzymes (from companies such as Promega, NEB, etc.) 11. LB medium (liquid and solid) with appropriate antibiotics

2.1.2. Generation of RNAi Transgenic Plants

1. Choose the appropriate transformation systems (particle bombardment or Agrobacterium transformation) (see General Introduction). 2. The hp-RNA constructs are co-transformed with a plasmid containing the selectable marker of choice (see Chapter 2) (see Part II. Transformation and Regeneration). 3. Regeneration and selection of the transformed plants are performed essentially as described elsewhere in this volume (see Chapters 4–10) (see Part II. Transformation and Regeneration). 4. Molecular characterization of transgenic plants (see Chapters 13 and 14) (see Part IV. Characterisation of Transgenic Plants).

2.2. Vigs 2.2.1. Generation of BSMV-Derived Vectors

1. For the VIGS experiments, the γRNA-based BSMV vectors (37) are used to silence genes in barley and wheat (see Note 2 and Fig.2). 2. PCR primers are used to amplify approximately 300–600bp of the target sequence with NotI and PacI restriction sites

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Fig.2. Genomic organization of Barley Stripe Mosaic Virus (BSMV). Tripartite genome composed of the α, β, and γ RNAs. The fragment homologous to the target gene is cloned downstream of the termination codon of the γb open reading frame. The positions of selected restriction sites are indicated.

appended to the 5′ end of the forward and reverse primers, respectively. 3. Digestion of PCR products with NotI and PacI and insertion into the γ.bPDS4 (37); PDS for phytoene desaturase) vector digested with NotI–PacI are carried out. 2.2.2. In Vitro Transcription of Viral RNAs

Capped in vitro transcripts are prepared from three linearized plasmids (pBSMVα with MluI, pBSMVβ with SpeI, and pBSMVγ. b+fragment homologous to target gene with BssHII) that contain the tripartite BSMV genome using the mMessage mMachine T7 in vitro transcription kit (Ambion), following the manufacturer’s protocol.

2.3. Quantitative RealTime PCR

1. Isolation of total RNA from leaves or other tissue materials using TRIzol reagent (Invitrogen Life Technology). Apply the usual rules of RNA handling and avoid ribonuclease contamination (see Note 3) 2. Reverse transcription (RT) with 10µg of total RNA 3. For RT, 0.07µg of oligo(dT)21 primers 4. Reverse transcriptase (7 units) (Invitrogen Life Technologies, Basel, Switzerland) and 1× own buffer 5. dNTPs (0.7mM of each) 6. dTT (10mM) and 1.5 units of RNase OUT (Invitrogen Life Technologies, Basel, Switzerland)

2.4. Short Interfering RNA Detection

1. Recover the lower-molecular-weight RNAs after removal of high-molecular-weight RNAs by precipitation with 10% polyethylene glycol 8000-0.5 M NaCl (14) (see Note 4).

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2. Dry 7µg of the lower-molecular-weight RNA fraction using a speedvac and re-suspend the pellets in 10µl of loading buffer (95% formamide, 20mM EDTA pH 8.0, 0.05% bromophenol blue, 0.05% xylene cyanol). 3. Heat RNA samples at 95°C for 5min and chill on ice prior to loading. 4. Load RNA samples on gel (15% polyacrylamide, 7M urea, 1×TBE pH 8.5), and run at 260V – 47mM for a 20-cm gel (see Note 5). 5. Transfer gel to Hybond N+ membranes (Amersham Biosciences) by electroblotting in 25mM sodium phosphate buffer, pH 6.5, at 15V/200mA, overnight. Cross-link membranes with UV (1,200–2,400µJ). 6. For hybridization, the ULTRAhyb-Oligo buffer from Ambion (17× buffer) can be used. It needs to be dissolved at 68°C before using. 7. Label 1µM of around ten DNA oligos complementary to the sequence of interest using 0.5 units of T4 Polynucleotide Kinase (PNK, Roche) and 6µl of [γ-32P]ATP (5,000Ci/ mmol). 8. Carry out RNA blot hybridizations at 35°C as described by Hamilton and Baulcombe (14).

3. Methods 3.1. Parameters for Selecting a Target Gene Fragment 3.1.1. Sequence

1. The starting point for generating a given RNA-silencing vector is a bioinformatics effort. By using known cDNA sequences or predicted gene sequences corresponding to the target gene of interest, primers are designed to amplify a portion of the cDNA by using reverse transcription-PCR (RT-PCR). When genomic sequences are not available for the genes of interest, we can use known target genes from other organisms to search through the available sequenced cDNAs. If the target gene is a member of a multi-gene family, multiple alignments of family members are needed to help guide the design of PCR primers. The region of the gene to be amplified and its similarity to other genes dictates whether the resulting RNAi construct is likely to target a single mRNA or transcripts of multiple related genes. 2. Both translated as well as untranslated regions (UTRs) have been used with equally good results. As the mechanism of silencing depends on sequence homology, there is potential for cross-silencing of related mRNA sequences. Where this is

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not desirable, a region with low sequence similarity to other sequences, such as 5′ or 3′ UTR, should be chosen. To reduce cross-silencing, blocks of sequence with identity over 20 bases between the construct and non-target gene sequences should be avoided. 3. Standard software can now be used for improving detection of sequence identity in order to accurately and systematically evaluate and minimize RNAi off-target effects between siRNA sequences and target genes (45) (see Note 6). 4. Gene suppression can also be achieved by expressing dsRNAs derived from promoters, rather than coding regions of genes. This will therefore induce transcriptional gene silencing (TGS) (46, 47). 3.1.2. Size

Gene fragments ranging from 50 bp to 1kb have been successfully used as targets. Two factors can influence the choice of length of the fragment. Shorter fragments result in a lower frequency of silencing, and very long hairpins increase the chance of recombination in bacterial host strains. We recommend a fragment length of 300–600 bp as a suitable size to maximize the efficiency of silencing obtained.

3.2. RNAi

There are at least three ways in which hpRNA constructs can be made. The construct may be generated from standard plant transformation vectors in which the hairpin-encoding region is generated de novo for each gene. Alternatively, generic gene silencing vectors such as the pStarling and pStargate series developed by CSIRO (Australia) for cereal transformation (http://www.pi.csiro.au/RNAi/vectors.htm) can be used. They simply require the insertion of PCR products, derived from the target gene, into the vectors by conventional cloning (pStarling) or by using the Gateway directed recombination system (pStargate).

3.2.1. Generation of Hairpin-RNA Vectors (hp-RNA)

Clone cDNA Fragments Corresponding to Targeted mRNAs

1. Fragments of around 300–600bp corresponding to the target sequence are isolated by RT-PCR using specific primers with incorporated BamHI and BglII restriction sites, which produce ends compatible with each other. This allows the gene fragments to be directionally cloned within the unique BamHI site of vector pAHC17 (44). 2. The intron fragment is produced with the same cloning strategy as described for the target genes. 3. PCR amplification is carried out for 35 cycles of 45s denaturation at 94°C, 45s annealing at 62°C. and 90min extension at 72°C. 4. The cDNA fragments recovered by BamHI and BglII digestion are cloned in the BamHI unique restriction site of plasmid pAHC17 (see Note 7 and Fig.3).

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Fig.3. Self-complementary hairpin (hp) construct derived from hp transgene used in the transformation experiments. Gene-specific sequences (black arrows indicating the orientation) in the antisense and sense orientations are cloned between fragments of an intron sequence and are controlled by the constitutive ubi promoter (hatched box) and the nopaline synthase terminator (dotted box).

5. Once the assembly of the inverted repeat is completed, it can then be cloned into an appropriate binary vector in case the Agrobacterium transformation is used. The pSTARLING Vector

1. The pSTARLING system is found to work very efficiently and is suitable for silencing a small number of genes, because it becomes soon laborious when individually silencing a large number of target genes. 2. This vector uses the maize ubiquitin promoter for high-level constitutive hairpin-RNAi production in monocot plants. 3. A PCR fragment can be inserted, using conventional restriction enzyme digestion and DNA ligation techniques, in the antisense orientation into the BamHI.PacI.AscI polylinker and in the sense orientation into the SpeI.SnaBI.KpnI polylinker.

The pSTARGATE Vector

1. This vector was designed as a high-throughput alternative to the pSTARLING vector using the commercially available Gateway cloning system (http://www.Invitrogen.com), in order to silence large sets of genes (i.e. members of a gene family or pathways). 2. As well as allowing directional cloning, the system incorporates a negative selection marker (ccdB) that selects against vectors that have not undergone a recombination reaction, resulting in a high frequency of recovery of recombined plasmids. 3. The pSTARGATE vector contains two recombination cassettes consisting of either attP1-ccdB-attP2 or attR1-ccdB-attR2 in an inverted repeat configuration, such that when gene fragments flanked by the appropriate att sites are recombined with the vector, an ihpRNA-encoding construct is produced.

Verification of the Engineered Plasmid Vector

1. All amplified cDNA fragments are sub-cloned into pGEM-T vector. 2. Transformation of E. coli is performed following each ligation reaction with selection of ampicillin-resistant colonies, preparation of plasmid DNA from small-scale cultures, and identification of recombinant plasmids by restriction mapping. 3. To be sure that the correct fragment of DNA has been cloned, which is a potential problem in a large project in which numer-

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ous cDNA fragments are being cloned in parallel and mixups could occur, DNA sequencing is used to verify each final vector. 4. Each construct is finally verified by sequencing the entire inverted repeat fragment and aligning the resulting sequence against the target gene sequence. Detection of Silencing Phenotypes

1. Transforming plants with hpRNA constructs typically generate a series if independent lines have different phenotypes and degrees of target mRNA reduction (22, 48). 2. Levels of the targeted mRNA can range from wild type to undetectable or with a full spectrum of the effect of RNAi (weak, intermediate, and strong) (see Note 8).

3.3. Vigs 3.3.1. VIGS Inoculation Protocol

1. Transcripts of each of the BSMV genomes (wild-type or genetically modified) are mixed in a 1:1:1 ratio and combined with the inoculation buffer FES (49). 2. Apply the mixture to 7-day-old plants by rub inoculation. Pipette the mixture between the pinched thumb and first finger of a gloved hand. 3. Hold the base of the plant with the other hand while the first and the second leaves are gently squeezed between gloved first finger and thumb. 4. The entire leaf surface is then coated with the mixture by sliding the gently pinched fingers from base to tip two times.

3.3.2. Silencing Endogenous Barley or Wheat PDS with BSMV

1. Greenhouse-grown barley and wheat seedlings are inoculated with the 1:1:1 mixtures of in vitro transcripts synthesized from plasmids containing the wild-type BSMV and β RNAs, and derivatives of the γ RNA that carry either no plant sequence (BSMV:00) or the PDS fragments. 2. Seven days after rub-inoculating the first and second leaves of 7-day-old seedlings with BSMV-PDS, evidence of photobleaching is first apparent in third and fourth leaves of the barley plants. Photobleaching also develops in wheat, but it is usually not detectable until 10 days after viral inoculation (see Notes 9 and 10).

3.3.3. Using BSMV-VIGS to Identify Genes Required in Disease R-Pathways

Eight days is usually the appropriate interval of time necessary between infecting with BSMV to initiate VIGS and the application of the pathogen to challenge the plant resistance system.

3.4. Test for Knock Down of Targeted mRNAs

1. To determine whether the RNA-inducing transgenes affect mRNA levels of targeted genes, quantitative RT-PCR is performed by using two primer pairs. One pair of primers is specific for the mRNA of interest and they are designed to

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3.4.1. Quantitative Real-Time PCR

measure effective endogenous mRNA levels and not the transgene transcripts. 2. The second pair of primers amplifies a control gene not targeted for RNA silencing. For example, the glyceraldehyde-3phosphate dehydrogenase gene (GAPDH, AF251217) is used as an internal standard. 3. To normalize results, three replicates (three cDNAs) are performed for each RNA sample. 4. Real-time PCR assays can be performed with an ABI PRISM 7700 Sequence Detection System (Applied Biosystems) using SYBR Green PCR Master Mix (Applied Biosystems) in a final volume of 26µl including cDNA template and appropriate primer pairs. The amplification conditions are 2min at 50°C, 10min at 95°C, and 40 cycles of 15s at 95°C and 1min at 60°C.

3.4.2. Short Interfering RNA Detection

1. The hybridization blots are also hybridized with the housekeeping GAPDH gene (AF251217, glyceraldehyde-3-phosphate dehydrogenase) as a control. 2. The relative intensity of the hybridization signals in the transgenics versus wild-type plants can be determined with a phosphoimager (Cyclone gene array system, Perkin–Elmer, Boston).

4. Notes 1. The inverted repeat can be stabilized in bacteria through separation of the self-complementary regions by a “spacer” region. When the spacer sequence encodes an intron, the efficiency of gene silencing is very high, with up to 100% of the transformants generated with a particular gene construct showing some degree of silencing (12). 2. BSMV, the type member of the hordeivirus family, is a positive-sense single-strand RNA virus with a tripartite genome composed of the α, β, and γ RNAs. Fragments of transcribed sequences from the plant gene to be targeted for silencing are inserted into a DNA plasmid, from which the γ RNA can be synthesized by in vitro transcription. The plant cDNA fragment is cloned immediately downstream of the termination codon of the γb open reading frame (Fig.2). 3. In general, the “cleaner” the RNA preparation (i.e. free of protein, carbohydrate, etc.), the better the electrophoresis will resolve the RNA. 4. If the concentration of RNA is high enough, precipitation often occurs immediately and visibly. Otherwise, incubation

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on ice for 30min aids precipitation. Precipitation can also be improved by dissolving the 10% PEG/0.5M NaCl buffer in 50% formamide instead of water. 5. Let the bromophenol blue get to just before the bottom of the gel before stopping run. The siRNAs will have run twothirds to three-quarters of the way down the gel. 6. Recently, some commercial siRNA supply companies, such as Dharmacon, Qiagen, Proligo, Ambion, and Invitrogen, and some academic institutions, like Cold Spring Harbor Laboratories and the Whitehead Institute, have developed their unique siRNA search programs. Readers can access those programs through their web sites. 7. Each RNAi construct contains a cDNA fragment derived from the respective target gene and oriented in the antisense and sense directions at the 5′ and 3′ ends of the construct, respectively, separated by the intron of choice (Fig.3). 8. The variation in the degree of silencing observed in the transformants showing both reduction and loss of function may be a useful feature for gene discovery and functional genomics. Complete silencing of genes encoding a key element in basic cell functions or at particular developmental stages may result in lethality, whereas the reduced gene expression may give viable plants with phenotypes indicative of the role of the target gene. 9. In wheat, photobleaching less frequently extends across the width of the leaf without interruption than in barley and often has a striped appearance (34). 10. Symptoms associated with the BSMV:00 control virus are less severe in wheat than in barley. Variations in the susceptibility to BSMV infection exist among different barley genotypes, whereas wheat lines are uniformly conducive to BSMV-VIGS (34).

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40. Regina, A., Bird, A., Topping, D., Bowden, S., Freeman, J., Barsby, T., Kosar-Hashemi, B., Li, Z., Rahman, S. and Morell, M. (2005) High-amylose wheat generated by RNA interference improves indices of large bowel health in rats. P.N.A.S. USA 103, 3546-3551. 41. McGinnis, K., Murphy, N., Carlson, A. R., Akula, A., Akula, C., Basinger, H., Carlson, M., Hermanson, P., Kovacevic, N., McGill, M. A., Seshadri, V., Yoyokie, J., Cone, K., Kaeppler, H. F., Kaeppler, S. M. and Springer, N. M. (2007) Assessing the efficiency of RNA interference for maize functional genomics. Plant Physiol. 143, 1441-1451. 42. Reynolds, A., Leake, D., Boese, Q., Scaringe, S., Marshall, W. S. and Khvorova A. (2004) Rational siRNA design for RNA interference. Nature Biotechnol. 22, 326-330. 43. Kimbara, J., Takashi, R. and Nasuda, S. (2004) Characterization of the genes encoding for MAD2 homologues in wheat. Chromosome Res. 12, 703-714. 44. Christensen, A. H., Sharrock, R. A. and Quail, P. H. (1992) Maize polyubiquitin genes: structure, thermal perturbation of expression and transcript splicing, and promoter activity following transfer to protoplasts by electroporation. Plant Mol. Biol. 18, 675-689. 45. Qiu, S., Adema, C. M. and Lane, T. (2005) A computational study of off-target effects of RNA interference. Nucleic Acid Res. 33, 1834-1847. 46. Cigan, A. M., Unger-Wallace, E. and HaugCollet, K. (2005) Transcriptional gene silencing as a tool for uncovering gene function in maize. Plant J. 43, 929-940. 47. Yan, H., Chretien, R., Ye, J. and Rommens, C. M. (2006) New constructs approaches for efficient gene silencing in plants. Plant Physiol. 141, 1508-1518. 48. Chuang, C. F. and Meyerowitz, E. M. (2000) Specific and heritable genetic interference by double-stranded RNA in Arabidopsis thaliana. P.N.A.S. USA 97, 4985-4990. 49. Pogue, G. P., Lindbo, J. A., Dawson, W. O. and Turpen, T. H. (1998) Tobamovirus transient expression vectors: tools for plant biology and high-level expression of foreign proteins in plants, in Plant Molecular Biology Manual (Gelvin, S. B. and Schilperoot, R. A., eds.), Kluwer Academic Publishers, Dordrecht, The Netherlands, pp. 1-27.

Chapter 13 Gene Insertion Patterns and Sites Philippe Vain and Vera Thole Abstract During the past 25 years, the molecular analysis of transgene insertion patterns and sites in plants has greatly contributed to our understanding of the mechanisms underlying transgene integration, expression, and stability in the nuclear genome. Molecular characterization is also an essential step in the safety assessment of genetically modified crops. This chapter describes the standard experimental procedures used to analyze transgene insertion patterns and loci in cereals and grasses transformed using Agrobacterium tumefaciens or direct transfer of DNA. Methods and protocols enabling the determination of the number and configuration of transgenic loci via a combination of inheritance studies, polymerase chain reaction, and Southern analyses are presented. The complete characterization of transgenic inserts in plants is, however, a holistic process relying on a wide variety of experimental approaches. In this chapter, these additional approaches are not detailed but references to relevant bibliographic records are provided. Key words: Transgene copy, transgenic locus, genomic DNA, polymerase chain reaction, Southern analysis, inheritance, segregation.

1. Introduction Transgene insertion patterns and sites in the plant nuclear genome vary widely between independent transformation events and between strategies used to deliver the foreign DNA into the plant genome. Over the past 30 years, Agrobacterium-mediated transformation has been the major DNA delivery system for novel transgenic technologies (1), starting with the transformation of dicotyledonous species in the 1980s (2) followed by monocotyledonous species in the 1990s (3). Direct transfer of DNA in protoplasts (4, 5) or via particle bombardment (6, 7) has played a significant additional role in developing fundamental and applied transgenic research (8). To date, most transformation approaches have relied on the integration of transgene(s) into the plant nuclear Huw D. Jones and Peter R. Shewry (eds.), Methods in Molecular Biology, Transgenic Wheat, Barley and Oats, vol. 478 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-379-0_13

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genome through near-random illegitimate recombination, sometimes involving microhomologies (9). Therefore, it is not known where and how the transgene(s) will integrate into the plant nuclear genome. Past analyses of tens of thousands of transformation events in functional genomic programs (i.e., T-DNA insertion lines) have shown that transgene(s) frequently integrates in generich plant nuclear regions; however, this might be due to the fact that selectable marker genes used during the transformation process have to be functional to enable the recovery of transgenic plants. Agrobacterium-based transformation technologies are generally best suited to generate populations of transgenic plants containing a low number of transgene copies with limited rearrangements, which can be integrated at one or multiple unlinked Mendelian loci. New generations of binary vectors also contribute to limit the transfer of unwanted and superfluous DNA sequences, such as vector backbone (10) or non-plant DNA (11), into the plant nuclear genome. Direct transfer of DNA often produces populations of transgenic plants containing more complex inserts at a single Mendelian locus (12, 13). Recent progress in the development of particle bombardment technologies using dephosphorylation of linear DNA fragments without backbone sequences and low quantities of DNA has shown to significantly improve transgene insert structure. However, sequencing of these transgenic loci revealed frequent small-scale rearrangements, which can lead to the creation of multiple new open reading frames (ORFs) as well as the frequent de novo insertion of chloroplastic DNA together with the transgenes. In this chapter two examples of transformation events are provided to illustrate the different methodologies. In both cases a binary vector containing HPT (coding for hygromycin resistance) as a selectable marker gene and GUS (coding for β-glucuronidase) as a reporter gene present in a single T-DNA was delivered either via particle bombardment (Event#1) or via Agrobacterium (Event#2). The transgenes are driven by promoter and terminator sequences enabling their expression in the plant nucleus, and it is assumed that neither integration nor expression of the transgenes occurs in the plastidial genomes.

2. Materials 2.1. Inheritance Studies

●

Disposables and equipment 1. Controlled environment rooms and/or glasshouse facilities for plant culture 2. Controlled tissue culture room (25°C)

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3. Incubator at 37°C (with orbital shaker) 4. Laminar flow hood with Bunsen burner/gas supply 5. Stereomicroscope (Leica MZ6) 6. Thermocycler 7. Gel-doc system with Multi-Analyst software (Bio-Rad) for gel analysis 8. Horizontal gel electrophoresis equipment (e.g., gel tank system with power supply) 9. Vacuum chamber connected to a pump 10. Balance 11. Fridge (4°C) and freezer (−20°C) 12. Micropipettes and corresponding pipette tips 13. Pipettes (10 ml and/or 20 ml) 14. Ice 15. Liquid N2 16. Filter paper (e.g., Whatman) 17. Disposable plastic plates (96-, 24- or 6-well) 18. Microfuge tubes of various sizes (0.2, 0.5, 1.5, and/or 2 ml) 19. Disposable plastic tubes of various sizes (15 and/or 50 ml) 20. Plastic Petri dishes (9 cm diameter) 21. Scalpel and forceps ●

Reagents 1. Ethanol (70%) 2. Sodium hypochlorite solution (Fluka 71696) 3. Sterile deionized water 4. MS medium: Murashige and Skoog medium including B5 vitamins (Duchefa M0231), 10 g/l sucrose, 6 g/l agarose (Sigma A7921) and 2 g/l phytagel, pH 5.8 5. GUS assay solution: 100 mg X-gluc (5-bromo-4-chloro-3indolyl-b-D-glucuronide) dissolved in 2 ml DMSO (dimethyl sulphoxide) adjusted to 200 ml with 10 mM EDTA, 0.1% Triton X-100, 100 mM sodium phosphate, pH 7.0 6. DNeasy Plant Mini kit (Qiagen, #69104) 7. Taq DNA Polymerase (e.g., GE Healthcare UK Ltd, 270799-06) and its 10× reaction buffer 8. dNTPs stock solution (2 mM) 9. Primers (10 µM stock solutions, e.g., GUS forward and reverse primer pair; housekeeping gene forward and reverse primer pair)

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10. Agarose 11. Ethidium bromide: 10 mg/ml stock solution 12. 10× TBE: 0.89 M Tris-base, 0.89 M boric acid, 0.025 M EDTA, pH 8.3 2.2. Southern Analyses

●

Disposables and equipment 1. Phosphor imager (e.g., Typhoon 9200, Amersham Biosciences) 2. Spectrophotometer and quartz cuvettes 3. Gel-doc system with Multi-Analyst software (Bio-Rad) for gel analysis 4. Incubators at 37°C and 65°C 5. Thermocycler 6. Densitometer (e.g., Bio-Rad 690) 7. Fridge (4°C), freezer (−20°C), freezer (−80°C) 8. Speedvac DNA110 (at medium heat) 9. Horizontal gel electrophoresis equipment (e.g., gel tank systems with power supply) 10. Table-top orbital shaker 11. Storage phosphor screen (e.g., Molecular Dynamics) 12. Film for autoradiography 13. DNA sequencing service or equipment 14. HybondN+ nylon membrane (Amersham) 15. Blotting equipment (e.g., plastic tray, glass plate, blotting paper) 16. Pestle and mortar or disposable plastic pestles 17. Disposable plastic tubes of various sizes (15 and/or 50 ml) 18. Microfuge tubes of various sizes (0.2, 0.5, 1.5 and/or 2 ml) 19. Cling film 20. Micropipettes and corresponding pipette tips 21. Pipettes (10 and/or 20 ml) 22. Ice 23. Liquid N2

●

Reagents 1. Nucleon Phytopure Plant DNA Extraction Kit (Amersham Biosciences) 2. DNeasy Plant Mini kit (Qiagen, #69104) 3. QIAquick Gel Extraction kit (Qiagen, #28101)

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4. QIAquick PCR Purification kit (Qiagen, #28104) 5. Restriction enzymes and their corresponding reaction buffers 6. DNA sequencing kit 7. Agarose 8. 10× TBE: 0.89 M Tris-base, 0.89 M boric acid, 0.025 M EDTA, pH 8.3 9. Lambda DNA 10. Ethidium bromide (10 mg/ml) 11. Sterile deionized water 12. DNA Polymerase I, Klenow fragment 13. [32P]-dCTP (Amersham) 14. Carrier DNA (autoclaved herring testes DNA, 5 g/l) 15. OLB (Oligo Labelling Buffer): 2 ml (1.25 M Tris–HCl, pH 8.0 and 0.125 M MgCl2), 36 µl b-mercapto-ethanol, 10 µl dATP, 10 µl dTTP, 10 µl dGTP, 5,165 µl of 2 M Hepes (pH 6.6), 3,099 µl of 90 OD260 U Random Hexamer (Pharmacia)/µl 16. BSA (bovine serum albumin, 50 mg/ml) 17. T1/10E: 3 mM Tris–HCl, pH 7.0 and 0.02 mM EDTA, pH 8.0 18. SDS (sodium dodecyl sulfate, 10% w/v) 19. 0.25 M HCl 20. 0.4 M and 3 M NaOH 21. 20× SSC: 3 M NaCl and 0.3 M sodium citrate, pH 7.0 22. 5× HSB: 175.3 g NaCl, 30.3 g PIPES and 7.45 g Na2EDTA 2H2O in 1 l H2O, pH 6.8 with NaOH 23. Denhardt’s reagent III: 2 g gelatin, 2 g Ficoll-400, 2 g polyvinylpyrrolidone-360, 10 g SDS, and 5 g Na2P2O·10H2O in 100 ml of H2O, dissolved at 65°C.

3. Methods 3.1. Number of Transgenic Loci

The number of transgenic loci present in a primary transgenic plant (T0) is generally determined using inheritance studies. Such analysis can be undertaken at any generation but is often conducted at the second generation (T1) after self-pollination or a test-cross with a wild-type plant. The relative number of T1 plants expressing and not expressing the transgene provides an observed segregation ratio of transgene phenotype. The relative number of

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T1 plants containing and not containing the transgene provides an observed segregation ratio of transgene genotype. The former is frequently used to estimate the number of transgenic loci. However, this approach often underestimates the real number of loci, which can be precisely determined using the latter approach. Comparative molecular analysis of T0 and progeny plants can also give information about the number of transgenic loci. In recent years, the precise determination of the locus number in transgenic plants (14, 15) has played an important role in developing “clean-gene” technologies that produce plants free of any selectable marker gene following transformation with multiple T-DNAs (16). The number of transgenic loci is not to be confounded with the number of transgene copies in a given plant genome because several transgene copies can integrate at a single transgenic locus and single transgene copies can integrate at different loci. ● Segregation of transgene phenotype Primary (T0) transgenic plants can be self-pollinated or crossed to produce T1 seeds. T1 plants are scored for the expression of the transgene(s). Any of the transgenes introduced into the plant genome can be used to phenotype T1 plants. The selectable marker gene(s) used to produce the original T0 plants is often used to screen T1 plants for transgene expression. However, transgene(s) of interest, which can be easily scored, can also be used to phenotype T1 plants (such as genes affecting starch metabolism detected by iodine staining). Antibiotics such as kanamycin or hygromycin can be used to identify progeny plants containing a functional NPTII or HPT gene, respectively (see Note 1). Herbicides such as glufosinate, sulfoylurea, or glyphosate can be used to detect progeny plants containing a functional BAR, ALS or EPSP gene, respectively. Herbicides enable facile phenotyping of T1 plants by spraying the seedlings or painting whole plant leaves. Reporter genes (such as GUS, LUC, or GFP) are also often used to assess transgene expression in T1 plants. Non-destructive phenotyping strategies of T1 plants (e.g., observation of GFP or LUC expression) are always preferable to destructive ones (e.g., GUS staining or antibiotic/herbicide selection), as they enable the subsequent molecular analysis of T1 plants non-expressing the transgene(s). For Event#1 and Event#2, the expression of the GUS gene in T1 plants produced by self-pollination of T0 plants was assessed using histochemical staining of either endosperm, leaf, or root tissue. 1. For endosperm staining, seeds are sterilized for 30 s with 70% ethanol followed by 15 min with 50% sodium hypochlorite solution, then rinsed three times with sterile deionized water, and stored for a few hours in the last rinsing water. Sterile seeds are cut into two halves with a sterile scalpel under a laminar flow hood. The part of the seed containing the embryo is used to produce a T1 seedling on jellified MS medium, and the other half is used for GUS staining. In most cases, phe-

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notyping the endosperm is comparable to phenotyping the embryo itself, as they have a similar genotype, however, with different ploidy levels. 2. For leaf or root staining, there is no requirement to sterilize seeds; however, seeds can be washed for 30 s with 70% ethanol and rinsed three times with deionized sterile water. For some species, such as wheat and barley, seeds can be kept for 2–4 days on a wet filter paper at 5°C in the dark before germination at 20–25°C in the dark for 5 days, then under a 16-h photoperiod. The leaf or root tissues can be sampled after 5–7 days at 20–25°C. This method is often preferred to endosperm staining for species that are sensitive to sodium hypochlorite (such as wheat and barley). 3. Tissue samples are submerged into the GUS assay mixture in 96-, 24-, or 6-well disposable plastic plates depending upon the size of the plant sample. 4. Samples (in open plates) are vacuum infiltrated (28 in.Hg) for 10–15 min, and then incubated at 37°C overnight, preferably with gentle shaking on an orbital shaker. 5. The following day, samples are observed for GUS staining. An example of GUS staining of endosperms from T1 seeds produced from Event#2 is shown in the Fig. 1. 6. Estimation of the number of transgenic loci: The relative number of T1 plants expressing and not expressing the GUS gene provides an observed segregation ratio. This observed ratio can be statistically compared to theoretical Mendelian inheritance patterns for one gene (Table 1) using a Chi-square analysis. In the case of Event#2, 83 T1 seeds expressed the GUS gene and 17 did not (Fig. 1). Chi-square analysis suggested the presence of one transgenic locus (Table 2). However, it must be emphasized that this approach, despite being the most commonly used in the scientific literature, provides only an estimate of the number of transgenic loci (see Note 2). In the case of Event#2, analysis based on the phenotype was misleading, as further molecular analysis (see Inheritance analysis based upon transgene genotype – Section 3.1) demonstrated that Event#2 contains in fact two independent transgenic loci instead of only one. True inheritance studies should be based upon the presence (genotype) and not the expression (phenotype) of the transgene in progeny plants. ●

Inheritance analysis based upon transgene genotype Tables 1 and 3 show the theoretical Mendelian inheritance patterns for one or two genes inserted at one, two, three, or four loci after self-pollination respectively. T1 plants are scored for the presence of the transgene(s). This is generally conducted by molecular analysis such as PCR or dot blot analysis. Genotyping

Fig. 1. Histochemical staining of endosperm to assess the expression of the GUS gene in a segregating population of progeny seeds. The example presented is Event#2: 83 out of 100 T1 seeds are expressing the GUS gene (dark coloration).

Table 1 Theoretical Mendelian inheritance patterns for a transgene present at one, two, three, or four loci. In this example, the T0 plant has been transformed with a single DNA fragment/vector containing a GUS reporter gene. The GUS gene is either present (G) or absent (g) in each T1 plant. The expected percentage of T1 plants containing or not the transgene in the segregating population is presented after either selfpollination or testcross with a non-transgenic wild-type plant Genotype of T1 plants a

Genotype of T0 plant

G (in %)

g (in %)

1 Locus (G)

75

25

2 Loci (G) (G)

93.75

6.25

3 Loci (G) (G) (G)

98.44

1.56

4 Loci (G) (G) (G) (G)

99.61

0.39

1 Locus (G)

50

50

2 Loci (G) (G)

75

25

3 Loci (G) (G) (G)

87.5

12.5

4 Loci (G) (G) (G) (G)

93.75

6.25

Self-pollination

Testcross with wild-type

a

(G) (G) = T0 plant containing two transgenic loci, each locus (G) contains the GUS gene. T0 plants are always hemizygous for a given transgene at each transgenic locus (i.e., the transgene integrates in only one of the homologous chromosomes in T0 plants)

Gene Insertion Patterns and Sites

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Table 2 Inheritance study of Event#1 and Event#2 based on the segregation of transgene phenotype and genotype. T1 plants were produced by self-pollination of T0 plants. For Event#2, gene silencing in 7 out of the 100 T1 plants leads to an incorrect evaluation of the number of transgenic loci when using transgene phenotype in segregation studies. For Event#1, gene silencing in 2 out of the 100 T1 plants did not affect the estimation of the number of loci Phenotype of T1 plants GUS−

(χ2)a

Genotype of T1 plants GUS

No GUS

69

31

(χ2)a

Event#1

Observed

71

29

1 Locus

Expected

75

25

ns

75

25

ns

2 Loci

Expected

93.75

6.25

S

93.75

6.25

S

3 Loci

Expected

98.44

1.56

S

98.44

1.56

S

4 Loci

Expected

99.61

0.39

S

99.61

0.39

S

Event#2

Observed

83

17

90

10

1 Locus

Expected

75

25

ns

75

25

S

2 Loci

Expected

93.75

6.25

S

93.75

6.25

ns

3 Loci

Expected

98.44

1.56

S

98.44

1.56

S

Expected

99.61

0.39

S

99.61

0.39

S

4 Loci a

GUS+

2

2

Chi-square (χ ) is calculated as the sum of (observed − expected) /expected. For “Event #2 – phenotype of T1 plants − 1 Locus,” the calculation is ((83−75)2/75) + ((17−25)2/25) = 3.4. This indicates that the observed segregation ratio is similar (ns) to the expected segregation for 1 locus (at P < 0.05). χ2 values above 3.84 indicate a significant difference between expected and observed ratios (S). Alternatively, the χ2 can be calculated using the Yate’s correction (for analysis with one degree of freedom) as the sum of (|observed − expected| − 0.5)2/expected. For “Event #2 – phenotype of T1 plants − 1 locus,” the calculation is ((|83 − 75| − 0.5)2/75) + ((|17 − 25| − 0.5)2/25) = 3.0

T1 plants using molecular analysis is more expensive and labour intensive than phenotyping T1 plants (see Segregation of trangene phenotype – Section 3.1). For this reason, it is rarely used to conduct inheritance studies despite the limitations associated with segregation analyses based on transgene phenotype (see Note 2). An optimized approach is to combine both phenotyping and genotyping of T1 plants when possible. Such an analysis for Event#2 is presented below: 1. Plants are grown and phenotyped following steps 2–5 in segregation of transgene phenotype- section 3.1. It is important to grow all T1 plants without selection in order to keep all progeny plants alive and available for further molecular analysis. Treatment of T1 seeds or plants with antibiotics or herbicides would kill the transgenic progeny containing silenced transgenes and would therefore skew the inheritance analysis (see Note 2). For Event#2, the 83 T1 seeds clearly expressing

Table 3 Theoretical Mendelian inheritance patterns for two genes inserted at one, two, three, or four loci after self-pollination. In this example, the T0 plant has been transformed with two DNA fragments or vectors, one containing a GUS reporter gene and the other a LUC reporter gene. The GUS gene is either present (G) or absent (g) in a T1 plant. The LUC gene is either present (L) or absent (l) in a T1 plant. The expected percentage of T1 plants containing or not the GUS and LUC transgenes in the segregating population is presented Genotype of T0 planta

Genotype of T1 plants G – L (in %) g – L (in %)

G – l (in %)

g – l (in %)

1 Locus (G/L)

75

0

0

25

2 Loci (G) (L)

56.25

18.75

18.75

6.25

2 Loci (G) (G/L)

75

0

18.75

6.25

2 Loci (G/L) (L)

75

18.75

0

6.25

2 Loci (G/L) (G/L)

93.75

0

0

6.25

3 Loci (G) (L) (L)

70.31

23.44

4.69

1.56

3 Loci (G) (G) (L)

70.31

4.69

23.44

1.56

3 Loci (G/L) (G/L) (G)

93.75

0

4.69

1.56

3 Loci (G) (G/L) (L)

89.06

4.69

4.69

1.56

3 Loci (G/L) (G/L) (L)

93.75

4.69

0

1.56

3 Loci (G/L) (G) (G)

75

0

23.44

1.56

3 Loci (G/L) (L) (L)

75

23.44

0

1.56

3 Loci (G/L) (G/L) (G/L)

98.44

0

0

1.56

3 Loci (G) (G) (G)

0

0

98.44

1.56

3 Loci (L) (L) (L)

0

98.44

0

1.56

4 Loci (G/L) (G/L) (G/L) (G/L)

99.61

0

0

0.39

4 Loci (G) (G) (G) (G)

0

0

99.61

0.39

4 Loci (L) (L) (L) (L)

0

99.61

0

0.39

4 Loci (G) (G) (G) (L)

73.83

1.17

24.61

0.39

4 Loci (G) (G) (L) (L)

87.89

5.86

5.86

0.39

4 Loci (G) (L) (L) (L)

73.83

24.61

1.17

0.39

4 Loci (G/L) (L) (L) (L)

75

24.61

0

0.39

4 Loci (G/L) (G) (L) (L)

92.58

5.86

1.17

0.39

4 Loci (G/L) (G) (G) (L)

92.58

1.17

5.86

0.39

4 Loci (G/L) (G) (G) (G)

75

0

24.61

0.39

4 Loci (G/L) (G/L) (L) (L)

93.75

5.86

0

0.39

(continued)

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Table 3 (continued) Genotype of T1 plants G – L (in %) g – L (in %)

Genotype of T0 planta

G – l (in %)

g – l (in %)

4 Loci (G/L) (G/L) (G) (L)

97.27

1.17

1.17

0.39

4 Loci (G/L) (G/L) (G) (G)

93.75

0

5.86

0.39

4 Loci (G/L) (G/L) (G/L) (L)

98.44

1.17

0

0.39

4 Loci (G/L) (G/L) (G/L) (G)

98.44

0

1.17

0.39

a

(G/L) (G) = T0 plant containing two transgenic loci: one transgenic locus (G/L) contains both the GUS and LUC genes and the other locus (G) contains only the GUS gene. T0 plants are always hemizygous for a given transgene at each transgenic locus (i.e., the transgene integrates in only one of the homologous chromosomes in T0 plants)

the GUS gene (Fig. 1) are considered to contain the GUS gene and are not analyzed further. The 17 T1 plants not expressing GUS are analyzed at the molecular level to determine whether they contain (transgenic but silenced) or not (non-transgenic segregant) the GUS gene. 2. Leaf samples matching the length of a 1.5-ml microfuge tube are collected from the 17 T1 plants not expressing the GUS gene. Leaf samples are also collected from a negative control (i.e., wild-type plant) and from a positive control (e.g., transgenic plant previously characterized by PCR). 3. DNA is extracted using the DNeasy Plant Mini kit (Qiagen) following the instructions of the manufacturer. 4. The quantity of the DNA template used for PCR varies depending upon the genome size of each cereal species. Twenty-five nanograms of rice DNA or 100–150 ng of barley/ wheat DNA is often used for a 25 µl PCR reaction. 5. Each PCR reaction contains plant DNA, 1× reaction buffer (including 2.5 mM MgCl2), 250 µM of dNTPs, 0.4 µM of each primer (e.g., corresponding to the GUS gene), and 2 units of Taq DNA polymerase. 6. Samples are denatured at 95°C for 1 min, followed by 30 cycles at 95°C for 30 s, 60°C for 30 s, and 72°C for 1 min. PCR is terminated by a 10 min extension step at 72°C. 7. Each PCR reaction (5–10 µl) is loaded onto a 1–1.2% (w/v) agarose gel (in 0.5× TBE) containing ethidium bromide (5 µl in a 100 ml gel) and subjected to electrophoresis at 80–100 V for 20 min. Examination of the gel under UV light (Fig. 2) showed that 7 out of the 17 T1 plants contained the GUS gene but did not express it (i.e., silenced GUS gene). 8. Steps 5–7 are repeated with PCR primers amplifying a single copy of a housekeeping gene or of an Restriction Fragment Length Polymorpgism (RFLP) probe in order to assess

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Fig. 2. PCR analysis to assess the presence of the transgene in non-expressing progeny plants. The example presented is Event#2: 7 out of 17 T1 plants that exhibited no GUS staining (Fig. 1) contain a silenced GUS gene.

whether the quality of the DNA samples was suitable for PCR amplification. Analysis of the samples with primers amplifying a housekeeping gene showed that the ten remaining samples from Event#2 were suitable for PCR amplification (Fig. 2). 9. Calculation of the number of transgenic loci: The observed ratio can be statistically compared to Mendelian inheritance patterns for one gene (Table 1) using a Chi-square analysis. For Event#2, 90 T1 plants contained the GUS gene and 10 did not, showing that the GUS gene integrated at two independent unlinked Mendelian loci in the plant genome (Table 2). Southern analysis across generations Southern analysis is generally used to characterize the arrangement of individual transgenic loci (see southern analysis – Section 3.2) but it can also give information on the number of transgenic loci by comparison of hybridization patterns across generations. For example, the differences in banding patterns between the primary T0 plant and a population of self-pollinated segregating T1 plants can be used to assess the number of transgenic loci. Such an analysis is presented for Event#2 below: 1. DNA is extracted from plant leaf material (see Note 3) using the Nucleon Phytopure Plant DNA Extraction Kit following the recommendations of the manufacturer. DNA concentration is calculated on the basis of absorbance at λ = 260 nm. DNA is kept in water at 4°C for short-term storage and at −20°C for long-term storage (see Note 4). ●

2. The quantity of DNA used for Southern analysis varies depending upon the genome size of each cereal species (typically 5 µg for rice or 20 µg for barley/wheat). DNA is digested in a 50–80 µl volume using 10–15 units of an enzyme per microgram of DNA (see Note 5). DNA from a negative control

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(i.e., wild-type plant) and a positive control (e.g., transgenic plant previously characterized by Southern analysis) are also digested. 3. The volume of each restriction digest is reduced to approximately 30 µl using a Speedvac and loaded onto a 0.8% agarose gel (in 1× TBE), which is run at 40 V overnight. The gel should contain one or two molecular makers (such as 600 ng of lambda DNA digested with either PstI or HindIII). Restriction enzyme buffer is added to the blank lanes to avoid distortion of the bands during electrophoresis. 4. A photo of the gel is taken under UV light after staining with ethidium bromide in order to check the digestion level and equal loading of the DNA samples, as well as to record the position of the bands from the molecular maker(s). 5. The gel is washed (with gentle agitation) for 15 min in 0.25 M HCl, rinsed with H2O, denatured for 15 min in 0.4 M NaOH, and blotted onto a HybondN+ nylon membrane using 0.4 M NaOH overnight. The membrane is rinsed for 2 min in 2 × SSC, wrapped in cling film, and stored 5°C (short-term) or −20°C (long-term). 6. The membrane is pre-hybridized for 2–5 h at 65°C by gentle shaking in a 50 ml solution containing 2 ml of 5× HSB, 1 ml of Denhardt’s reagent III, and 1 ml of carrier DNA (denatured by boiling for 7 min). 7. Probes (approximately 500 bp in length) can be produced by restriction digest of the plasmid used for transformation followed by gel purification (e.g., QIAquick Gel Extraction kit) or via PCR followed by column purification (e.g., QIAquick PCR Purification kit) following the manufacturer’s recommendations. Twenty-five nanograms (in 15 µl) of each probe is boiled for 7 min and kept on ice for 5 min. The probe is labeled by adding 5 µl of OLB, 2 µl of BSA, 2 µl of DNA Polymerase I Klenow fragment (5 U/µl), and 2 µl of 32PdCTP and by incubating this mixture for 2 h at 37°C. The probe is denatured for 5 min using 2.6 µl (i.e., 1/10 volume) of 3 M NaOH and then added to the pre-hybridization solution covering the membrane. The membrane is gently shaken in this mixture at 65°C overnight. 8. The membrane is washed twice with 500 ml of 2 × SSC, 1% SDS for 15 min followed by two additional 15 min washes with 500 ml of 0.2 × SSC, 1% SDS. The membrane is wrapped in cling film. 9. Estimation of the number of transgenic loci: The membrane is analyzed by autoradiography (when using films) or by analysis of the storage phosphor screen in a phosphor imager. For Event#2, the comparison of banding patterns between T0

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Fig. 3. Southern analysis of T0 and T1 progeny plants produced by self-pollination. The example presented is Event#2 (Xba I digest, GUS probe). 1, 2 and 1 + 2 indicate the hybridization pattern of T1 plants containing either the locus 1, the locus 2, or the locus 1 + locus 2, respectively. Nil indicates a non-transformed segregating plant.

plant and segregating T1 plants suggests the presence of two transgenic loci (Fig. 3). The limitations of this approach to precisely determine the number of transgenic loci are detailed in Note 6. This approach can also be used in subsequent generations to assess the stability of the structure of the transgenic loci. When more than one DNA fragment/vector has been used for transformation, this analysis should be conducted using either a probe common to each fragment used for transformation or a mixture of two probes (i.e., one probe for each fragment/vector). 3.2. Configuration of Each Transgenic Locus

When transgenes are inserted at a single locus, T0 plants can be analyzed at the molecular level in order to determine the number of transgene copies inserted and the configuration of the transgenic locus. When more than one transgenic locus has been identified (using protocols detailed in Section 3.1), each independent locus should be analyzed separately (see Note 7). This can be achieved using a segregating population of plants or lines homozygous for each individual locus. Southern analysis Southern analysis is the most common approach used to characterize the configuration of transgenic loci. A combination of restriction enzymes and probes covering the entire DNA sequence used for transformation (including vector backbone sequences) generally enables the reconstitution of the overall structure of each locus. A first step is often to estimate the number of transgene copies (intact and/or rearranged) at each locus. Different types of analyses will be conducted depending upon the complexity of a transgenic locus. An analysis of a simple (one to a few copies) or complex (many copies) transgenic locus is presented below: 1. DNA is extracted; the membrane is produced and hybridized as detailed in steps 1–8 – Southern Analysis across generations – Section 3.1. A restriction enzyme cutting once in

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the middle of the fragment/vector used for transformation (XbaI in the plasmid used as an example) is generally utilized to digest the plant DNA. Probes corres-ponding to transgene sequences 5′ (HPT in our example) and/or 3′ (GUS in our example) of this restriction site are used for hybridization (see Note 8). 2. The membrane is analyzed by autoradiography or using a phosphor imager. This preliminary analysis is often suffi-cient for an initial estimation of the complexity of each transgenic locus. i. Analysis of a simple transgenic locus: Southern analysis showing up to four hybridization bands of similar intensity for most enzyme–probe combinations, such as Event#2 locus1 and Event#2 locus2. (a) Southern analysis (steps 1–8 – Southern Analysis across generations – Section 3.1) is conducted with a range of restriction enzymes (see Note 9) and probes following the general guidelines presented in Table 4. This approach enables the study of both border and internal regions of a transgenic locus. It is recommended to use several enzymes and probes with the same characteristics for a more precise analysis. For example, in Event#2 locus1 (Fig. 4), an additional enzyme to HindIII (flanking the GUS expression unit) and probes corresponding to the promoter and terminator sequences of the GUS gene should be used, as it is possible that multiple and rearranged copies of the transgene have been inserted. Probes corresponding to antibiotic resistance genes present in the vector backbone should also be used to detect potential backbone transfer (Table 4). (b) Determination of the structure of Event#2 locus1: the presence of multiple bands for XbaI and HindIII digests with the GUS probe suggests the presence of more than one copy of the GUS gene (one intact and one probably deleted). However, only single bands are observed for the HPT and NPTI genes, the latter demonstrating that sequences derived from the vector backbone have been transferred and integrated into the plant genome. This is generally due to the inefficient recognition/processing of the T-DNA left-border (LB) leading to a ‘read-through’ beyond the borders of the T-DNA sequence (see Note 10). The size of the bands suggests a structure of the transgenic locus described in Fig. 4. (c) Determination of the structure of Event#2 locus2: only single bands are observed when using GUS- and HPTspecific probes. The absence of a hybridization signal for the NPTI probe indicates the absence of backbone

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Table 4 Guidelines for restriction digest and probing in Southern analysis of transgenic plants Type of restriction enzyme

Type of probe

Example provided

Enzyme cutting once in the fragment/vector used for transformation, between the expression units

Probe corresponding to each transgene sequence located 5′ or 3′ of this restriction site. The bands often contain both transgenic DNA and plant genomic sequences flanking the transgenic locus. Multiple bands often indicate multiple copies and/or transgenic loci

Event#2: Locus1 – XbaI with GUS or HPT probe (Fig. 4)

Enzyme flanking promoter: transgene:terminator expression units

Event#2: Locus1 – HinProbes corresponding to each transdIII with GUS probe gene sequence. If multiple bands are (Fig. 4) then use probes obtained, use probes corresponding corresponding to GUS to promoter or terminator sequences promoter and terminator for a more detailed analysis sequences

Enzyme flanking all expression units

Probes corresponding to each transgene sequence. The size of the band informs on the intactness of the insert

Enzymes not cutting into the fragment or vector used for transformation

Probes corresponding to each transgene Event#1 – PacI, NheI, BstXI, ApaI or KpnI with sequence. When single bands are GUS probe (Fig. 6) obtained with many (five or more) non-cutting enzymes, it suggests the presence of a single transgenic locus without large intervening plant nuclear DNA sequences

Any enzyme cutting in the fragment used for transformation

Probe from vector backbone sequence, Event#1 – XbaI or HindIII preferably sequence corresponding with NPTI probe (Fig. to the antibiotic resistance gene used 6) for vector selection in bacteria (E. coli or Agrobacterium)

Event#2: Locus2 – PmeI with GUS or HPT probe (Fig. 5)

transfer. The size of the bands suggests a structure of the transgenic locus described in Fig. 5. ii. Analysis of a complex transgenic locus: Southern analysis shows four or more hybridization bands with a wide range of signal intensity for most enzyme–probe combinations, such as Event#1 (Fig. 6). For such events, hybridization patterns are often unreliable (17) and cannot be used on their own to determine the exact configuration of the transgenic locus or the number of transgene copies (see Note 11). Nevertheless, densitometric analysis of hybridization signals can

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Fig. 4. Southern analysis of Event#2 – locus 1. DNA from T2 homozygous plants was digested using either XbaI (X), PmeI (P), or HindIII (H). Membranes were probed using sequences from the GUS, HPT or NPTI (kanamycin resistance gene in the vector backbone) genes.

Fig. 5. Southern analysis of Event#2 – locus 2. DNA from T2 homozygous plants was digested using XbaI (X), PmeI (P), or HindIII (H). Membranes were probed using sequences from the GUS, HPT, or NPTI (kanamycin resistance gene in the vector backbone) genes.

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Fig. 6. Southern analysis of Event#1. DNA from T0 plants or plasmid used for particle bombardment was digested using enzymes either not cutting [PacI (P), NheI (N), BstXI (B), ApaI (A), KpnI (K)] or cutting [XbaI (X), HindIII (H), SpeI (S)] in the plasmid used for transformation. The black triangle indicates increasing concentration of the PmeI restriction enzyme to digest plant DNA. The number of GUS or HPT copies (c) in reconstitution standards is represented. Membranes were probed using sequences from the GUS, HPT, or NPTI (kanamycin resistance gene in the vector backbone) genes.

be used to estimate the number of copies of each transgene and/or the number of intact expression units (i.e., promoter + transgene + terminator sequence). Southern analysis with restriction enzymes not cutting into the fragment/vector used for transformation can also give information on the potential presence of intervening plant genomic DNA within the transgenic locus: (a) DNA is extracted as in step 1 – Southern Analysis across generations – Section 3.1. The concentration of both wild-type plant genomic and plasmid DNA used f or reconstitution standards needs to be precisely assessed for reliable densitometric analyses (see Note 12). (b) DNA is digested as in step 2 – Southern Analysis across generations – Section 3.1. Reconstitution standards equivalent to 1, 2, 5, 10, 20, 40, 80 transgene copies are spiked into 5 µg wild-type DNA (see Note 13). It is recommended to also include a positive control(s) to each membrane (see Note 14). (c) Southern analysis (as in steps 3–8 – Southern Analysis across generations – Section 3.1) is conducted with plant DNA digested with HindIII flanking the GUS expression unit or with PacI, NheI, BstXI, ApaI, or KpnI that are not cutting into the plasmid used for transformation. Four different probes are used: first, a single-copy RFLP probe is

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applied and then probes corresponding to the GUS, HPT, and NPTI gene. The former analysis is essential to confirm the equal loading of DNA amounts for each sample (see Note 14). Digestion of plasmid DNA with HindIII yields a single band of 2.9 kb (Fig. 6). Digestion of the genomic DNA of Event#1 with HindIII shows some hybridization bands corresponding to intact monomers of the GUS expression unit (2.9 kb). Other bands, of higher and lower molecular weight, represent chimeric fragments resulting from DNA rearrangements. Similar profiles are obtained using a combination of a SpeI-digest and the HPT probe (Fig. 6). Digestion with the ‘non-cutting’ enzyme ApaI shows two bands with the GUS probe, suggesting that this complex transgenic locus contains intervening plant genomic DNA. A large number of NPTI gene copies are also present in the Event#1 locus confirming the presence of plasmid backbone (as the entire plasmid was used for particle bombardment). This information does not allow determination of the configuration for the Event#1 locus. (d) Determination of transgene copy number in Event#1: The intensity of all hybridization signals above background is quantified using either different film exposures of each membrane in a densitometer (Bio-Rad 690) or an analysis of a storage phosphor screen in a phosphor imager. A regression is produced on the basis of the signals obtained from the reconstitution standards establishing a relationship between signal intensity and copy number. The sum for all signals from each lane provides an estimate of the number of transgene copies for each enzyme–probe combination (see Note 15). Signals from the HindIII, SpeI, and XbaI digests suggest the presence of 36 GUS and 14 HPT copies in Event#1 (Fig. 6). • Additional methods Southern analyses generally provide useful information on the overall architecture of the transgenic locus. However, the full characterization of each transgenic locus can be obtained by sequencing the insert itself (18, 19), the plant genomic regions flanking the insert, and sometimes the genomic DNA prior to transgene insertion. These approaches are not detailed in this chapter but routine protocols have been published for the high-throughput identification of flanking genomic sequences in model plants such as A. thaliana (20, 21) or rice (22). After sequencing of the plant genomic regions flanking the insert, this information can be used to design specific PCR primers orientated towards the insert, enabling the step-by-step sequencing of the entire insert. Primers from flanking regions can also be used to

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sequence wild-type plant DNA in order to determine whether pre-existing genomic sequences have been deleted at the point of insertion. DNA sequencing of the insert also enables the identification of putative chloroplastic DNA that could have integrated along with the intended transgene(s). 3.3. Conclusions

The experimental procedures and strategies detailed in this chapter are commonly used to analyze transgene insertion patterns and loci in transformed plants including cereals and grasses. During the past 20 years, these molecular analyses have greatly contributed to our understanding on how and where transgenes integrate into the plant genome and enabled scientists to develop strategies to improve the molecular makeup of transgenic plants (1). These approaches also underpin the safety assessment of the genetically modified (GM) crops to be commercialized in the EU as detailed in the European Food Safety Authority (EFSA) guidance document of the scientific panel on GM organisms (http://www.efsa.europa.eu/etc/medialib/efsa/science/gmo/ gmo_guidance/gmo_guidance_ej374.Par.0001.File.tmp/gmo_ guidance_ej374_gmm.pdf).

4. Notes 1. Antibiotic selection is often inefficient when used on whole seeds (i.e., germination on a wet filter paper or gel-based culture medium containing antibiotics) or plants (i.e., leaf painting with antibiotic solutions) because of the naturally high resistance levels of cereals. When an antibiotic must be used, mature embryos should be dissected from the sterilized seeds and germinated, embryo axis down, on a gel-based medium using the same antibiotic concentration previously used to produce the T0 plants. 2. Gene silencing in progeny plants leads to falsely consider that progeny plants not expressing the transgene are non-transgenic. This can inflate the observed number of progeny plants considered as non-transgenic and consequently lead to underestimation of the number of transgenic loci (see Event#2 in Table 2). Gene silencing in progeny plants seems to occur more frequently when multiple transgenic loci are present in the original T0 plant. In rice plants transformed using a single type of T-DNA, the number of transgenic loci estimated on the basis of transgene expression is incorrect in 36% of the cases (23). When more than one type of DNA fragment/ vector is used in co-transformation experiments (e.g., multi-

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ple T-DNAs delivered via Agrobacterium or multiple DNA fragments delivered by particle bombardment), it is necessary to monitor the expression of one transgene per type of fragment/vector in the progeny plants. For example, if one T-DNA contains the GUS gene and another T-DNA contains the LUC gene, the expression of both transgenes will be assessed in each T1 plant to provide the observed segregation ratio. This observed ratio will then be compared to Mendelian inheritance patterns for two genes (Table 3) using a Chi-square analysis. In rice, relying on transgene expression to estimate the number of loci in plants transformed with two different T-DNAs is incorrect in 78% of the cases (15). Therefore, the segregation of transgene phenotype should be used only in conjunction with additional molecular analyses of transgenic plants to precisely determine the number of transgenic loci. 3. Young leaves are often preferred to old plant material for DNA extraction. In certain cereal species, such as rice, plants can be left in the dark for 1 or 2 days before DNA extraction in order to minimize the presence of polysaccharides and/or phenolic compounds in the extracts. This improves the quantity and quality of the plant DNA recovered. 4. Frequent freeze–thaw cycles as well as micropipetting of plant genomic DNA solution with small tips can shear DNA and should be avoided. 5. Digestion in a thermocycler with a heated lid prevents condensation on the lid of the microfuge tubes and helps keeping digestion volumes constant. 6. The comparison of hybridization profiles between T0 and progeny plants can be difficult when large numbers of transgene copies or transgenic loci are present in the plant genome or when the biological characteristics of the plant do not allow facile self-pollination (such as auto-incompatibility in some grass species). The number of progeny plants analyzed must be large enough to detect the hybridization pattern of each independent transgenic locus. As a rule of thumb, at least 8, 20, and 30 progeny plants obtained after self-pollination should be analyzed by Southern blotting to detect the presence of two, three, or four loci, respectively. However, at least 32 and 128 progeny plants will need to be analyzed to have a reasonable chance to observe each transgenic locus individually when plants contain three or four loci. In practice, it is preferable to use the approach detailed in inheritance analysis based upon transgene genotype – Section 3.1 rather than to compare hybridization profiles when plants contain more than two transgenic loci.

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7. When no information is available on the number of transgenic loci, the Southern analysis of T0 plants should be taken with caution, as the complexity of banding patterns may reflect transgene insertion at different loci and not the presence of multiple copies at one locus. The latter is frequent when transgenic plants are produced via Agrobacteriummediated transformation. In rice, more than half of the plants produced contain T-DNA sequences inserted at two or more loci (23). 8. Membranes should be checked for background signal using standard exposure before re-probing (i.e., using the same length of exposure as when membranes are probed). Artifactual background signals would compromise the precise determination of transgene copy number and the organization of the transgenic locus. Signals corresponding to high copy numbers (20 or more) can take a long time to decay. Membranes are stripped three times boiling in 500 ml of 0.1 × SSC and 0.5% SDS for 5 min. 9. If possible, restriction enzymes sensitive to CpG or CpNpG methylation or having star activity should be avoided in Southern analysis in order to prevent under- or over-digestion of plant DNA leading to under- or overestimation of the complexity of the transgenic loci. 10. Additional probes corresponding to vector backbone sequence directly outside the T-DNA left border (LB) or right border (RB) can also be used in Southern analysis to detect the potential transfer of small backbone sequences into the plant genome. 11. Factors affecting restriction digest, such as the presence of impurities in the DNA preparation, suboptimal quantity of enzyme, or digestion in small volumes, can lead to changes in banding patterns. For example, decreasing restriction enzyme (e.g., PmeI) quantities reduced the signal intensity corresponding to the intact HPT-GUS T-DNA fragment and proportionally increased high-molecular-weight band signals (Fig. 6). This suggests that some of the high-molecular-weight bands observed in complex banding patterns of Southern analyses could be artifactual when suboptimal restriction digest conditions are used. Finally, it implies that banding patterns should be used with caution, especially when considering the relative intensity of hybridization signals in the pattern, as they can be artifacts. 12. The concentrations of plant genomic and plasmid DNA used for reconstitution standards should be determined several (three or more) times. Imprecision in DNA concentrations will proportionally influence subsequent copy number determinations. It is important to note that for a given DNA sample, the concentration calculated using spectrophotometry

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will often be inferior (sometimes half) to the concentration determined using an ethidium bromide-stained gel analysis of plant DNA compared to commercially titrated DNA (such as lambda DNA). All DNA samples should therefore be analyzed using the same methodology and if possible at the same time. 13. Reconstitution standards are prepared by serial dilutions of plasmid DNA into wild-type genomic DNA so as to introduce between 1 and 40 transgene copies per plant genome equivalent. Plasmid DNA should be kept in T1/10E when diluted at low concentrations to prevent degradation. Two plasmid molecules are added to every wild-type plant genome to represent one transgene copy in a plant homozygous for the transgene (such as T2 plants for Event#2). When primary transformants (T0) are analyzed, only one molecule is added per plant genome, as all T0 plants are hemizygous for the transgene. For example in rice, 63 pg of the plasmid (5,792 bp) used in this study was equivalent to one gene copy for 5 µg of a T2 rice plant DNA (5,792 bp × 660 Da per bp × 1.65 × 10−24 g per Da = 6.3 × 10−18 g per plasmid. Five micrograms rice DNA/0.5 pg per rice genome = 107 genomes. 6.3 10−18 × 107 = 63 pg of plasmid for each 2C rice genome). Reconstitution standards corresponding to high copy number produce intense hybridization signals and can take a long time to decay even after chemical stripping of the membrane (see Note 8 for a protocol). More than one transgene can be added to each wild-type DNA in reconstitution standards provided that their lengths after restriction digest are sufficiently different to prevent signal interference. 14. It is useful to extract a large quantity of DNA from one (preferably two) well-characterized transgenic plant lines (as a positive control) as well as from a wild-type plant (as a negative control and as DNA to be mixed with plasmid DNA in reconstitution standards). The DNA should be divided into aliquots corresponding to the amount required for a single digest and stored at −20°C. This enables including the same quantity of DNA from the same source to all subsequent experiments/membranes to be analyzed by densitometry. When probing with a genomic RFLP probe, the signals obtained from both positive and negative controls can be used to normalize loading of the transgenic plant DNA. When probing with a transgene sequence, the variation in the positive control signals helps estimating the error in the copy number evaluation between membranes. 15. It is recommended to analyze at least twice the same transgenic plant within the same membrane (e.g., using two different restriction enzymes) in order to verify that the copy number determined by densitometry is similar in both cases.

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References 1. Vain, P. (2007) Thirty years of plant transformation technology development. Plant Biotechnol. J. 5, 221–229. 2. Zambryski, P., Joos, H., Genetello, C., Leemans, J., Van Montagu, M. and Schell, J. (1983) Ti-plasmid vector for the introduction of DNA into plant cells without alteration of their normal regeneration capacity. EMBO J. 2, 2143–2150. 3. Hiei, Y., Ohta, S., Komari, T. and Kumashiro, T. (1994) Efficient transformation of rice (Oryza sativa L.) mediated by Agrobacterium and sequence analysis of the boundaries of the T-DNA. Plant J. 6, 271–282. 4. Paszkowski, J., Shillito, R. D., Saul, M., Mandak, V., Hohn, T., Hohn, B. and Potrykus, I. (1984) Direct gene transfer to plants. EMBO J. 3, 2717–2722. 5. Shimamoto, K., Terada, R., Izawa, T. and Fujimoto, H. (1989) Fertile transgenic rice plants regenerated from transformed protoplasts. Nature 338, 274–276. 6. Gordonkamm, W. J., Spencer, T. M., Mangano, M. L., Adams, T. R., Daines, R. J., Start, W. G., Obrien, J. V., Chambers, S. A., Adams, W. R., Willetts, N. G., Rice, T. B., Mackey, C. J., Krueger, R. W., Kausch, A. P. and Lemaux, P. G. (1990) Transformation of maize cells and regeneration of fertile transgenic plants. Plant Cell 2, 603–618. 7. McCabe, D. E., Swain, W. F., Martinell, B. J. and Christou, P. (1988) Stable transformation of soybean (Glycine max) by particle acceleration. Bio-Technology 6, 923–926. 8. Vain, P. (2006) Global trends in plant transgenic science and technology (1973–2003). Trends Biotechnol. 24, 206–211. 9. Kohli, A., Leech, M., Vain, P., Laurie, D. A. and Christou, P. (1998) Transgene organization in rice engineered through direct DNA transfer supports a two-phase integration mechanism mediated by the establishment of integration hot spots. P.N.A.S. USA 95, 7203–7208. 10. Martineau, B., Voelker, T. A. and Sanders, R. A. (1994) On defining T-DNA. Plant Cell 6, 1032–1033. 11. Rommens, C. M., Humara, J. M., Ye, J. S., Yan, H., Richael, C., Zhang, L., Perry, R. and Swords, K. (2004) Crop improvement through modification of the plant’s own genome. Plant Physiol. 135, 421–431. 12. Vain, P., Worland, B., Kohli, A., Snape, J. W., Christou, P., Allen, G. C. and Thompson, W. F. (1999) Matrix attachment regions increase transgene expression levels and stability in transgenic rice plants and their progeny. Plant J. 18, 233–242.

13. Svitashev, S. K. and Somers, D. A. (2001) Genomic interspersions determine the size and complexity of transgene loci in transgenic plants produced by microprojectile bombardment. Genome 44, 691–697. 14. Cluster, P. D., Odell, M., Metzlaff, M. and Flavell, R. B. (1996) Details of T-DNA structural organization from a transgenic Petunia population exhibiting co-suppression. Plant Mol. Biol. 32, 1197–1203. 15. Afolabi, A. S., Worland, B., Snape, J. W. and Vain, P. (2004) A large-scale study of rice plants transformed with different T-DNAs provides new insights into locus composition and T-DNA linkage configurations. Theor. Appl. Genet. 109, 815–826. 16. Lu, H. J., Zhou, X. R., Gong, Z. X. and Upadhyaya, N. M. (2001) Generation of selectable marker-free transgenic rice using double right-border (DRB) binary vectors. Aust. J. Plant Physiol. 28, 241–248. 17. Vain, P., James, V. A., Worland, B. and Snape, J. W. (2002) Transgene behaviour across two generations in a large random population of transgenic rice plants produced by particle bombardment. Theor. Appl. Genet. 105, 878–889. 18. Svitashev, S. K., Pawlowski, W. P., Makarevitch, I., Plank, D. W. and Somers, D. A. (2002) Complex transgene locus structures implicate multiple mechanisms for plant transgene rearrangement. Plant J. 32, 433–445. 19. Son, D. Y., Ahn, K. M. and Lee, S. I. (2003) Sequencing, cloning and expression of CP4EPSPS roundup ready soybean insert. Food Sci. Biotechnol. 12, 133–136. 20. Spertini, D., Beliveau, C. and Bellemare, G. (1999) Screening of transgenic plants by amplification of unknown genomic DNA flanking T-DNA. Biotechniques 27, 308–314. 21. Strizhov, N., Li, Y., Rosso, M. G., Viehoever, P., Dekker, K. A. and Weisshaar, B. (2003) Highthroughput generation of sequence indexes from T-DNA mutagenized Arabidopsis thaliana lines. Biotechniques 35, 1164–1168. 22. Sallaud, C., Meynard, D., Boxtel, J. V., Gay, C., Bes, M., Brizard, J. P., Larmande, P., Ortega, D., Raynal, M., Portefaix, M., Ouwerkerk, P. B. F., Rueb, S., Delseny, M. and Guiderdoni, E. (2003) Highly efficient production and characterization of T-DNA plants for rice (Oryza sativa L.) functional genomics. Theor. Appl. Genet. 106, 1396–1408. 23. Vain, P., Afolabi, A. S., Worland, B. and Snape, J. W. (2003) Transgene behaviour in populations of rice plants transformed using a new dual binary vector system: pGreen/ pSoup. Theor. Appl. Genet. 107, 210–217.

Chapter 14 Fluorescent In Situ Hybridization to Detect Transgene Integration into Plant Genomes Trude Schwarzacher Abstract Fluorescent chromosome analysis technologies have advanced our understanding of genome organization during the last 30 years and have enabled the investigation of DNA organization and structure as well as the evolution of chromosomes. Fluorescent chromosome staining allows even small chromosomes to be visualized, characterized by their composition and morphology, and counted. Aneuploidies and polyploidies can be established for species, breeding lines, and individuals, including changes occurring during hybridization or tissue culture and transformation protocols. Fluorescent in situ hybridization correlates molecular information of a DNA sequence with its physical location on chromosomes and genomes. It thus allows determination of the physical position of sequences and often is the only means to determine the abundance and distribution of DNA sequences that are difficult to map with any other molecular method or would require segregation analysis, in particular multicopy or repetitive DNA. Equally, it is often the best way to establish the incorporation of transgenes, their numbers, and physical organization along chromosomes. This chapter presents protocols for probe and chromosome preparation, fluorescent in situ hybridization, chromosome staining, and the analysis of results. Key words: Chromosome, fluorescent microscopy, physical mapping, biotin, digoxigenin DAPI.

1. Introduction Molecular cytogenetics investigates the properties of chromatin and chromosomes and analyses the physical organization of DNA sequences along chromosomes or within interphase nuclei. With the development of non-radioactive hybridization methods and fluorescent microscopy, multi-colour analysis has become possible and has greatly advanced chromosome research. In fluorescent chromosome banding (1) (see Fig. 1a), fluorescent dyes bind directly to DNA either uniformly or specifically to base pairs Huw D. Jones and Peter R. Shewry (eds.), Methods in Molecular Biology, Transgenic Wheat, Barley and Oats, vol. 478 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-379-0_14

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Fig. 1. Fluorescent analysis of chromosomes. (a) Fluorescent chromosome banding: root tip metaphase chromosomes of a triploid hybrid desert banana with 33 small chromosomes stained with DAPI (blue under UV-excitation). The centromeric heterochromatin of all chromosomes fluoresces more strongly. Some cytoplasm is visible as weak fluorescence between chromosomes. (b) Example of FISH with a highly repetitive DNA family probe (labelled with biotin d-UTP and detected with streptavidin conjugated to Alexa594, red fluorescence) to rye metaphase chromosomes. The chromosome-specific distribution pattern identifies all seven chromosome pairs for details of probes (see (2). Bar 7µm. (See Color Plate 7 )

and certain fractions of chromosomes, and different types of heterochromatin can be differentiated and some information about the DNA sequences within chromosomes can be elucidated (3). As the unbound dye is essentially non-fluorescent and its fluorescence is a magnitude greater on binding to DNA, even small chromosomes and those surrounded by cytoplasm and cell walls can be analysed. Considerably more information is gained by fluorescent in situ hybridization (FISH), where labelled probes are hybridized to chromosomes and nuclei in situ (4) (see Fig. 1b) and where the cytological information about chromosomal organization can be directly combined with molecular data about DNA sequence (5, 6). FISH is used for mapping of DNA sequences to their physical location within the genome, for correlating linkage groups to specific chromosomes and providing markers for chromosomes or chromosome segments to allow chromosome identification, and for the determination of chromosomal rearrangements (6–8). In plant molecular biology, FISH has proved particularly valuable to understand the large-scale organization of genomes and chromosomes and of chromosome regions that are characterized by repetitive sequences and to follow their diversity and evolution (2, 9–12). Many of the answers obtained from chromosomal FISH are difficult to discover using any other method: restriction digests of repeated sequences show one or more bands in gel electrophoresis, which are difficult to assign to loci, and sequencing projects are not able to access long and relatively homogeneous stretches of repetitive sequences. The method also does not require segregation genetics, which is impossible to carry out in sterile crops and difficult in long-life-cycle species including trees. Research in understanding genome organization and the three-

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dimensional spatial distribution of DNA sequences at interphase and meiosis has benefited from the advent of FISH and other molecular cytogenetic methods of investigation, chromatin organization, and modelling (13, 14). While single-copy in situ hybridization to show the physical location of a gene in mammalian, and particularly human, research frequently complements genetic data, in plants it remains a specialized technique (15, 16). In plants, wellcharacterized single-copy sequences as short as 800 bp have been localized, but 2.5 kb or longer sequences are easier to work with, and more than 40 kb of target sequence is required for reliable hybridization to both chromatids of all chromosomes carrying the locus; otherwise only statistical analysis of several metaphases will identify the location of the hybridized probe. The detection and identification of the chromosomal positions of transgenes (17–20, 30) or viral sequences (21–23) incorporated into host genomes represents a unique situation where the considerable effort required for FISH analysis of low or single-copy sequences are justified for several reasons. First, the univocal identification of whether an alien DNA molecule has indeed been incorporated into the genome is very difficult (sometimes even impossible) by Southern hybridization or by polymerase chain reaction (PCR); second, the chromosomal position of the integrated transgene by mapping would require the generation of a segregating population for every transgenic line; third, vectors carrying transgenes or viral sequences are several kilobase long and, if integrated in tandem, provide an acceptable although relatively small target for FISH. While it has been increasingly recognized that the location of these sequences within the genome can have a major influence on the level and stability of their expression and hence is important to know (23), a further very important justification of embarking on chromosomal analysis of transgenic lines is the need to establish whether they have maintained their chromosomal integrity or whether aneuploidies, polyploidies, or rearrangements are present. In situ hybridization to chromosomes was first described using radioactively labelled satellite DNA (24, 25). Now in situ hybridization to chromosomes almost entirely uses non-radioactive methods based on fluorescence that allow more accurate localization of the probe and the use of several probes simultaneously (4). The biotin and digoxigenin systems have found by far the widest applications as indirect labels for FISH and are detected by fluorophore-conjugated avidin (or its derivatives) or antibodies. Alternatively, direct methods use fluorescent labelled nucleotides that do not need a special detection step, making the method faster and normally cleaner, but at the same time slightly less sensitive. Double-target FISH, with red and green fluorescence for detection of probes and DAPI as bluefluorescing counterstain for chromosomes, is routinely used in our

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laboratory, while three- or four-target hybridization experiments, with addition of blue or far-red or intermediate fluorophores are used more widely in human and mammalian cytogenetics. Ratiolabelled probes, where some probes carry more than one label (mixed either during or after separate labelling reactions), can be used with single-labelled probes to increase the number of targets detected in both animals (26, 27) and plants (28). In this chapter, preparation and staining of chromosomes will be described that can be used for assessing the chromosomal integrity and polyploidy after transformation or for FISH experiments. Some discussion of probe preparation and labelling is given and, finally, the steps of FISH to detect transgenes and how to capture and analyse the images are described.

2. Materials 2.1. Fixation of Material

1. Metaphase arresting agent: choose one of the following. (a) Ice water (typically for seedlings of temperate plants): fill a 5- to 15-ml tube two-thirds with fresh or bottled water (see Note 1); shake vigorously to aerate, keep at−20°C until the water starts to freeze, shake again, and store in an ice bucket. (b) 8-Hydroxyquinoline (for many species): make a 2 mM solution in water (pale yellow, taking 12 h to dissolve); store in dark at 4°C for up to 6 months. (c) Colchicine (for most plants): prepare a 0.05–1% (v/w) solution in water; store in dark at 4°C for up to 1 month. 2. Fixative: freshly prepared (less than 30 min old). Three parts 96% ethanol to one part glacial acetic acid. 3. Small tubes (5–10 ml) with tight screw caps (e.g. bijou tubes or freezer vials).

2.2. Chromosome Preparation

1. Enzyme buffer: prepare a 10× stock solution at pH 4.6 by mixing four parts 100 mM citric acid with six parts 100 mM tri-sodium citrate. For use dilute 1:10 with distilled water. 2. Enzyme solution: 2% (w/v) cellulase from Aspergillus niger (Calbiochem, 21947, 4,000 units/g) or a mixture of 1.8% Calbiochem and 0.2% “Onozuka” RS cellulase (5,000 units/g) and 3% (v/v) pectinase from Aspergillus niger (solution in 40% glycerol, Sigma P4716, 450 units/ml). Make up in 1× enzyme buffer. Store in 2–5 ml aliquots at−20°C (see Note 2). 3. Acetic acid: 10–20 ml 45% (v/v) and 60% (v/v) acetic acid in water.

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4. Microscope slides: soak in chromium trioxide solution in 80% (v/v) sulfuric acid for 3 h or more, wash in distilled water, and air dry; wipe with 96% alcohol before use (see Note 3). 5. Glass microscope coverslips: 18 mm× 18 mm size, No. 1 (see Note 4). 6. Dissecting needles and fine forceps (e.g. No. 5). 7. Dry ice or liquid nitrogen. 2.3. Probe Labelling

1. Labelling kit: based on random priming or nick translation (e.g. Invitrogen, Roche). 2. Labelled nucleotides (if they are not included in the labelling kit): dixoginen d-UTP from Roche, or fluorescent conjugated d-UTPs or CTPs from Amersham/GE Healthcare or Molecular Probes. 3. Purification tubes to remove unincorporated nucleotides, unwanted enzymes, and salts unless included in the kit (see Note 5).

2.4. Pre-treatment of Slide Preparations

1. 2× SSC: diluted from 20× stock solution (20× is 3 M NaCl, 0.3 M sodium citrate, adjusted to pH 7). 2. RNase solution: 10 mg/ml in 10 mM Tris–HCl, pH 8; store 0.5–1 ml aliquots at−20°C; dilute with 2× SSC to 100µg/ml before use. 3. 10 mM HCl. 4. Pepsin solution (optional see Note 6): make up stock of 500µg/ ml (approximately 4,000 U/mg) in 10 mM HCl; store 0.5–1 ml aliquots at−20°C; dilute to 1–10µg/ml for use. 5. Ethanol series: 96%, 80–90% (v/v) and 70% (v/v) in water. 6. Paraformaldehyde: in the fume hood, add 4 g paraformaldehyde (electron microscope grade) to 80 ml water and heat to 60°C for about 10 min, clear the solution with a few drops of concentrated (10 M) NaOH, let cool down to below 30°C before use, and adjust pH to 8 with 1N H2SO4. 7. Plastic coverslips: 40 mm× 25 mm pieces from autoclavable plastic bags of the type used for contaminated waste. 8. Coplin jars holding eight slides and 50–100 ml solution.

2.5. Hybridization Mixture, Denaturation, and Hybridization

1. Formamide: store 0.5–1 ml aliquots in at −20 °C (see Note 7). 2. Dextran sulfate: 50% (w/v) solution in water, heat to dissolve and sterilize by forcing through a 0.22µm filter; store 0.5–1 ml aliquots at −20°C. 3. SDS solution: 10% (w/v) sodium dodecyl sulfate (also called sodium lauryl sulfate) in water, filter, sterilize; store at room temperature.

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4. Salmon sperm DNA: 1µg/ µl sonicated or autoclaved DNA (also suitable are herring sperm or E. coli DNAs); store 0.5–1 ml aliquots at −20°C. 5. 20× SSC solution: 3 M NaCl, 0.3 M sodium citrate, adjusted to pH 7, filter (0.22µm), sterilize if it has not been autoclaved; store 0.5–1 ml aliquots at −20°C. 6. EDTA: 100 mM, pH 8; store 0.5–1 ml aliquots in at −20°C. 7. Plastic coverslips: 25 mm× 25 mm pieces from autoclavable plastic bags of the type used for contaminated waste. 2.6. Stringent Washes and Detection of Probe

1. 2 × SSC and 0.1 × SSC: dilute from 20 × stock (see Section 2.4); prepare 500 ml each and pre-heat to 45°C. 2. Formamide: store 40 ml aliquots at −20°C (see Note 7). 3. Stringent wash solutions: prepare 100 ml of 20% formamide/ 0.1 × SSC for high stringency washes, or 20% formamide/ 2 × SSC for low stringency washes (see Note 8). 4. Detection buffer: 4× SSC containing 0.2% (v/v) Tween-20; prepare 500 ml and preheat to 45°C. 5. BSA block: 5% (w/v) bovine serum albumin in detection buffer. 6. Detection reagent: prepare 50µl detection reagent per slide (see Note 9). Where two labels are being detected, the two reagents are mixed in the detection buffer. – Digoxigenin labels: anti-digoxigenin (FAB-fragment) conjugated to a suitable fluorochrome (Roche); typically 1:200 dilution in detection buffer (see Note 10). – Biotin labels: avidin, streptavidin, extra-avidin, or antibiotin conjugated to suitable fluorochrome (e.g. Vector, Sigma, Molecular Probes, Roche); typically 1:200–500 dilution in detection buffer (see Note 10). 7. Plastic cover slips: 25 mm × 40 mm pieces cut from autoclavable plastic bags of the type used for contaminated waste. 8. Staining jars for washing slides, containing about 100 ml of solution.

2.7. Counterstaining and Mounting of Slides

1. DAPI staining solution: 2′, 6-diamidino-2-phenylindole; prepare a stock of 0.1 mg/ml in water; store at−20°C; dilute to 2–4µg/ml in McIlvaine’s buffer pH 7 (18 ml of 100 mM citric acid and 82 ml 200 mM Ha2HPO4) and store 1 ml aliquots in −20°C. 2. Antifade reagent: commercially available, e.g. Citifluor AF1 (Agar), Vectashiled (Vector), Slow Fade (Molecular Probes), or Fluorguard (Sigma). 3. Glass microscope cover slips: 24 mm× 30 mm or 40 mm No. 0 (see Note 4).

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3. Methods The basic procedures of in situ hybridization (see Fig. 2) are derived from those used for Southern hybridization: the targets are the chromosomes and nuclei prepared on a microscope slide, while the probes are the labelled DNA sequences to be detected (in the case of this book the transgene and any controls, Fig. 3a). The probe and chromosomes are denatured (either separately or together) and are allowed to form hybrid molecules (Fig. 2). Most protocols use an overnight hybridization step that is essential for low-copy FISH, while for repetitive DNA sequences a few hours are sufficient. As with Southern hybridization, several washes are performed to remove unbound probe and to determine the stringency of hybridization – the similarity between probe and target DNA sequences which is required before they can remain stably hybridized in a double-stranded DNA helix. For chromosome preparations, stringency is controlled by formamide (a helix de-stabilizing agent) in addition to the temperature, and sodium ion concentration in the hybridization and washing solutions. In direct FISH, chromosomes can be immediately counterstained,

Fig. 2. Flow diagram of fluorescence in situ hybridization analysis.

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Fig. 3. FISH analysis and some common background problems. Wild Petunia metaphase chromosomes (2n = 14) after FISH with an endogenous pararetrovirus, EPRV probe (labelled with biotin d-UTP and detected with streptavidin conjugated to Alexa594, red fluorescence under green excitation (for probe description see (22) and a control probe (labelled with digoxigening d-UTP and detected with anti-digoxigenin conjugated to FITC, green fluorescence under blue excitation). Chromosomes are counterstained with DAPI (blue fluorescence under UV excitation). For each panel, the three fluorescent images were recorded with a digital camera and overlaid using Adobe Photoshop. (a) Well-spread metaphase chromosomes with little cytoplasm and background hybridization allows clear identification of FISH signal: four 5S rDNA sites (green) and two major (arrow heads) and two minor (arrows) EPRV sites (red) are detected at the centromere of four different chromosome pairs. The minor EPRV sites show an irregularly shaped “double dot” signal, one on each chromatin, indicating a few integrated copies, while the major sites resemble large bands of several tens of integrated virus copies. Unspecific ERPV signal next to the chromosomes (X) or appearing as perfect sphere with a hallow (*) can be discarded. (b) Background fluorescence to cytoplasm obscures the FISH signal. Strong perfect double dots (**) at an unusual terminal position with no match at the homologous chromosome are a false site. (c) Strong unspecific background fluorescence on and next to chromosomes obscures the proper signal. (d) Strong unspecific star-like dots of red and green fluorescence indicate that the probes have disintegrated causing aggregation of label. Weak FISH signal of the green control probe to the centromeres of most chromosomes is still visible. Proper red EPRV signal is not detectable. Note that unspecific background signal tends to be spherical with clear edges, while the proper hybridization signal is made up of many small dots resulting in an uneven appearance. Compare the minor EPRV signal in (a) to the accidental double dots in (b) and the major site in (a) with the unspecific large dots in (d). Bar: 10µm. (See Color Plate 8 )

most often with DAPI, while indirect FISH with biotin and digoxigenin labels needs a detection step using immunocytochemistry principles. For fluorescent staining of chromosomes (Fig. 1a), to check ploidy level of lines and individuals follow protocols of

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fixation and chromosome preparation (see 3.1 and 3.2) followed by counterstaining, mounting, and analysis (see 3.7 and 3.8). 3.1. Fixation of Material

A critical factor contributing to the success of FISH and chromosome staining is the preparation of clean chromosome spreads with plenty of metaphases. Plant material used for fixation and chromosome preparations needs to be healthy, disease free, and growing rapidly. It can be any dividing tissue, but the time of analysis during the growth of transgenic plants may determine which material is best: root tips from young seedlings, freshly appearing roots at the edge of pots of older plants, or hydroponic cultures. For some species, meristematic cells from young shoots, leaves, or emerging buds can also be used. For most application it is important to have maximum number of metaphase chromosomes. To accumulate metaphases, condense the chromosomes and destroy the spindle microtubules to allow better spreading of the chromosomes, and the tissue is usually pre-treated. The spindle microtubule inhibitor colchicine (widely used for chromosome counting) gives heavily condensed chromosomes, while other chromosome condensation agents such as ice water or 8-hydroxyquinoline give more extended chromosomes that are more suitable for accurate FISH analysis. 1. All steps are carried out in clean vessels with clean forceps. During collection and pre-treatment (steps 2–4), avoid contact of growing material with fixatives and chemicals (see Note 11). 2. Choose suitable plant material from which to obtain metaphases. Make sure these have been growing under optimum conditions. 3. Prepare chosen metaphase arresting agent as described under Section 2.1 and transfer to tubes. 4. Transfer roots (1–2 cm long) or buds to 3–5 ml of metaphase arresting agent (see Note 12). As a guide, use about five times as much solution as material. Screw cap on tightly. 5. Incubate as follows: – Ice water: 24 h. – 8-Hydroxyquionoline: at temperature of plant growth for 30–45 min and then at 4°C for 30–45 min. – Colchicine: 2–4 h at room temperature or 12–16 h at 4°C. 6. Quickly blot the material dry and plunge it into fresh fixative. Make sure that the fixative is not contaminated with water for quick penetration. In addition to labelling tubes with a felt

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pen, use pencil on a sticker, or a piece of paper that can be submerged into the fixative. 7. Leave for 1 h at room temperature before storing at 4°C or−20°C (see Note 13). 3.2. Chromosome Preparation

For both fluorescent staining and FISH, metaphase chromosome and interphase preparations are made on glass microscope slides. Pre-treatment with proteolytic enzymes is recommended to remove cell walls and cytoplasm prior to squashing the material in acetic acid between a cover slip and glass slides. All steps are carried out at room temperature unless stated otherwise. Use small Petri dishes for the washing steps and prepare material as described under Section 2.2. 1. Wash fixative from plant material twice in enzyme buffer for 10 min. 2. Transfer plant material to 1–2 ml enzyme solution and incubate at 37°C for 30 min to 2 h, depending on the material (see Note 2). 3. Transfer plant material from enzyme solution to enzyme buffer. 4. Transfer material for two or three preparations to 0.5 ml 45% acetic acid and leave for a few minutes. If the material is very soft, this step can be omitted. 5. Dip the microscope slide into 96% ethanol and dry with a lintfree tissue, place a drop of 60% acetic acid on the slide, and put a small piece of root tip, anther, or other material into the drop. Change the acetic acid if necessary. 6. Under a dissecting microscope, tease apart the tissue or tap with a glass rod; remove any particles. Metaphases tend to float free. 7. Wipe a cover slip with a tissue and place onto the preparation. Examine with a phase-contrast microscope. Squashing with light to very heavy hand pressure might be required, or the cover slip can be tapped to give the optimum preparations. 8. Check slides under the microscope for cell density and metaphase index, and put good preparations on a metal plate on a dry ice block for 5–10 min (not more); then flick off coverslip with a razor blade edge. Allow preparation to air dry. 9. Screen slides, without mounting, using a phase contrast microscope. Look for well-spread nuclei with no clumps, lack of surface film, and no dirt. Metaphase chromosomes should be well separated and free of cytoplasm, debris, scratches, and dirt. Take notes about each slide to refine the preparation method next time. Small chromosomes can be checked by staining with DAPI (see Section 3.7)

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10. Mark the position of the preparation by scratching lines on the glass slide. The position of the preparation will not be visible during later stages on the wet slides. 11. Suitable slides can be stored desiccated at 4°C for a few days or for months at−20°C (see Note 13). 3.3. Probe Labelling

Protocols for labelling DNA are not given here, as suitable instructions are included by the manufacturers in the kits based on random priming or nick translation. Also frequently used are direct PCR labelling methods that amplify specific sequences from cloned or total genomic DNA sequences while incorporating labelled molecules. For successful FISH, the probe must incorporate a suitable proportion of label molecules to allow detection. In order for polymerases to work efficiently, the labelled nucleotide (d-UTP or d-CTP) is mixed in a 1:2 ratio with unlabelled TTP or CTP. Manufacturers of labelled nucleotides usually make recommendations of the concentrations and dilutions that give optimal incorporation; these are a good starting point, but the absolute amounts of very expensive labelled nucleotides can often be reduced by 50–70%, particularly with fresh reagents that have not been through multiple freeze–thaw cycles. A further crucial factor is the length of the probe after labelling, and it must not be too long (typically 200–600 bp) to allow penetration to the DNA packaged within the chromosome. Random priming and nick translation will automatically result in probe lengths suitable for FISH, but sometimes the DNAse component in the nick translation enzyme mixture might need adjusting, and PCR labelling of larger insert clones will not result in good probes. Further important factors for successful labelling are the purity of the template DNA and the lengths of the DNA template sequence to be used for labelling. If a cloned sequence is used, make sure that the miniprep DNA is clean without bacterial contamination and that the insert can be cut out cleanly. For shorter inserts (100 bp to 2 kb), PCR amplification using the universal M13 sequencing primers is recommended and will yield very clean template DNA. It is best to run the PCR product on a gel and to cut out and purify the correct band prior to labelling by random priming or nick translation. For longer products, the miniprep DNA can be directly labelled after linearizing the plasmid or cutting the insert out. However, we have found that large DNA molecules do not often give good templates possibly because the polymerases become inefficient if not are stopped by the end of DNA template molecules. Therefore cutting large DNA molecules by sonication, heating, or enzyme digestions is favourable before labelling (see (4, 19)). For transgene detection, I recommend the use of biotin labelling (see Fig. 3a), as biotin is the smallest hapten and generally the

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most easily incorporated, and specific biotin kits are available. Secondly, many different avidin, streptavidin, or antibiotin antibodies are available linked to very strong and durable fluorochromes (e.g. Alexa dyes, Molecular Probes). As a second control or reference probe, labelled with digoxigenin or direct fluorophores, I recommend the use of the 5S rDNA, as it is universal and in many species has major and minor sites (see Fig. 3a) that provide excellent quality control of the FISH experiment. 45S rDNA is also acceptable, but often has such strong sites that it overpowers the transgene hybridization signal. Alternatively, repetitive DNA sequences that allow the identification of chromosomes can be very useful. 3.4. Pre-treatments of Slide Preparations

Before the in situ hybridization procedure, the slide preparations are pre-treated to enhance probe access to the target sites (e.g. by removing surface proteins with pepsin or protease treatments), and to reduce non-specific probe and detection reagent binding (e.g. RNase treatment and pepsin/protease treatments). The preparation is then stabilized by refixation in paraformaldehyde and alcohol, which also helps to retain the preparation during the many washes. 1. Steps are carried out in coplin jars at room temperature unless otherwise stated. 2. Add 200µl RNase of solution to the marked area on each slide. Cover with a plastic coverslip and incubate for 1 h in a humid chamber. 3. Remove the cover slip and wash slides twice for 5 min each in 2× SSC. 4. Place slides in 10 mM HCl for 2 min, shake fluid from each slide and quickly add 200µl of pepsin solution to each slide, cover with a plastic coverslip, and incubate for 10 min at 37°C (see Note 6). 5. Wash off cover slips in distilled water and then wash twice for 5 min in 2× SSC. 6. Place slides in a coplin jar with paraformaldehyde fixative for 10 min in a fume hood. 7. Wash twice for 5 min in 2× SSC. 8. Dehydrate through an ethanol series (70%, 90%, 96%, 2 min each) and allow to air dry.

3.5. Hybridization Mixture, Denaturation, and Hybridization

1. In a microcentrifuge prepare the hybridization master mixture as described in Table 1. Mix well. 2. For each probe mixture, add 34µl master mix and 2µl transgene probe and 1µl reference or control probe to a centrifuge tube, and make up to 40µl with distilled water.

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Table 1 Hybridization master mix Component

Final concentration in hybridization mix

Amount for one slide (µl)

Amount for eight slidesa (µl)

100% Formamide

50%b

20b

180b

20× SSC

2×

4

36

50% Dextran sulfate

10%

8

72

Salmon sperm DNA 1µg/µl

0.025µg

1

9

100 mM EDTA

1.25 mM

0.5

4.5

10% SDS

0.125%

0.5

4.5

34

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Total a

Prepare master mix for one more slide than needed. Hence numbers given here are amounts for one slide multiplied by nine b For low stringency, use 40%, 16µl, and 134µl, respectively, and add water accordingly to make up the volume

3. Denature probe mixtures in a water bath at 70°C for 10 min and place in ice for 5 min. 4. Place 40µl of probe mixture over the area of preparation on the slide and cover preparations with a plastic coverslip, taking care to remove any bubbles by lifting and replacing the coverslip. 5. Place slides onto a heating block and raise temperature to that required for denaturation. Typical time and temperature are 75°C and 8 min. Temperatures between 70 and 95°C and times from 6 to 12 min may be used depending on the species, method of preparation, and storage time of material before and after making the preparation ( see Note 14 ). 6. Lower the temperature to 37°C over 10–20 min and maintain it at 37°C for the hybridization. 7. Leave slides on the heated block if they can be maintained in a moist atmosphere, or transfer to a humid box in an oven or floating in a water bath for hybridization. Be careful that preparations do not dry out, nor accumulate drops of condensation on them. Leave overnight (16 h). 3.6. Stringent Washes and Detection of Hybridization Sites

1. Prepare wash solutions and detection buffer as described in Section 2.6. 2. Remove slides from hybridization chamber and check that they have neither dried out nor become wet from condensation. 3. Float off coverslips in 2× SSC at 42°C; allow coverslips to fall away. Be careful not to scratch slides against each other.

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4. Wash in 2× SSC at 42°C for 2 min. 5. Incubate slides twice for 5 min in the stringent wash solution at 42°C. Measure the temperature of the wash solution with slides accurately and record. Use 0.1× SSC for high stringency or 2× SSC for low stringency (see Note 8). 6. Wash slides three times in 2× SSC at 42°C. 7. Take slides out of the water bath and allow them to cool for 10–15 min. 8. Transfer slides to detection buffer for a minimum of 5 min. 9. Apply 200µl BSA block to the marked area of each slide, cover preparation with a plastic cover slip, and leave for 5–30 min. 10. Prepare detection solution depending on probe label used (see Section 2.6). Shake off BSA block and place 50µl of detection solution onto the slides, cover with a plastic cover slip, and incubate at 37°C in a humid chamber for 60 min (see Note 9). 11. Remove cover slips and wash in detection buffer for 3× 5 min. 3.7. Counterstaining and Mounting

Most preparations require a DNA counterstain to visualize the chromosomes. It is important that counterstaining fluoresces in a colour not used for probe detection and that it is not too bright so that hybridization sites are not obscured; hence staining and then mounting is recommended. Some protocols mix the counterstain with the mountant, which saves one stage of preparation, but this can lead to over-staining and high backgrounds. The first fluorescein FISH experiments used propidium iodide that stains chromosomes red when excited with blue or green light. For double-target FISH, when red and green are used to visualize fluorescent probes, DAPI that fluoresces blue under UV excitation (Fig. 3) is ideal, as it also often reveals heterochromatin banding. DAPI is also an ideal stain to analyse chromosome preparations for checking their quality and counting chromosomes, in particular if they are small (Fig. 1a). 1. Place a drop (50–100µl) of staining solution onto the slide and cover with a plastic coverslip, and incubate at room temperature in the dark for 10 min (see Note 9). 2. Remove the coverslip and wash the slide briefly in detection buffer (see Section 4.2.6). 3. Drain buffer and place a drop of antifade mountant onto the slide surface and apply a glass cover slip. Squash cover slip with high hand pressure between sheets of filter paper to squeeze out excess mountant (see Note 15). 4. Observe slide. The antifade may take a few hours to penetrate preparations, so wait overnight before detailed observation and photography.

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An epi-fluorescence microscope is used to visualize sites of probe hybridization and chromosome staining. While this chapter does not aim to give instructions for the use of the microscope, some general problems and guidelines are given below. 1. The location of the microscope in a completely dark room, on a stable surface and set-up for comfortable operation is essential, as fluorochromes and in situ hybridization signals may be very weak and fade rapidly even when viewed with good antifade mountants. Make sure that the system is set up correctly and that the lamp is properly centred. 2. The optics and immersion oil used for visualization must be specified for fluorescent applications including UV. Immersion oil must be kept in the dark at room temperature: heating in the sun in a salesman’s car, mixing oils, or absorption of water will make oils autofluorescent or UV opaque. 3. The most useful filter sets in most applications are those for individual fluorochromes; multi-bandpass sets are available, but often the brightness of the different fluorochromes is not matched enough to make them useful, and cross-excitation of the different fluorochromes will often make analysis of results difficult. Make sure filters have not degraded through use. 4. Use film or a digital camera system for capture of images. Both give resolution suitable for publication. However, it must be noted that a digital system, no matter how expensive, cannot make up for deficiencies in the set up of the microscope. Where film is used, colour print film is usually preferable to slide (reversal) film since it has higher exposure latitude and is cheaper to buy and print than slide films. 5. Always mount slides in antifade solution, as this greatly prevents fading. 6. When viewing slides, be extremely careful not to fade the chromosomes and in particular the weak fluorescent FISH signal before you have recorded it. Low-power lenses will generally not fade the signal but when viewing under 63× and 100× fading is rapid and it is necessary to set up the camera system beforehand to be ready to press the exposure button as soon as the field is in focus. Photograph the FISH signal first and avoid viewing your chromosomes with DAPI before you have taken the FISH signal and are satisfied with its quality. 7. When presented with a very weak FISH signal that is almost invisible to the naked eye, focusing your image can be difficult. You can normally focus red and green fluorescence interchangeably, but DAPI will not allow you to do the same. 8. Whether digital cameras or film is used, it is now most convenient to use a microcomputer-based image-processing program (e.g. Adobe Photoshop, Corel Draw, or one of the

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equivalent packages) to analyse, overlay, and arrange images. However, care must be taken not to over-process images, and supervisors must be well aware that they are responsible for the supervision of image processing by their laboratory members! 3.9. FISH Analysis of Transgenes

Probably one of the most difficult parts of your experiment is to decide whether the signal particularly from a low-copy transgene, you have observed or recorded is real. 1. Check that metaphases are complete and that the morphology of the chromosomes is preserved. Chromosomes can be a little swollen, but should not have a ghost-like, blown-up appearance; they should not be frayed, have holes, or be distorted. 2. Make sure that the control probe has resulted in the expected signal. 3. Investigate background signal and unspecific cross-hybridization that obscure the real FISH signal (see Fig. 3b and c). Any outof-focus signal that produces a clear hallow can be discarded (see Fig. 3a). Very strong star-like dots (see Fig. 3d) are often the indication that probes or detection reagents have deteriorated. 4. Count the sites of transgene FISH signals that are located on chromosomes. Analyse several metaphases and, if possible, identify the chromosome and position of FISH signal. In a normal diploid cell, a disomic locus should give a signal on both chromatids of both homologues at the same position (see Note 16). When looking for a target of less than 50 kb, it is unlikely that all present sites will be visible in the same metaphase because of coiling and squashing, so be prepared not to see the maximum number of sites in any given metaphase. However, looking at 5–10 metaphases or more should confirm which is real signal and which is the background. Strong, reliable signal will indicate that several copies of the transgene have integrated in tandem (18,19 (see Fig. 3a). 5. Most importantly, compare slides with the same probe mixture or detection combination to exclude failure, contamination or inefficiencies in probe labelling, antibody dilution, or other reagents used.

4. Notes 1. Water quality for seed germination and ice water treatment is very important. Chlorines or amines used for water purification or heavy metals from pipes reduce the metaphase index; so the use of bottled water is recommended.

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2. Modifications such as addition of pectolyase (0.1–1% solution) or viscozyme (0.1–0.5%), or alteration of ratios of different enzymes, may be necessary to obtain optimized preparations from different species. 3. High-quality glass slides are essential for clean preparation; treatment with strong acid removes not only oils but also surface anions and allows cells and chromosomes to stick to the slide; test cleaned slides under phase contrast to avoid those that have impurities. If chromic acid is not available, use 6N HCl. Slides with specially treated surfaces might also be useful; however, some will cause increased background and some are expensive. 4. The optimum preparations are often near the periphery of the cover slip. Therefore, use a small (e.g. 18× 18 mm) coverslip for making preparations (Sections 2.2 and 3.2), and larger (e.g. 24× 30 or 40 mm) coverslips for mounting (Sections 2.7 and 3.7). 5. Ethanol precipitation using sodium acetate or lithium chloride can be used instead of a purification column. 6. This step can be omitted if the preparation is clean and individual chromosomes can be identified with phase contrast. Adjust the concentration to the material. 7. A good grade, but not the highest, is required. To avoid degradation in opened bottles, separate into aliquots of 1 ml for the hybridization mix (Sections 2.5 and 3.5) and 40 ml for stringent washes (Sections 2.6 and 3.6) before freezing at−20°C. If the aliquots do not freeze completely, then there are impurities, and another grade or fresher batch should be used. 8. For low-copy transgenes, reduced hybridization stringency is recommended, although cross-hybridization to false target has to be monitored. 9. Most fluorochromes fade very rapidly in light (the energy destroys the molecule) and must be kept away from bright lights while preparing solutions and when on slides. 10. Dilutions of antibody vary widely between batches even from one manufacturer and must be optimized in any application. Typical dilutions vary from 1:50 to 1:600. 11. Traces of fixatives on glassware, tools used to handle plant material, or in the atmosphere of, for example, refrigerators will reduce greatly the metaphase index. Do not cross-contaminate other plant material with the fixative or fixative vapours. 12. When using roots from freshly germinated seeds and the material is not needed for subsequent planting, the whole seed is preferably used for ice treatment and fixation.

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13. Fixed material can be stored for several weeks to months before making chromosome preparations as long as it does not get warm. But even when stored cold, it tends to get hard and it becomes difficult to remove cytoplasm. Storing or shipping ready-made chromosome slide preparations (Section 3.2) may sometimes be the better option when FISH has to be done later or in another remote lab. 14. This step is critical to ensure that all chromosomal DNA is denatured; but if too long or at too high temperature, chromosomes will lose their morphology. Many makes of PCR machines have modifications for holding slides (29) and these are particularly good at enabling the temperature and time to be controlled accurately. Alternatives include use of thermostatted heated plates or placing slides in a tray heated in a waterbath; in both cases the slide temperature must be accurately monitored by placing a thermometer next to the preparations. Some protocols denature the slide preparations separately from the probe by dipping into a 70% formamide, 2× SSC mixture at 60–80°C for 6–10 min, before applying the hybridization mixture and coverslip. 15. For long-term storage or transport, cover slips can be sealed with gum or nail varnish. If fluorescence becomes very faint, particularly after long storage, remove the cover slip, wash slide briefly in detection buffer, and remount (sometimes restaining with DAPI may be necessary). 16. Be aware that some transgenes might only be present on one of the two homologues or might have integrated at several locations.

Acknowledgements I would like to thank John Bailey, University of Leicester, for help in refining our laboratory’s FISH protocols, and Pat Heslop-Harrison for reading the manuscript and continuous discussion. Chee How Teo and Alessandra Cotento from my lab, and Katja Richert-Poeggeler, Federal Biological Research Centre for Agriculture and Forestry (BBA), Braunschweig, Germany, are acknowledged for letting me use figures from our joint research projects. Support is acknowledged from the EU-FP5 network PARDIGM QLK3-CT-2002–02098, Generation Challenge Programme and FAO/IAEA Coordinated research Projects.

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References 1. Schweizer, D. (1981) Counterstain-enhanced chromosome banding. Human Genet. 57, 1–14. 2. Contento, A., Heslop-Harrison, J. S. and Schwarzacher, T. (2005) Diversity of a major repetitive DNA sequence in diploid and polyploid Triticeae. Cytogenet Genome Res. 109, 34–42. 3. Kowalska, A., Bozsaky, E., Ramsauer T., Rieder, D., Bindea G., Lorch T., Trajanoski Z. and Ambros P. F. (2007) A new platform linking chromosomal and sequence information Chromosome Res. 15, 327–339. 4. Schwarzacher T. and Heslop-Harrison J. S. (2000) Practical in situ Hybridization. Bios, Oxford, 213 + xii. 5. Schwarzacher, T. (2003) DNA, chromosomes, and in situ hybridization. Genome 46, 953–962. 6. Kato, A., Vega, J. M., Han, F., Lamb, J. C. and Birchler, J. A. (2005) Advances in plant chromosome identification and cytogenetic techniques. Curr. Opin. Plant Biol. 8, 148–154. 7. Hasterok, R., Marasek, A., Donnison, I. S., Armstread, I., Thomas, A., King, I. P., Wolny, E., Idziak, D., Draper, J. and Jenkins, G. (2006) Alignment of the genomes of Brachypodium distachyon and temperate cereals and grasses using BAC landing with fluorescent in situ hybridization. Genetics 173, 349–362. 8. Forsström, P. O., Merker, A. and Schwarzacher, T. (2002) Characterization of mildew resistant wheat-rye substitution lines and identification of an inverted chromosome by fluorescent in situ hybridization. Heredity 88, 349–355. 9. Heslop-Harrison J. S. (2000). Comparative genome organization in plants: from sequence and markers to chromatin and chromosomes. Plant Cell 12, 617–635. 10. Brandes, A., Thompson, H., Dean, C. and Heslop-Harrison, J. S. (1997) Multiple, repetitive DNA sequences in the paracentromeric regions of Arabidopsis thaliana L. Chromosome Res. 5, 238–246. 11. Dechyeva, D., Gindullis, F. and Schmidt, T. (2003) Divergence of satellite DNA and interspersion of dispersed repeats in the genome of the wild beet Beta procumbens. Chromosome Res. 11, 3–21. 12. Lim, K. Y., Kovarik, A., Matyasek, R., Chase, M. W., Knapp, S., McCarthy, E., Clarkson, J. J. and Leitch, A. R. (2006) Comparative genomics and repetitive sequence divergence in the species of diploid Nicotiana section Alatae. Plant J. 48, 907–919.

13. Schwarzacher, T. (1997) Three stages of meiotic homologous chromosome pairing in wheat: cognition, alignment and synapsis. Sexual Plant Reprod. 10, 324–331. 14. Weierich, C., Brero, A., Stein, S., von Hase, J., Cremer, C., Cremer, T. and Solovei, I. (2003) Three-dimensional arrangements of centromeres and telomeres in nuclei of human DNA murine lymphocutes. Chromosome Res. 11, 485–502. 15. Desel, C., Jung, C., Cai, D. G., Kleine, M. and Schmidt, T. (2001) High-resoltion mapping of YACs and the single-copy gene Hs1pro-1 on Beta vulgaris chromosome by multi-colour flurosecence in situ hybridization. Plant Mol. Biol. 45, 113–122. 16. Fransz, P. F., Stam, M., Montijn, B., Ten Hoopen, R., Wiegant, J., Kooter, J. M., Oud, O. and Nanniga, N. (1996) Detection of singlecopy genes and chromosome rearrangements in Petunia hybrida by fluorescence in situ hybridization. Plant J. 9, 767–774. 17. Pedersen, C., Zimny, J., Becker, D., JähneGärtner, A. and Lärz, H. (1997) Localization of introduced genes on the chromosomes of transgenic barley, wheat and triticale by fluorescence in situ hybridization. Theor. Appl. Genet. 94, 749–757. 18. Leggett, J. M., Perret, S. J., Harper, J. and Morris, P. (2000) Chromosomal localization of cotransformed transgenes in the hexaploid cultivated oat Avena sativa L. using fluorescence in situ hybridization. Heredity 84, 46–53. 19. Salvo-Garrido, H., Travella, S., Schwarzacher, T., Harwood, W. A. and Snape, J. W. (2001) An efficient method for the physical mapping of transgenes in barley using in situ hybridization. Genome 44, 104–110. 20. Salvo-Garrido, H., Travella, S., Bilham, L. J., Harwood, W. A. and Snape, J. W. (2004) The distribution of transgene insertion sites in barley determined by physical and genetic mapping. Genetics 167, 1371–1379. 21. Harper G., Osuji J. O., Heslop-Harrison J. S. and Hull R. 1999. Integration of banana streak badnavirus into the Musa genome: molecular and cytogenetic evidence. Virology 255, 207–213. 22. Richert-Pöggeler, K. R., Noreen, F., Schwarzacher, T., Harper, G. and Hohn, T. (2003) Induction of infectious Petunia vein clearing (pararetro) virus from endogenous provirus in petunia. EMBO J. 22, 4836–4845. 23. Staginnus, C., Gregor, F., Mette, M. F., Teo, C. H., Borroto-Fernández, E. G., Laimer da Câmara Machado, M., Matzke, M.

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Schwarzacher and Schwarzacher, T. (2007) Endogenous pararetroviral sequences in tomato (Solanum lycopersicum) and related species. BMC Plant Biol. 7, 24. Gall, J. G. and Pardue, M. L. (1969) Formation and detection of RNA-DNA hybrid molecules in cytological preparations. P. N.A.S. USA 63, 378–383. John, H. A., Birnstiel, M. L. and Jones, K. W. (1969) RNA-DNA hybrids at the cytological level. Nature 223, 582–587. Nederlof, P. M., van der Flier, S., Wiegant, J., Raap, A. K., Tanke, H. J., Ploem, J. S. and van der Ploeg, M. (1990) Multiple fluorescence in situ hybridization. Cytometry 11, 126–131. Müller S., Neusser M. and Wienberg J. 2002. Towards unlimited colors for fluorescence

in-situ hybridization (FISH) Chromosome Res. 10, 223–232. 28. Mukai Y., Ankara Y. and Yamamoto M. 1993. Simultaneous discrimination of the three genomes in hexaploid wheat by multicolour fluorescence in situ hybridization using total genomic and highly repeated DNA probes. Genome 36, 489–494. 29. Heslop-Harrison, J. S., Schwarzacher, T., Anamthawat-Jónsson, K., Leitch, A. R., Shi, M. and Leitch, I. J. (1991) In situ hybridization with automated chromosome denaturation. Technique 3, 109–115. 30. Kohli A., Twyman R. M., Abranches R., Wegel E., Stoger E. and Christou P. (2003) Transgene integration, organization and interaction in plants. Plant Mol. Biol. 52, 247–258.

Chapter 15 Establishing Substantial Equivalence: Transcriptomics María Marcela Baudo, Stephen J. Powers, Rowan A. C. Mitchell, and Peter R. Shewry Abstract Regulatory authorities in Western Europe require transgenic crops to be substantially equivalent to conventionally bred forms if they are to be approved for commercial production. One way to establish substantial equivalence is to compare the transcript profiles of developing grain and other tissues of transgenic and conventionally bred lines, in order to identify any unintended effects of the transformation process. We present detailed protocols for transcriptomic comparisons of developing wheat grain and leaf material, and illustrate their use by reference to our own studies of lines transformed to express additional gluten protein genes controlled by their own endosperm-specific promoters. The results show that the transgenes present in these lines (which included those encoding marker genes) did not have any significant unpredicted effects on the expression of endogenous genes and that the transgenic plants were therefore substantially equivalent to the corresponding parental lines. Key words: Bread wheat, transgenics, transcriptomics, transgene expression, substantial equivalence, gluten proteins.

1. Introduction Feeding an expanding world population is a major challenge for the twenty-first century (1) and there is no doubt that transgenesis could make a major contribution to improving crop yields and quality. However, the acceptability of transgenic crops in Western Europe is low, owing partly to public concerns about the safety of the technology. Regulatory authorities also require detailed studies of transgenic crops to be carried out before they are approved for commercial production, including the demonstration that they are “substantially equivalent” to crops produced by conventional breeding (2). Huw D. Jones and Peter R. Shewry (eds.), Methods in Molecular Biology, Transgenic Wheat, Barley and Oats, vol. 478 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-379-0_15

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Although “substantial equivalence” is difficult to define precisely, it is usually considered that the composition of the material should be within the range for conventionally bred lines grown under the same conditions. We have used a custom cDNA microarray (3–5) to determine the substantial equivalence between the transcriptomes of transgenic and conventionally bred lines of wheat expressing the same genes encoding high-molecular-weight (HMW) subunits of wheat glutenins, under the control of their own endospermspecific promoters. The data were initially analysed to identify genes that showed statistically significant (p < 0.05) differences in expression, and then from this set of genes those with fold changes greater than a critical value (1.5) were identified. The transgenic lines also expressed the bar and uidA marker genes and contained the ampR gene and plasmid backbone sequences (6–8). We, therefore, also compared transgenic lines transformed with whole plasmids or with excised DNA fragments containing only the HMW subunit gene and the bar selectable marker gene (5) (see Note 1). The results demonstrated that the expression of the transgene studied had little statistically significant impact on the global genomic expression in the developing grain, particularly when compared to the greater differences (numbers of significantly differentially expressed genes and fold changes) observed between sibling lines produced by conventional breeding (5). The methods described here were developed for use with a cDNA microarray system, and such arrays are still widely used, particularly for small-scale analyses of small numbers of individually selected genes (often called “boutique arrays”). However, most large-scale gene expression studies now use Affymetrix oligonucleotide-based arrays (e.g., the Wheat GeneChip® Probe Array), which provide a much wider coverage and greater reproducibility and flexibility. Many of the methods described here are equally applicable to this system (e.g., for RNA preparation), but specific aspects of GeneChip® expression analysis are also covered.

2. Materials 2.1. Plant Material

1. Conventional bred lines: L88-31 (9), L88-18 (9) and cv. Cadenza (B1084-0-1) (5). 2. Transgenic bread wheat line in the L88-31 line background: homozygous line selected from B102-1-1 (6, 7). 3. Transgenic bread wheat lines in the cv. Cadenza background: “clean” fragment transgenic line B1355-4-2(18) and “whole plasmid” transgenic line B1118-8-4(6) (5).

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1. Containment glasshouse or controlled environment chamber maintained at 18–20°C/10–14°C day/night cycle; 16-h day/8-h night, 50–70% humidity, 750 mE/s/m2 irradiance. 2. Automatic controlled watering.

2.3. SDS– Polyacrylamide Gel Electrophoresis (SDS–PAGE) (see (10))

1. Separating gel buffer: 1.25 M Tris-borate, 1% (w/v) SDS, pH 6.8 (no adjustment required). 2. Stacking buffer: 1.0 M Tris–HCl, pH 6.8, 10% (w/v) SDS. 3. Ammonium persulfate (prepared fresh), 10% (w/v). 4. Sample buffer: 6.55 ml of stacking buffer, pH 6.8, 3.3% (w/v) SDS, 10% (v/v) of glycerol, and 1.54% (w/v) of DTT (100 mM final concentration). Make up to 100 ml with water. 5. Running buffer: 10Xseparating gel buffer. 6. Acrylamide solution: 40% (w/v) acrylamide and 2% (w/v) NN ¢-methylenebisacrylamide.

2.4. Total RNA Extraction (see Note 2)

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Extraction of total RNA from wheat endosperm (11) 1. Extraction buffer: 2% (w/v) CTAB (hexadecyltrimethylammonium bromide), 2% (w/v) PVP (polyvinylpyrrolidinoline K 30), 100 mM Tris–HCl, pH 8.0, 25 mM EDTA, 2.0 M NaCl, 0.5 g/l spermidine (see Note 3). 2. 2-Mercaptoethanol (see Note 4). 3. Chloroform: isoamyl alcohol (24:1) (see Note 4). 4. 10 M LiCl (lithium chloride). 5. SSTE buffer: 1.0 M NaCl, 0.5% SDS, 10 mM Tris–HCl, pH 8.0, 1 mM EDTA.

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Extraction of total RNA from wheat endosperm (12) 1. Homogenization buffer (see Note 5): 1.4% (w/v) SDS, 0.1 M sodium acetate, pH 8.0, 0.5 M NaCl, 0.05 M EDTA, pH 8.0, 0.1% (w/v) 2-mercaptoethanol. 2. Tris– HCl, pH 8.0, buffered phenol/chloroform (1:1, see Note 6).

2.5. cDNA Microarray Labelling

1. cDNA synthesis: oligo (dT)23 anchor primer (Sigma-Genosis, Haverhill, UK) 0.5 µg/µl/70 µM, Superscript III Reverse transcriptase (200 U/ml) and 5× First Strand Buffer (Invitrogen, Paisley, UK), 50× aa-dNTP mix (Sigma, 10 µl dATP 100 mM, 10 µl dCTP 100 mM, 10 µl dGTP 100 mM, 5 µl dTTP 100 mM, 5 µl amino-allyl -dUTP 100 mM). 2. Purification of aa-dUTP-cDNA: MiniElute columns (Qiagen, Crawley, UK), 75% (v/v) ethanol. 3. Amino-allyl-labelled first strand cDNA: ready to use succinimidyl esters of Alexa dyes (Alexa Fluor dye 555/647, Molecular Probes) (see Note 7), 1 M sodium bicarbonate

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labelling buffer, pH 9.0, MiniElute Columns (Qiagen, Crawley, UK). 2.6. Microarray Hybridization and Washing

1. 2× hybridization mix: 400 ml of 50% (w/v) formamide, 450 ml of 10× SSC, 16 ml of 0.2% (w/v) SDS, 2 mg/ml of poly(dA). 2. Wash chamber (50 ml Falcon tube). 3.

2.7. Real Time RT-PCR

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Solution A (2× SSC, 1% (w/v) SDS up to 50 ml with d-H2O), solution B (1× SSC, 0.2% (w/v) SDS, make up to 50 ml with d-H2O), solution C (0.1× SSC, 0.2% (w/v) SDS, make up to 50 ml with d-H2O). cDNA synthesis 1. SuperScriptTM III RT and RNaseOUT®. 2. 2× RT-Reaction Mix (Invitrogen): 2.5 mM oligo (dT)20, 2.5 ng/ml random hexamers and 10 mM MgCl2 and dNTPs.

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Real time RT-PCR reaction: components 1.

SYBR Green I dye, Platinum Taq DNA Polymerase (60 U/ml).

2.

dNTPs (400 mM dGTP, 400 mM dATP, 400 mM dCTP, 400 mM dUTP).

3.

40 mM Tris–HCl, pH 8.4, 100 mM KCl, 6 mM MgCl2.

4.

UDG (uracyl-DNA glycosylase 40 U/ml).

5.

ROX Reference Dye (glycine conjugate of 5-carboxyX-rhodamine, succinimidyl ester).

3. Methods 3.1. Background to cDNA Microarrays

For the detailed transcriptomic studies we used a wheat cDNA microarray of 19,846 spots containing 9,246 unigene sequences (http://www.cerealsdb.uk.net/index.htm) (3). Duplicated unigene set arrays were spotted onto Codelink-activated slides (Amersham Biosciences Ltd, UK). Arrays were hybridized in reverse dye labelling to fluor dye aa-dUTP-cDNA samples using Alexa Fluor dyes 555 and 647, and hybridizations were performed as single-pair comparisons between the transgenic wheat line (B102-1-1 or B1118-8-4 or B1355-4-2) and its background, non-transgenic line (L88-31 or Cadenza) at two stages of endosperm development (14 and 28 days post-anthesis (dpa)), and with leaf tissue at 8 days post-

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germination (dpg) (5). Each of these comparisons was made using three biological replicates for the line, tissue, and developmental stage selected with two technical replicates for each biological replicate, as provided by a dye swap. cDNA-based microarrays have advantages in that they are economical to use and allow the operator full control over the content and design (customized arrays). In contrast, Affymetrix oliogonucleotide-based arrays are more flexible, as they are a single dye system allowing for simpler experimental designs, are more specific (with improved discrimination between similar isoforms and members of multigene families), and provide more quantitative and readily comparable data. The GeneChip® Genome Array uses a set of 25-mer oligonucleotide “probes” designed to match each transcript sequence. In the case of the wheat chip, there are 11 probes in each set (probe set), with the majority being designed to consensus sequences from the assembly of public domain expressed sequence tags (ESTs). Thus, each probe set is representative of many ESTs rather than a single one as in cDNA arrays. The probes are chosen by automated procedures designed to discriminate between different transcripts, and to satisfy other criteria such as relatively consistent GC content. For each probe designed for a target transcript (perfect matches, PM) there is a matching one with a single base change in the middle of the sequence (mismatches, MM). This gives an estimate of non-specific hybridization, which can be corrected for. However, there is a debate about the real value of the signals from the MM probes, and many widely used analysis approaches do not use them (see Section 3.10). Discrimination between transcripts with similar sequences is most effective with shorter (e.g. 25-mer) probes, since a single base mismatch is sufficient to destabilize the hybridization and the fixed length means the hybridization conditions can be standardized to suit all the probes. Conversely, longer, variable length probes such as those used in cDNA microarray platforms will inevitably hybridize with any transcript that shares sequence similarity along a portion of its length, so the actual signal integrates over several different transcript molecules. 3.2. Plant Material and Growth Conditions

Endosperms and leaves of three transgenic lines of hexaploid bread wheat (Triticum aestivum) were used for transcriptome comparison studies. The transgenic wheat lines B102-1-1 (in the L88-31 background) (6, 7) and B1118-8-4 (in the Cadenza background) (5) were produced by co-bombardment with the p1Ax1 plasmid (13) containing the HMW glutenin subunit 1Ax1 (Glu-1Ax) gene under the control of its own endosperm-specific promoter, and a plasmid carrying

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the selectable bar gene and the marker gene uidA under the control of the maize ubiquitin promoter (14). The transgenic line B1355-4-2 (5), also derived from Cadenza, was obtained by co-transformation using “clean” fragments corresponding to the HMW glutenin subunit 1Ax1 gene and the bar gene coding sequences. A conventionally bred, sister line to L8831, L88-18 (9), was also used for transcriptome comparison. For the two conventionally bred lines (L88-31, L88-18) and transgenic line (B102-1-1), the transcriptomes compared were B102-1-1 vs. L88-31, L88-18 vs. L88-31, and B102-1-1 vs. L88-18. For the comparison of transformation methods (i.e., clean fragments vs. whole plasmids), the comparisons were B1355-4-2 vs. Cadenza, B1118-8-4 vs. Cadenza, and B13554-2 vs. B1118-8-4. Details of the relevant gene composition Nomme of the different bread wheat lines studied are shown in Table 1.

Table 1 Relevant gene composition of the different wheat lines under study. The table is based on data in Lawrence et al. (9), Barro et al. (6), and Rooke et al. (7). HMW high molecular weight Endogenous HMW subunit genes

HMW subunit Marker transgenes genes

Parental line. Sister line derived from same cross as L88-6

1A null, 1Bx17, 1By18, 1D null

None

None

L88-18

Control line. Sister line derived from same cross as L88-6 and L88-31

1Ax1, 1Bx17, 1By18, 1D null

None

None

B102-1-1

Transgenic. L88-31 transformed with 1Ax1 gene as whole plasmid

1A null, 1Bx17, 1By18, 1D null

1Ax1

bar, uidA

Cadenza

Commercial cultivar

1A null, 1Bx14, 1By15, 1Dx5, 1Dy10

None

None

B13554-4-2

Cadenza transformed with 1Ax1 gene as clean fragment

1A null, 1Bx14, 1By15, 1Dx5, 1Dy10

1Ax1

bar

B1118-8-4

Cadenza transformed with 1Ax1 gene as whole plasmid

1A null, 1Bx14, 1By15, 1Dx5, 1Dy10

1Ax1

bar, uidA

Line

Characteristics

L88-31

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1. Plants were grown in pots and arranged in a balanced row and column design, providing three biological replicates per treatment (wheat line by developmental stage). 2. Two plants were grown per pot for the endosperms harvested at 14 and 28 dpa. Only two tillers were kept per plant. Selected pots (three biological replicates in the design) contained a third plant, which was sampled for leaf tissue at 8 dpg. 3. Spikes were checked daily and tagged when anthesis was observed in the central spikelets. 4. Seed endosperms were manually dissected from caryopses under aseptic conditions; samples from each pot comprised a minimum of 24 endosperms taken from the central parts only of two spikes. 5. Samples were taken at the same time of the day to avoid effects of diurnal rhythms. 3.3. SDS–PAGE

3.4. RNA Extraction

The expression of the HMW subunit protein was determined for all wheat lines by SDS–PAGE of total grain protein (Fig. 1) using 10% (w/v) acrylamide gels and a Tris-borate buffer system (10). Extraction of total RNA from wheat endosperms The method used was adapted from Chang et al. (11). 1. Grind 2–3 g of tissue to a fine powder in liquid nitrogen using a pre-cooled, small mortar and pestle (-70°C) (see Note 8).

●

Fig. 1. SDS–PAGE of high-molecular-weight (HMW) glutenin subunits from the bread wheat lines used for transcriptomic analysis. Lane 1 non-transformed background L88-31 line; lane 2 transgenic B102-1-1 line; lane 3 conventionally bred L88-18 line (sibling line of L88-31); lane 4 background non-transformed Cadenza (B1084) line; lane 5 “clean fragment” transgenic (B1355) line; lane 6 “whole plasmid” transgenic (B1118) line. The HMW glutenin subunits are indicated by the bracket. The position of HMW subunit 1Ax1 protein is indicated by the asterisks in the different wheat lines (transgenic lines in lane 2, 5, and 6 and conventionally bred line in lane 3). Data are from Baudo et al. (5).

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2. Add the ground tissue quickly to 15 ml of extraction buffer at room temperature (with 300 µl of 2-mercaptoethanol added) and mix completely by inverting the tube (see Note 9). 3. Extract twice with an equal volume of chloroform to isoamyl alcohol (15 ml final volume), separating phases by centrifugation at 15,000 × g at room temperature (RT) for 10 min. 4. Add 0.25 volume 10 M LiCl to the supernatant and mix. Precipitate the RNA at 4°C overnight and harvest by centrifugation at 15,000 × g for 20 min at 4°C. 5. Suspend the resulting pellet in 500 ml of SSTE. 6. Extract the suspension once with an equal volume of chloroform:isoamyl alcohol. 7. Add two volumes of ethanol to the supernatant and precipitate at −70°C for at least 30 min, or 2 h at −20°C. 8. Spin for 20 min at 15,000 × g to pellet RNA. 9. Wash with 75% (v/v) ethanol. 10. Dry pellet and re-suspend in nuclease-free water. Extraction of total RNA from 8-dpg seedlings The extraction method was adapted from Cheng et al. (12). 1. Grind a known amount of tissue (approximately 1 g of young leaves) in liquid nitrogen to a fine powder using a pre-cooled, small mortar and pestle (–70°C) (see Note 9). ●

2. Transfer the frozen powder quickly into 5–10 ml of homogenization buffer prepared in a second mortar (see Note 10), and continue to grind until extract is homogeneous (see Note 11). 3. Transfer the homogenate into a 50-ml oak-ridge centrifuge tube with a cap and incubate at 65°C for 10 min (homogenate final volume 5–10 ml). 4. Cool the tube on ice and add 0.2 volume of 5 M potassium acetate, pH 5.5 (see Note 12). Mix gently but thoroughly and incubate on ice for 10–15 min. 5. Centrifuge (10,000 × g, 4°C, 15 min) and retain the supernatant in a fresh centrifuge tube. 6. Add to the supernatant an equal volume of phenol: chloroform (1:1, v/v) mixture, tightly cap the tube, and shake vigorously for 10 min. Centrifuge (10,000 × g, 21°C, 10 min). Retain and transfer the upper, aqueous layer to a new tube (see Note 13). 7. Repeat the phenol:chloroform extraction and partition of the aqueous layer as in the previous step.

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8. Add 0.1 volume of 3 M sodium acetate (pH 5.3) and 2.5 volumes of ethanol to the retained aqueous phase. Mix well and incubate overnight at -20°C to give more efficient precipitation of nucleic acids. 9. Centrifuge (10,000 × g, 4°C, 30 min) to pellet the nucleic acids and discard the supernatant. Leave the tube inverted for a few minutes. 10. Dissolve the pellet in a small volume (300–700 µl) of nuclease-free water. Transfer the nucleic acid solution to an Eppendorff tube and add 0.67 volume of 10 M LiCl to precipitate the RNA. Mix well and incubate on ice for 20–30 min (see Note 14). 11. Pellet the precipitated RNA by micro-centrifugation (15,000 × g, room temperature, 20 min) and discard the supernatant. 12. Dissolve the pellet in the smallest possible volume (approximately 200–300 µl) of nuclease-free water, repeat the precipitation with LiCl, and pellet the RNA by centrifugation as before. 13. Dissolve the pellet in 200 µl of nuclease-free water and add 15 µl of 5 M potassium acetate (pH 5.5) and 800 µl of ethanol. Mix well and pellet the RNA by microcentrifugation (15,000 × g, room temperature, 20 min) (see Note 15). 14. Remove the supernatant and wash the pellet by the addition of 1.0 ml of 80% (v/v) ethanol followed by centrifugation (15,000 × g, room temperature, 10 min). 15. Dry the pellet at room temperature by leaving the tubes open on the bench for no more than 10 min. Dissolve in 100–200 µl of nuclease-free water. Divide each sample into aliquots to avoid contamination or degradation during repeated thawing and refreezing. Removal of genomic DNA from total RNA samples After the RNA extractions, the RNA fractions were treated with DNA-free (DNase Treatment & Removal Reagents Kit, Ambion) following the manufacturer’s instructions. The system is designed for the removal of contaminating DNA from RNA samples and for the removal of DNase I enzyme after treatment without the need for heat or phenol extraction. ●

RNA quantification and quality control The concentration, integrity, and quality of the RNA are determined using the Nanodrop ND 1000 spectrophotometer (Labtech Int, UK) and Agilent 2100 Bioanalyser (RNA 6000 Nano Assay, Agilent Technologies, Palo Alto, CA, USA) (see Note 16). ●

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Sample storage RNA samples are stored for short periods (up to 3 months) at -20°C, or for longer periods at -80°C. ●

3.5. cDNA Synthesis

1. Add to 100 µg of DNAse-treated total RNA (up to 20 µl volume), 8 µl of oligo (dT)23-anchored primer and nuclease-free water to provide a final mixture volume of 28 µl. 2. Incubate the priming reaction mixture at 70°C for 10 min and then place it on ice for 5 min. 3. Add to the priming reaction mixture the cDNA synthesis reaction mixture: 10 µl 5× first strand buffer, 10 µl of 0.1 M DTT, 5 µl of 50× aa-dNTP mix, 2 µl of Superscript III reverse transcriptase (200 U/L) and nuclease-free water to give a final volume of 50 µl. 4. Incubate at 42°C for 2–3 h. 5. Purify the aa-dUTP-cDNA product using MinElute Columns following the manufacturer’s instructions (Qiagen). 6. The final eluate can be collected as a 10 µl sample. The cDNA product will be used to prepare the probe for the microarray hybridization and for real time RT-PCR. This last reaction can be performed without a 50× aa-dNTP mixture (see Note 17).

3.6. cDNA Microarray Labelling

1. To couple the cDNA with a reactive fluorescent dye, divide the total cDNA volume (see step 6 – Section 3.5) into two aliquots (5 µl each) for reverse dye labelling, and set up the reactions for the different dyes into separate tubes. 2. Add to each tube: 5 µl of amino-allyl purified cDNA (see step 3 – Section 3.5), 3 µl of 1 M sodium bicarbonate labelling buffer (pH 9.0), 2 µl of Alexa Fluor dye 555 or 647, and 10 ml of nuclease-free water. 3. Mix well using a pipette and incubate in the dark for 1 h at room temperature. 4. Remove the uncoupled dye from labelled aa-dUTP-cDNA using mini Elute columns (Qiagen) (see Note 18).

3.7. cDNA Microarray Hybridization

1. Incorporate 20 µl of mixed labelled cDNA (from step 2 – Section 3.6 for both Alexa dyes) to 25 µl of 2x hybridization mix and 2 ml of poly(dA). 2. Denature the probe at 95°C for 3 min. 3. Apply the labelled probe to cover slip and place the slide with the printed Code Link side face down. 4. Place the slide hybridization chamber in an oven at 42°C and incubate overnight.

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5.

Place the slide in a Falcon (blue cap) tube containing wash solution A (see step 2 – Section 2.6) and invert for 15 min at room temperature (see Note 19).

6.

Transfer the slide to a second Falcon tube also containing wash solution A and invert for 15 min, at room temperature.

7.

Transfer the slide to a third Falcon tube containing wash solution B (see step 3 – Section 2.6) and invert for 8 min, at room temperature.

8.

Transfer the slide to a fourth Falcon tube containing wash solution C (see step 3 – Section 2.6) and invert for 5 min, at room temperature.

9.

Place the slide in a dry Falcon tube and spin immediately at 8,000 × g to dry.

10.

Scan hybridized slides using the Axon Instruments GenePix 400B dual laser scanner.

In order to make an assessment of the differential expression of genes in the transcriptomes for any pair of wheat lines, the microarray slides are subjected to image analysis to determine the intensities of the two fluorescent dyes for any observation (spot). After data normalization, a statistical analysis is applied to fit a model, account for the experimental design used, and test the significance of differential expression. 1. Image analysis and normalization The spots on the scanned slides are visualized using the GenePix software (Gene Pix version 5, Axon Instruments, USA) and all are investigated manually to exclude those where hybridization is poor, or where pixelation is not well defined (weak signal). The data provided from such image analysis, given all pixels from each spot, include the mean intensities of the fluorescent (647 or 555) dyes and their log2 ratio. In particular, these values are used in the analysis of differential expression. The data are imported from GenePix to the GeneSpring package (GenSpring 6.2, Silicon Genetics, USA) for normalization of the log2 ratios of the intensity values. In our study, a plot of the log2 ratio (of differential expression) (M) vs. the log2 product (of intensities) (A) revealed that a locally weighted scatter plot smooth (LOWESS) normalization could be applied to remove undesirable trends in the data. In other words, the log2 ratios should show a constant range across the levels of intensity (log product) for all spots, but the plot revealed that there was some trend with increasing A. The purpose of normalization is therefore to remove systematic variation (e.g., due to experimental procedures) from the log2 ratio data. The LOWESS normalization does this by determining the relationship between M and A on a point-by-point basis and

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accounting for that relationship to adjust the M values accordingly (adjusted M=M-LOWESS-fitted M). As there were two separate experiments, the data from the L88-31, L88-18, and B102-1-1 lines were treated separately from the data for the Cadenza lines. 2. Statistical analysis With reference to methods discussed by Kerr (15), the normalized data can be analysed using the GenStat (GenStat 7th Edition, GenStat Procedure Library Release PL15, Lawes Agricultural Trust, Rothamsted Research, Harpenden, UK) statistical system (see (5) and supplementary material supporting (5) for further details). A linear mixed model is fitted to the log2 ratios to take account of the experimental design (biological and technical replicates) as random effects terms before assessing the (fixed) effect of the genes. For this model term, its parameters (one for each gene) “provided with standard errors” are assessed in terms of the overall residual variability (noise) of the data having accounted for the model (signal). The ratio of each parameter to its standard error gives a t-statistic on the residual degrees of freedom and this allows the statistical significance of the differential expression from 0 (on the log2 scale) to be assessed. In our study, genes with significant differential expression (p < 0.05) were filtered on expression and the number of replicates present, so that only those genes with differential expression greater than 1.5-fold and with two or more replicates were retained for further analysis. Full details of the modelling procedure are included in the appendix at the end of this chapter. 3. Confirming differential expression using real time RT-qPCR Real time RTq-PCR should be applied to selected transcripts to confirm expression data from the microarrays. See Section 3.13. 3.9. Microarray Data Presentation

For a simple substantial equivalence experiment, a scatter plot of gene expression in control against expression in trangenic is appropriate. Examples are shown in Fig. 2 from the two experiments undertaken in (5). The GeneSpring package is used to display the results, plotting the pairs of mean intensities for each gene in each comparison of wheat lines and highlighting the small numbers of (statistically significant and differentially expressed) genes of interest. The results are also summarized numerically in Table 2. The results suggested that transgenesis did not affect the expression of significant numbers of endogenous genes and that the transgenic plants were substantially equivalent to their corresponding non-transgenic control (or parental) lines (5). The results also confirmed that the method of transformation (i.e., with clean fragments or whole plasmids) had little impact on the gene expression patterns. The test of substantial equivalence places emphasis on statistical rigour and, for cDNA arrays, takes

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Fig.2. Scatter plot representation of 14-day post-anthesis endosperm transcriptome for the following pair-wise comparisons: transgenic B102-1-1 line vs. control L88-31 line (left), conventionally bred L88-18 vs. control L88-31 line (middle), and transgenic B102-1-1 line vs. conventionally bred L88-18 line (right). The scatter plot shows the visualization of data obtained after the analysis of 14-dpa endosperm tissue transcriptome for the different pair-wise comparison performed between L88-31, L8818, and B102-1-1 bread wheat lines. Dots represent the normalized relative expression level of each arrayed gene for the transcriptome comparisons described. Dots highlighted in black represent statistically significant, differentially expressed genes (DEG) at an arbitrary cut off >1.5. The inner line on each graph represents no change in expression. The two offset lines in each graph are set at a relative cut-off of two-fold. The white-black palette bar (right site of the figure) shows the different degree of gene expression level, and the data trust scale. The vertical axis of the bar represents the relative expression levels (expressed as fold change): white-light grey tones represent no significant change in expression, greys under expression, and dark grey to black over-expression. The horizontal axis of the bar represents the degree to which the data can be trusted: dark or un-saturated tones represent low trust, and bright or saturated tones represent high trust. Data are from Baudo et al. (5).

account of issues such as dye bias and spatial variation across the chip (see Note 20 for an appraisal of current data analysis of cDNA array experiments). Whereas a simple scatter plot of expression in control vs. expression in transgenic is appropriate for simple substantial equivalence experiments, more complex designs may require other means of display. A powerful method of providing an overview of transcriptomic data, whether from cDNA, oligo array, or other platform, is hierarchical clustering (16). This methodology groups correlated gene expression of samples and/or genes together in a tree structure to visualize different levels of relatedness. Often it is used for a tree of genes in one dimension and a tree of samples in the other with gene expression represented by colour to give a “heat map” display. If the treatment being tested has an effect on expression, then differently treated samples will appear as different branches with all replicates appearing as leaves within these; the genes that differ in expression will also be clustered together in the other dimension. Non-hierarchical forms of clustering co-expressed genes such as k-means, Quality Threshold (QT), and self-organizing maps

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Table 2 Numbers of statistically significant and differentially expressed genes in the pairwise comparisons of various transgenic and control (non-transformed) bread wheat lines under study. The table shows the total number and percentages of statistically significant and differentially expressed genes at an arbitrary 1.5-fold cut-off for the different transcriptome comparisons. Data are from Baudo et al. (5) Lines used for comparison

14 dpa endosperms

28 dpa endosperms 8 dpg leaves

No

%

No

%

No

%

5

0.05

2

0.02

6

0.06

Control untransRelated control formed line without line with endogenous 1Ax1 gene (L88-31) 1Ax1 gene (L88-18)

92

0.99

527

0.59

26

0.27

Line with 1Ax transgene (B102-1-1)

154

1.63

118

1.25

4

0.04

Line transformed Control untransformed line with 1Ax1 clean (Cadenza) fragment (B13554)

6

0.06

9

0.1

1

0.01

Line transformed with Control untransformed line 1Ax1 gene as whole (Cadenza) plasmid (B1118)

97

0.07

12

0.13

2

0.02

Line transformed with Line transformed with 1Ax1 26 gene as whole 1Ax1 gene as clean plasmid (B1118) fragment clean gene (B13554)

0.28

4

0.04

3

0.03

Line with 1Ax1 transgene (B102-1-1)

Control untransformed line without 1Ax1 gene (L88-31)

Related control line with endogenous 1Ax1 gene (L88-18)

of are also frequently used. The average expression of such gene clusters can be an informative method of simplifying transcriptome data. Co-expression implies shared transcriptional control and possible functional relationships. Further inspection of the genes found in such clusters can then be carried out to determine whether any share known functions (such as protein storage, response to stress, or defence) or participate in common pathways. For the majority of genes in crops such as wheat, function can be inferred only from sequence similarity. Clustering, display,

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and annotation tools are available in open-source resources such as Bioconductor (http://www.bioconductor.org/) and in commercial packages like GeneSpring (Agilent Technologies, Inc). 3.10. Background to Wheat GeneChip® Analysis

We present here an overview of the steps that should be followed for the Affymetrix GeneChip® expression analysis. Details of the standard protocols can be found in “Affymetrix GeneChip® Expression Analysis Technical Manual” (see Note 21). GeneChip® probe arrays are manufactured by Affymetrix (see Note 22). Many universities and private companies are now fully equipped with the Affymetrix GeneChip TM® array platform and offer customers various levels of service for array probe processing (GeneChip® array probe purchase, cDNA labelling, array hybridization, scanning, array analysis, etc.). The Affymetrix Wheat GeneChip® array, created within the Affymetrix GeneChip® Consortia Program, contains 61,127 probe sets representing 55,052 transcripts for all 42 chromosomes in the wheat genome. The design of the array was based on public domain data from GenBank® and dbEST (http://www.affymetrix. com/community/research/consortia.affx). The gene chip wheat genome array can be used for gene expression studies in the different wheat species: T. aestivum (UniGene Build ~38, April 24, 2004), T. tauschii, T. monococcum, T. turgidum, T. turgidum ssp. durum. The array includes probes designed to ESTs and full-length sequences from all these species through May 2004. The GeneChip® probe arrays are manufactured using a process that combines photolithography and combinational chemistry. Each 1.7 cm2 chip comprises tens to hundreds of thousands of different oligonucleotide probes, with each probe “spot” (Probe Cell) being 20 mm. Each target transcript is measured by a probe set consisting of 11 PM probes and 11 MM probes which are 25 bases long. These PM and MM probes (Probe Pairs) are placed next to each other. The gene expression level can be calculated on the basis of the intensity differences between the PM and MM for all the probe pairs using Affymetrix software (see Note 22) or using the intensity from the PM probes alone (RMA and gcRMA analysis).

3.11. Wheat GeneChip® Expression Analysis

1. RNA sample preparation The isolation and purification of RNA (total RNA or purified poly (RNA species) can be carried out using established protocols for the specific tissue (protocols similar to those described in Section 3.4). There are also many commercially available kits for RNA isolation. For example, the TRIZOL-Reagent® (Invitrogen, see Note 23) protocol is recommended for total RNA extraction from wheat flag leaves. Modification of standard protocol for homogenization (TRIZOL instructions for RNA isolation, step 1) and RNA precipitation (TRIZOL-Reagent® instructions

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for RNA isolation, step 3) should be included and combined if the extract has high contamination with proteoglycans and polysaccharides. During the homogenization step, an additional centrifugation of the initial homogenate (see Note 24) should be performed. During the RNA precipitation step, total RNA recovered from aqueous phase should be precipitated in 2-Propanol and a high-salt precipitation solution (see Note 25). For high purity (particularly for A260/A230 ratio >1.8), we also recommend that the RNA is cleaned up through an RNeasy column (Qiagen, see Note 26). The RNA clean-up is recommended after genomic DNA contamination is removed from total RNA samples (see Section 3.4). The nucleic acid concentration and quality are checked by using the Nanodrop ND 1000 spectrophotometer (Labtech Int, UK) and Agilent 2100 Bioanalyser (RNA 6000 Nano Assay, Agilent Technologies, Palo Alto, CA, USA), respectively (see Note 16). 2. cDNA synthesis and labelling (see Note 27) Double-stranded cDNA is synthesized from total RNA (or purified poly(A)RNA). A biotin-labelled cRNA is then produced by in vitro transcription (IVT) from the cDNA. Fragmentation of the cRNA before hybridization onto the GeneChip® probe array has been shown to be critical for maximum sensitivity. 3. Target hybridization (see Note 28) A hybridization cocktail is prepared including the fragmented cRNA (target) and probe array controls. The hybridization to the probe array takes place during 16-h incubation. 4. Probe array washing and staining 1. Fluidic station set up: The fluidic station is used to wash and stain the probe array. It is operated using a GeneChip® Operating System (GCOS)/Microarray Suite on a PCcompatible workstation. This step includes setting up and priming the fluidic station (see Notes 29 and 30). 2. Probe array washing and staining (see Note 31): After 16 h of hybridization, the hybridization cocktail is removed from the probe array (see Note 28) and the probe array is filled completely with appropriate volume of recommended wash buffer (see Note 31). 5. Probe array scan (see Note 31) Once scanned, each complete probe array image is stored in a data file identified by the experiment name and saved with a data image file (.dat) extension. The GCOS captures and analyses the array image and experimental data: probe cells are defined and the intensity for each cell is computed (see Note 22). Owing to the higher quality control during manufacture, many of the issues on image analysis for cDNA arrays (Section 3.8), e.g., spatial trends, are considered not to apply to Affymetrix chips. Output

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files containing a signal value for every PM and MM probe are generated (cel files). 3.12. Wheat GeneChip® Data Analysis

As Affymetrix chips are so widely used across many organisms, considerable effort has been invested in developing and testing different methods of analysing the signals to give the best measure of gene expression. The method developed by Affymetrix uses the difference between the PM and MM probes averaged across a “probe set” to give a measure of expression (MAS5). However, there is some doubt as to the usefulness of MM signals and some of the alternative methods seem empirically to outperform MAS5. The Robust Multichip Average (RMA) algorithm takes the PM data from all the samples in an experiment (i.e., all the cel files) and not only normalizes to the median expression value of each chip but also imposes the same variance on the expression data from each chip (17). The gcRMA algorithm is a variant of RMA that takes into account the GC composition of each probe to weight the contribution to the probe set signal (18) and has been shown to perform well for Affymetrix array data compared to alternative methods when assessed by agreement to spike-ins (19) and to real-time RT-PCR measurements (20). The gcRMA algorithm can be applied using the open source Bioconductor package (http://www.bioconductor.org/) or using commercial software, e.g., GeneSpring® 7 (Agilent Technologies, Inc). Once the method of expression has been chosen (i.e., MAS5, RMA, gcRMA, or other), the subsequent analysis is the same, with the exception that unnormalized data such as MAS5 must first be normalized (e.g., by dividing by the median expression value for each chip). It is recommended practice to first filter the probe sets considered, to remove all except those that show absolute expression above a threshold (this can be judged from the signal given by non-wheat controls included on the chip) in at least one sample. These probe sets are further filtered to those that show differential expression above a threshold between any pair of samples; usually a fold change of 1.4 is considered the smallest that can be detected. For a substantial equivalence experiment, the design will normally consist of two or more genotypes with at least three biological replicates of each. To identify a significantly differentially expressed set of genes between genotypes, analysis of variance (ANOVA) is applied to the expression values on a log scale (since they usually have log-normal distribution) for every probe set. Given the very large number of probe sets typically tested at this stage, even after filtering, a multiple-testing correction can be applied. The Benjamini–Hochberg false discovery rate (FDR) correction (21) is an appropriate choice. Applying ANOVA to many probe sets with a typical p-value threshold of 0.05 and Benjamini–Hochberg multiple-testing correction, about 5% of the resulting list of genes that pass can be expected

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to be present by chance alone. However, if few or no genes pass this correction, it can be omitted and the predicted FDR taken into account in interpretation. For example, if 1,000 probe sets are tested at p < 0.05 and 50 pass with no multiple testing correction, this is no more than would be expected by chance alone. The criteria for substantial equivalence in interpreting the lists of significant genes are then the same as described for cDNA array experiments (5). 3.13. Validation of Transcriptomic Data by Real Time RT-PCR

Two general methods for the quantitative detection of genes (amplicon) have become established: gene-specific fluorescent probes (e.g., TaqMan chemistry) or specific double strand DNA binding agent (SYBR green chemistry) (22). We used SYBR green chemistry to validate by real time RT-PCR the expression of selected DEGs identified in the cDNA microarray. Specific primers were designed for the different genes (see Note 32). ●

Preparation of real time RT-PCR reactions 1. PCRs are performed in optical 96-well plates with an ABI® PRISMA 7500 Sequence Detection System (Applied Biosystems, Foster City, CA, USA). 2. Total RNAs (2 µg of DNase treated RNA, from Section 3.4) are reverse transcribed using the Reverse Transcriptase (RT) and buffer (SuperScriptTM III RT, Invitrogen) following manufacturer’s instructions. 3. The PCR reactions are performed with an estimated 100 ng of cDNA, 12.5 µl of 2x Platinum® qPCR Super MixUDG with SYBR green (Invitrogen), 0.5 µl of ROX Reference Dye (Invitrogen), and specific pair of primers (200 ng of each specific primer) in a final volume of 25 µl (see Note 33). 4. The following standard thermal profile is used for all PCR reactions: one cycle at 50°C for 2 min, one cycle at 95°C for 2 min, 40 cycles at 95°C for 15 s and 60°C for 1 min (see Note 34).

●

Real-time PCR data extraction and analysis 1. Raw data extraction. Threshold cycle (Ct) values were collected from each sample (23). In order to compare the Ct values from different cDNA samples, the Ct of all the validated genes were normalized to that of the housekeeping gene actin (see Note 35). 2. Data analysis. The relative expression of the selected genes was calculated using the equations as proposed by Pfaffl (24). These algorithms include the correction for gene amplification efficiency. PCR amplification efficiency for the different target genes and reference gene

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was estimated using the equations as proposed by Ramakers et al. (25). 3.14. Submission of Microarray Dataset to a Public Database: ArrayExpress

Microarray data should be deposited in a public repository database. ArrayExpress (26) is a public database at European Bioinformatics Institute (EBI) for high-throughput functional genomic data. This database consists of two parts: the ArrayExpress Repository (MIAME, the compliant primary archive) and the ArrayExpress DataWarehouse (a database of selected gene expression profiles from the repository which is consistently reannotated). ArrayExpress is one of the three databases recommended by MGED (Microarray Gene Expression Data Society) for deposition of public microarray data. It stores data used in publications in a confidential form while allowing access to authorized users such as journal editors and referees, with specified data being made publicly available upon publication of the paper to which they relate. The microarray data submission includes, in general, four main steps: (1) creating a new account for submission (MX account), (2) protocol submission (array preparation protocols, sample growth and extraction protocols, sample labelling, hybridization, scanning, analysis protocols, etc.), (3) array design submission (name, design, technology, etc.), and (4) experiment submission (experiment design, publications, samples, extracts, etc.). For more information, see webpage: http:// www.ebi.ac.uk/miamexpress/Help.

3.15. Statistical Modelling

The method of Residual Maximum Likelihood (REML) (27), implemented in the GenStat Seventh Edition (2003) statistical system, was used to fit a mixed model (consisting of random and fixed effects) to the data as a complete set, for any particular comparison (for example, B102-1-1 to L88-31 at 14 dpa), with up to six observations per gene. The design of the experiment in terms of the variance structure relating to biological replicates and technical replicates was assessed in terms of the model deviance, changes in which are distributed as X2 on the corresponding change in degrees of freedom when testing between models. Fixed terms in the model are assessed using the Wald test (28), the test statistic for this also being distributed as X2. The modelling procedure therefore takes account of variation due to the design terms (random effects) and fits a fixed term for them (9,246 genes). Having assessed the significance of random and fixed terms, the model used for B102-1-1 to L88-31 at 14 dpa was yijk = BioRepi + (BioRep ´ TechRep)ij + Genek + eijk, where yijk represents the log2 ratio of gene expression of B1021-1 to L88-31 at 14 dpa for biological replicate (BioRep) i, i = 1,…, 3, technical replicate (TechRep) j, j = 1, 2 (i.e., dye swap: j

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= 1 for dye 555/dye 647 ratio, j = 2 for dye 647/dye 555 ratio) and Gene k, k = 1,…, 9,246. The error term is given by eijkl. To fit the model, all terms were set up as factors (indicator columns) in GenStat. Use of the x between BioRep and TechRep model terms implies an interaction. The main effect of biological replicate followed by this interaction indicates that the random part of the model is the technical replicate (dye swap) nested within biological replicate. This form for the random part therefore matches the design used. The Gene term in the model contains parameters for all 9,246 genes. On the log-scale, the ratio of any one parameter for a gene to its standard error forms a t-test on the overall degrees of freedom for the model, for that gene. This allowed the statistical significance of differential expression for any gene (e.g., when comparing B102-1-1 to L88-31 at 14 dpa using the above model) to be assessed. Similar modeling was used for the comparisons of L88-31 with L88-18 and for B1021-1 with L88-18 at the three material/time points. For the three Cadenza lines, data from each set of the three experimental material/time points (seed/14 dpa and 28 dpa; leaf/8 dpg) were modelled by combining data from all three comparisons using a design term to indicate the precise comparison: B1355-4-2(18) vs. control Cadenza, B1118-8-4(6) vs. control Cadenza, or B1118-8-4(6) vs. B1355-4-2(18), and two indicator variables to denote the comparison with B1355-4-2(18) or with B1118-8-4(6) for the fixed effect of each gene. Having assessed the significance of the design terms, the best model was the same for all three material/time points: yijkl = Comparisoni + (Comparison ´ BioRep)ij + (Comparison ´ BioR ep ´ TechRep)ijk + (Genel ´ Cad1) + (Genel ´ (Cad2 ) + eijkl, where yijkl denotes the log2 ratio for comparison i = 1, 2, 3 (B13554-2(18) vs. control Cadenza, B1118-8-4(6) vs. control Cadenza, and B1118-8-4(6) vs. B1355-4-2(18)); biological replicate j, j = 1, 2, 3; technical replicate k, k = 1, 2 (dye swap: dye 555/dye 647, dye 647/dye 555) and gene l, l = 1,…, 9,246 for a comparison with B1355-4-2(18) (Cad1) as “treated line” (numerator of log2 ratios) (Cad1 = 1) or “control line” (denominator of log2 ratios) (Cad1 = –1) or not present (Cad1 = 0), or for comparison with B1118-8-4(6) (Cad2) as “treated line” (Cad2 = 1) or not present (Cad2 = 0), and where eijkl is the error in fitting yijkl. The dot between terms indicates their interaction, so that in the above model, for the random terms, the biological replicates are nested within the comparisons being made, while the technical replicates (dye swaps) are nested within the biological replicates. Hence, again here, the form of the random part of the model found to be best also relates intuitively to the form of the design

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used for the Cadenza study. Also, because of the form of the fixed part of the model used, i.e., with the indicator variables, the test of any gene for a particular comparison benefits from the extra information derived for that gene over all comparisons, all three tests for each gene being made using the same overall residual variation.

4. Notes 1. Data deposition. The gene expression data presented here have been deposited in ArrayExpress database (http:// www.ebi.ac.uk/arrayexpress/Submissions/index.html) with the accession number A-MEXP-177. 2. Unless stated otherwise, all chemical solutions should be nuclease free. 3. Extraction buffer is best stored at room temperature. 0.5 g/l of spermidine should be added to the extraction buffer after autoclaving. 2-Mercaptoethanol should be added to the aliquoted buffer immediately before use. 4. Reagents should be handled carefully and under a flow hood as some are toxic, flammable, or irritants. 5. Fresh solutions from concentrated stocks of the individual constituents should be extracted just before they are to be used. 6. Tris–HCl, pH 8.0, buffered phenol:chloroform (1:1) is prepared by carefully adding 800 ml of 10 mM Tris–HCl, pH 8.0, to 400 g of redistilled ultra-pure phenol crystals and 0.4 g of 8-hydroxiquinolene (antioxidant). The mixture is stirred for 1–2 h and then allowed to stand and separate. The upper aqueous buffer layer is discarded. The lower yellow-coloured phenol:chloroform layer is retained for use and is stable at room temperature for 1–2 months, if kept in the dark. 7. Dissolve one vial of the Alexa-dye in 2 ml of DMSO (dimethyl sulfoxide) 8. Pre-cool the mortar, pestle, and spatulas in liquid nitrogen and keep frozen before use. 9. Warm 15 ml of extraction buffer + 300 µl of 2-mercaptoethanol to 65°C in a water bath. 10. Keep a second mortar and pestle and a buffer for homogenization at room temperature.

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11. Add more buffer in small amounts (1–2 ml) if the homogenate is too viscous. 12. Add potassium acetate (pH 5.5) to precipitate K+-SDS/protein/genomic-DNA/carbohydrate complexes. 13. Be careful not to contaminate the retained aqueous layer with material from the precipitated, interfacial, denatured protein layer. 14. The efficiency of precipitation of RNA by LiCl is dependent on the nucleic acid concentration, and there is a sudden and marked reduction in precipitation efficiency when the RNA concentration falls bellow 100 mg/ml. 15. To exchange K+ for the residual Li+ ions bound to the purified RNA. 16. For well-purified RNA, the A260/A280 ratio should be approximately 2.0 and the A260/A230 ratio should be >1.8. Concentration and purity can be determined using the Nanodrop ND 1000 spectrophotometer and the Agilent 2100 Bioanalyzer (http://www.agilent.com). 17. Alternatively, cDNA for relative quantification by real time RT-PCR can be produced with SuperScriptTM III RT and RNaseOUT kit (Invitrogen), following the manufacturer’s instructions. 18. To elute the aa-dUTP-labelled cDNA, add 10 µl of nucleasefree water to the centre of the Mini Elute column membrane, allow the column to stand for 5 min and centrifuge for 5 min. 19. Place each slide in a 50-ml Falcon tube. Cover all tubes with aluminium foil. 20. In our study, the data were analysed using the GenStat system because GeneSpring was not convenient for the modelling that was required to test for the significance of individual genes in the context of an overall model for all genes rather than on a gene-by-gene basis. In many studies, the normalization procedure is most conveniently done using the GenSpring system. However, since our study was carried out, the GenStat Statistical System has been developed further (in the eighth and ninth editions), allowing a more sophisticated approach to be taken in modelling microarray data. The data after the image analysis can now be more simply imported to GenStat for normalization and full analysis. Furthermore, subsequent and on-going research suggests that a different modelling technique could be applied that accounts specifically for the spatial variability of spots within blocks on a slide (29, 30). Alternatively, hierarchical mixture modelling (31, 32) of the normalized expression values

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could be used in order to deal with the complex variability of the data while still having power to detect the differential expression where there are few observations per gene. Finally, the analysis of multiple laser scans of microarray slides (33) followed by functional regression modelling may allow the effects of the intrinsic noise level of the scanner on the censoring of highly expressed genes to be overcome. 21. Where a large number of genes are tested using the modelling approach discussed here, an investigation of the false discovery rate of significant genes would be useful through the fitting of a mixture distribution to the p-values for genes using theory (34), now incorporated as the FDRMIXTURE procedure in GenStat. Finally, the problem of having a small number of data points for each of a large number of genes to be tested can be alleviated by using a variance shrinkage method (35). This uses a test based on variance estimates that are gene-specific but combining information across the genes. This is more powerful than tests on individual genes but avoids the problem of false discovery rate associated with using an assumption of common underlying variance for all genes (as taken in the modelling described here). 22. Protocols can be downloaded from the web page: http:// www.affymetrix.com/support/technical/manual/netaffx_ MAGE_ML_manual.affx. 23. See http://www.affymetrix.com for current technology references. Up to 1.3 million different oligonucleotide “probes” (where a probe is any standard 25-mer oligo sequence synthesized on the array used to detect a complementary target in solution) are synthesized on each array. Each probe is located in a specific area called a “probe cell”. Each probe cell contains between hundreds of thousands and millions of copies of a given nucleotide. The “probe pair” refers to the fundamental detection unit on an Affymetrix probe set consisting of a perfect match (PM) and corresponding mistmach (MM) oligo. On the array, a set of probe pairs (the “probe set”) represents the selected expression sequences. 24. TRIZOL-Reagent, Invirtogen catalog number 15596-026. 25. Following homogenization (TRIZOL-Reagent® instructions for RNA isolation, step 1), remove insoluble material from the homogenate by centrifugation at 12,000 x g for 10 min. 26. Total RNA in the aqueous phase should be precipitated in 0.25 ml of 2-propanol (Sigma) followed by adding 0.25 ml of a high-salt precipitation solution (0.8 M sodium citrate and 1.2 M NaCL) per 1 ml TRIZOL-Reagent®. 27. RNs easy column (Qiagen) used to clean up and concentrated eluted total RNA, following manufacturer’s instructions.

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28. Protocols described in GeneChip® expression Analysis Technical Manual, Section 2, Chapter 1. 29. Protocols described in GeneChip® Expression Analysis Technical Manual, Section 2, Chapter 2. 30. Protocols described in GeneChip® Expression Analysis Technical Manual, Section 2, Chapter 3, Part 7. 31. Refer to GeneChip® Fluidics Station User’s Guide for instructions. 32. Protocols described in GeneChip® Expression Analysis Technical Manual, Section 2, Chapter 3.9 for “Probe wash and Stain” and Chapter 3.15 for Probe Array Scan). Review the scanner user’s manual for safety precautions and more information. 33. Specific pairs of primers for SYBR-green detection and quantification of selected differentially expressed genes (DEG) for microarray validation (EST clone sequences searched in http://www.cerealsdb.uk.net/index.htm) were designed using Primer Express® software (ABITM PRISMA) following the TaqMan® Probe and Primer Design guides. 34. A master mix with sufficient cDNA and reaction components is prepared prior to dispensing into individual wells to ensure that each reaction (i.e., three technical replicates for each of the three biological replicates per sample tested) contains an equal amount of cDNA, and that pipetting and other errors are reduced. 35. A dissociation curve analysis is performed for each pair of specific primers to detect if non-specific amplification has occurred. To generate a baseline-subtracted plot of the logarithmic increase in fluorescence signal (DRn) vs. cycle number, the baseline data were collected for most of the amplifications between cycle 3 and 15. 36. The Actin gene (clone ID: H01_p335_plate_6; http://www. cerealsdb.uk.net/index.htm) used as internal control gene does not show differential expression between the lines, tissues and developmental stages under study. 37. Availability: http://www.ebi.ac.uk/arrayexpress.

Acknowledgements Rothamsted Research receives grant-aided support from the Biotechnology and Biological Sciences Research Council of the UK. The transcriptomic studies were supported by a grant under the BBSRC, Gene Flow Initiative (ref. GM 14152). The authors would

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like to thank Mr. Adrian Price at Rothamsted Research for discussions of methods for microarray data analysis. We also acknowledge our colleagues Prof. Michael Holdsworth (University of Nottingham), Prof. Keith Edwards (University of Bristol), Ms. Rebecca Lyons, and Dr. Gabriela M. Pastori (Rothamsted Research).

References 1. Evans, L. T.(1993)Crop Evolution, Adaptation and Yield.Cambridge University Press,Cambridge. 2. FAO/WHO (1996) Biotechnology and Food Safety, Report of a Joint FAO/WHO Consultation. 3. Wilson, I. D., Barker, G. L. A., Beswick, R. W., Shepherd, S. K., Lu, C., Coghill, J. A., Edwards, D., Owen, P., Lyons, R., Parker, J. S., Lenton, J. R., Holdsworth, M. J., Shewry, P. R. and Edwards, K. J. (2004) A transcriptomics resource for wheat functional genomics. Plant Biotechnol. J. 2, 495–506. 4. Wilson, I. D., Barker, G. L. A., Lu, C., Coghill, J. A., Beswick, R. W., Lenton, J. R. and Edwards, K. J. (2005) Alteration of the embryo transcriptome of hexaploid winter wheat (Triticum aestivum cv. Mercia) during maturation and germination. Funct. Integ. Genomics 5, 144–154. 5. Baudo, M. M., Lyons, R., Powers, S., Pastori, G. M., Edwards, K. J., Holdsworth, M. J. and Shewry, P. R. (2006) Transgenesis has less impact on the transcriptome of wheat grain than conventional breeding. Plant Biotechnol. J. 4, 369–380. 6. Barro, F., Rooke, L., Békés, F., Gras, P., Tatham, A. S., Fido, R. J., Lazzeri, P., Shewry, P. R. and Barcelo, P. (1997) Transformation of wheat with HMW subunit genes results in improved functional properties. Nat. Biotechnol. 15, 1295–1299. 7. Rooke, L., Steele, S. H., Barcelo, P., Shewry, P. R. and Lazzeri, P. (2003) Transgene inheritance, segregation and expression in bread wheat. Euphytica 129, 301–309. 8. Shewry, P. R., Halford, N. G., Tatham, A. S., Popineau, Y., Lafiandra, D. and Belton, P. S. (2003) The high molecular weight subunits of wheat glutenin and their role in determining wheat processing properties. Adv. Food Nutr. Res. 45, 221–302. 9. Lawrence, G. J., Macritchie, F. and Wrigley, C. W. (1998) Dough and baking quality of

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wheat lines in glutenin subunits controlled by Glu-A1, Glu-B1 and Glu-D1 loci. J. Cereal Sci. 7, 109–112. Shewry, P. R., Tatham, A. S. and Fido, R. J. (1995) Plant Gene Transfer and Expression Protocols: Separation of Plant Proteins by Electrophoresis, Vol. 49. Humana, Totowa. Chang, S., Puryear, J. and Cairney, J. A. (1993) Simple and efficient method for isolating RNA from pine trees. Plant Mol. Biol. Rep. 11, 113–116. Cheng, G. P., Wilson, I. D., Kim, S. H. and Grierson, D. (2001) Inhibiting expression of a tomato ripening-associated membrane protein increases organic acids and reduces sugar levels of fruit. Planta 212, 799–807. Halford, N. G., Field, J. M., Blair, H., Urwin, P., Moore, K., Robert, L., Thompson, R., Flavell, R. B., Tatham, A. S. and Shewry, P. R. (1992) Analysis of HMW glutenin subunits encoded by chromosome 1A of bread wheat (Triticum aestivum L.) indicates quantitative effects on grain quality. Theor. Appl. Genet. 83, 373–378. Christensen, A. H. and Quail, P. H. (1996) Ubiquitin promotor-based vectors for highlevel expression of selectable and/or screenable marker genes in monocotyledonous plants. Transgen. Res. 5, 213–218. Kerr, M. K. (2003) Linear models for microarray data analysis: hidden similarities and differences. J. Comput. Biol. 10, 891–901. Eisen, M. B., Spellman, P. T., Brown, P. O. and Botstein, D. (1998) Cluster analysis and display of genome-wide expression patterns. P.N.A.S. USA 95, 14863–14868. Irizarry, R. A., Hobbs, B., Collin, F., BeazerBarclay, Y. D., Antonellis, K. J., Scherf, U. and Speed, T. P. (2003) Exploration, normalization, and summaries of high density oligonucleotide array probe level data. Biostatistics 4, 249–264. Wu, Z. J., Irizarry, R. A., Gentleman, R., Martinez-Murillo, F. and Spencer, F. (2004)

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27. Patterson, H. D., and Thompson, R. (1971). Recovery of inter-block information when block sizes are unequal. Biometrika 58, 545–554. 28. Welham, S. J., and Thompson, R. (1997). Likelihood ratio tests for fixed model terms using residual maximum likelihood. J. Royal Stat. Soc. Ser B 59, 701–714. 29. Burgueño, J., Crossa, J., Grimanelli, D., Leblanc, O. and Autran, D. (2005) Spatial analysis of cDNA microarray experiments. Crop Sci. 45, 748–757. 30. Baird, D., Johnston, P. and Wilson, T. (2004) Normalisation of microarray data using a spatial mixed model analysis which includes splines. Bioinformatics 20, 3196– 3205. 31. Smyth, G. K. (2004) Linear models and empirical Bayes methods for assessing differential expression in microarray experiments. Stat. Appl. Genet. Mol. Biol. 3. 32. Newton, M. A., Noueiry, A., Sarkar, D. and Ahlquist, P. (2004) Detecting differential gene expression with a semiparametric hierarchical mixture method. Biostatistics 5, 155–176. 33. Khondoker, M. R., Glasbey, C. A. and Worton, B. J. (2006) Statistical estimation of gene expression using multiple laser scans of microarrays. Bioinformatics 22, 215–219. 34. Allison, D. B., Gadbury, G. L., Heo, M., Fernández, J. R., Lee, C. -K., Prolla, T. A. and Weindruch, R. (2002) A mixture model approach for the analysis of microarray gene data. Comput. Stat. Data Anal. 39, 1–20. 35. Cui, X., Gene-Hwang, J. T., Qui, J., Blades, N. J. and Churchill, G. A. (2003) Improved statistical tests for differential gene expression by shrinking variance components estimates. Biostatistics 6, 59–75.

Chapter 16 Establishing Substantial Equivalence: Proteomics Alison Lovegrove, Louise Salt, and Peter R. Shewry Abstract Wheat is a major crop in world agriculture and is consumed after processing into a range of food products. It is therefore of great importance to determine the consequences (intended and unintended) of transgenesis in wheat and whether genetically modified lines are substantially equivalent to those produced by conventional plant breeding. Proteomic analysis is one of several approaches which can be used to address these questions. Two-dimensional PAGE (2D PAGE) remains the most widely available method for proteomic analysis, but is notoriously difficult to reproduce between laboratories. We therefore describe methods which have been developed as standard operating procedures in our laboratory to ensure the reproducibility of proteomic analyses of wheat using 2D PAGE analysis of grain proteins. Key words: Substantial equivalence, wheat, 2-D PAGE, genetically modified, proteomics, mass spectrometry.

1. Introduction Almost all of the wheat consumed by humankind has been processed into food products, including baked goods (bread, cakes, pastries and biscuits), noodles and pasta, and ingredients used to confer specific functional properties. This pervasiveness of wheatderived products in the food chain means that it is usually encountered as flour or in processed forms rather than as grain. Also, the strong cultural and religious significance of many wheat-derived products means that the introduction of transgenic forms is more emotive for wheat than for other transgenic crops. The establishment of routine procedures for the identification of transgenic wheat and to determine its substantial equivalence (1) to conventionally bred wheat is therefore of concern for consumers and regulatory authorities (2). Several approaches can be Huw D. Jones and Peter R. Shewry (eds.), Methods in Molecular Biology, Transgenic Wheat, Barley and Oats, vol. 478 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-379-0_16

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used for this (3) including proteomics, which is well established for wheat grain (4–7) and has been applied to determine substantial equivalence in other transgenic comparisons (8–10). It is not possible to determine the complete proteome of any given cell, as the protein composition is dynamic and varies depending on the internal and external environment at the time of sampling. Also, the vast differences in the amounts of different protein components mean that it is often not possible to identify and quantify minor components. Nevertheless substantial progress has been made in the proteomic analysis of wheat and other cereals. The traditional method of separating proteins in proteomic studies is 2D PAGE, and this is still the method used in most studies of wheat (4–7). However, many laboratories are now using 2DLC-MS either instead of or together with 2D-PAGE. These two techniques are complementary, and they often identify different proteins. However, the cost of 2DLC-MS may still be prohibitive for many laboratories, and thus 2D PAGE is still the most widely used and widely available technique. The development of immobilized pH strips has made 2D-PAGE much less technically demanding and greatly improved reproducibility. However, 2D-PAGE still has limitations, with membrane proteins, highly charged (basic and acidic) proteins and very large proteins being difficult to resolve. Furthermore, low abundance proteins are also difficult to identify in the presence of highly abundant components. This is a particular problem with wheat grain as the major storage proteins can mask many other proteins if a total protein extract is used. However, these problems can be largely overcome by cell fractionation or sequential extraction techniques, or by the judicious use of restricted pH gradients. The methods described are based on those used in a number of laboratories including our own.

2. Materials All chemicals must be of the highest quality, AristaR if possible. Where the supplier is not quoted it was Sigma, Poole, Dorset, UK. 2.1. Extraction Buffers

1. For white flour: 5 mM Trizma base, 1 mM calcium chloride (CaCl2), no pH adjustment required. 2. Total protein extraction buffer: 2 M thiourea, 7 M urea, 1% (w/v) DTT, 2% (w/v) CHAPS and 0.5% IPG-buffer 3–10 (GE Healthcare, UK). 3. Osborne fractionation buffers: water-saturated butan-1-ol: 50 ml butan-1-ol plus 5 ml water and shake in bottle, allow to

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settle, use top layer; 0.5 M NaCl; 70% (v/v) ethanol; 50% (v/ v) propan-1-ol, 2% (v/v) 2-mercaptoethanol, 1% (v/v) glacial acetic acid; 5% (v/v) glacial acetic acid. 4. Acid:acetone precipitation buffers Solution I: 10% (w/v) TCA made up in acetone containing 0.07% (w/v) DTT Prepare 10% TCA 1 day before extraction and store at −20°C for at least 12 h prior to use. Add DTT fresh on day of use Solution II: Acetone containing 0.07% (w/v) DTT Store acetone at −20°C, 1 day before extraction, for at least 12 h prior to use. Add DTT fresh on day of use Discard solutions after use and prepare fresh solutions for subsequent extractions. 2.2. Isoelectric Focussing (IEF)

1. Re-hydration buffer: 7 M urea, 2 M thiourea, 2% (w/v) CHAPS, 40 mM dithiothreitol (DTT), 0.5% IPG ampholytes (of appropriate pH), a few grains of bromophenol blue. 2. DryStrip™ cover oil (GE Healthcare, UK). 3. Immobiline DryStrips™ (GE Healthcare, UK). 4. Paper electrode wicks (GE Healthcare, UK). 5. Equilibration Buffer: 50 mM Tris–HCl pH 8.8, 6 M urea, 2% (w/v) SDS, 30% (v/v) glycerol, a few grains of bromophenol blue. Add 1% (w/v) DTT, or 4% (w/v) iodoacetamide to fresh equilibration buffer (reduction and carbamidomethylation).

2.3. SDS– Polyacrylamide Gel Electrophoresis (SDS–PAGE)

1. Tris–HCl (1.5 M), pH 8.8. 2. SDS (10% (w/v)). 3. Ammonium persulphate (APS, 10% (w/v)) made up fresh just before use. 4. N, N, N′, N′-tetramethylethylenediamine (TEMED). 5. Water-saturated butan-1-ol. 6. ‘Acrylamide’ solution; 30% Duracryl, 0.65% Bis (Proteomic Solutions, France). 7. Bottom tank buffer: 25 mM Trizma base; 0.015 M glacial acetic acid. 8. Top tank buffer: 0.2 M Trizma base; 0.2 M Tricine; 0.4% (w/ v) SDS. 9. Displacement solution: 0.375 M Tris–HCl, pH 8.8, 50% (v/v) glycerol, a few grains of bromophenol blue.

2.4. Gel Staining

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Coomassie Colloidal Brilliant Blue G-250 1. Fix 50% (v/v) methanol, 10% (w/v) TCA. 2. Destain: 25% (v/v) methanol, 7% (v/v) acetic acid. 3. Rinse: 25% (v/v) methanol.

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4. Staining solution: 17% (w/v) ammonium sulphate, 34% (v/v) methanol, 0.5% (v/v) acetic acid, 0.1% (w/v) Coomassie Blue G250. Usually bought pre-made (e.g. from Sigma; stock is diluted to 1 l with 800 ml water. Forty millilitre of diluted stain is then added to 10 ml of methanol (see Note 1 regarding fixation of cereal proteins). ●

Silver stain 1. Fix: 40% (v/v) ethanol, 10% (v/v) acetic acid (substitute with 10% (w/v) TCA if fixing cereal proteins). 2. Sensitizing solution: 30% (v/v) ethanol, 0.2% (w/v) sodium thiosulphate, 0.5 M sodium acetate. 3. Silver stain: 0.25% (w/v) silver nitrate. 4. Developer: 0.24 M sodium carbonate, 200 µL of 37% formaldehyde in 1 l (0.0074% final concentration). 5. Stop: 0.04 M EDTA.

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Sypro Ruby™ protein stain (BioRad, UK) 1. Fix: 10% (v/v) methanol, 10% (v/v) TCA. 2. Sypro Ruby™ protein stain. 3. Destain: 10% (v/v) methanol, 6% (w/v) TCA.

2.5. Destaining of Gel Pieces Prior to Enzyme Digest and Mass Spectrometry

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Coomassie and Sypro-stained gel pieces: 1. Water 2. Acetonitrile 3. NH4HCO3 (0.1 M)

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Silver-stained gel pieces: 1. Sodium thiosulphate (20% (w/v)); potassium ferricyanide (1% (w/v)) mixed 1:1:1 with water.

2.6. Reduction and Alkylation (Carbamidomethylation)

1. DTT (10 mM)

2.7. Enzyme Digests

1. Tryptic digest buffer: 25 mM NH4HCO3, 5 mM CaCl2

2. Iodoacetamide (55 mM) 3. NH4HCO3 (0.1 M)

2. Trypsin concentration 0.125 µg/µl (Promega UK) (see Note 2) 2.8. Desalting and Concentration of Peptides

C-18 zip tips (Millipore) ●

MALDI-MS: 1. Wetting solution: 1:1 acetonitrile: water 2. Sample solution: 0.1% (v/v) TFA in water 3. Equilibration solution: 0.1% (v/v) TFA in water 4. Wash solution 0.1% (v/v) TFA in water

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5. Elution buffer 0.1% (v/v) TFA in 1:1 acetonitrile:water ●

ESI-MS: 1. Wetting solution: 1% (v/v) formic acid 2. Sample solution: 1% (v/v) formic acid in 4% (v/v) methanol 3. Washing solution: 1% (v/v) formic acid 4. Elution solution: 1% (v/v) formic acid in 70% (v/v) methanol

2.9. MALDI Target Spotting

1. Two milligram per millilitre α-cyano-4-hydroxycinnamic acid in 49.5% (v/v) acetonitrile, 49.5% (v/v) ethanol, 1% (v/v) 0.1% trifluoroacetic acid (TFA) 2. Tryptic digest of, for example, alcohol dehydrogenase (ADH)

3. Methods It is notoriously difficult to ensure the reproducibility of 2-D PAGE within and between laboratories. In our laboratory strict adherence to standard operating procedures (SOPs) is enforced to ensure that different operators obtain reproducible separations. Examples of 2D gel patterns from wheat white flour samples are shown in Fig. 1. The Tris–CaCl2 extraction method given does not extract ‘total proteins’ but does extract a representative sample of the proteins present in wheat flour except for the gluten proteins which are present in high amounts and which would obscure other less abundant proteins if extracted in total. Similar problems may be experienced with other tissues containing highly abundant proteins, notably Rubisco present in green tissues. We have provided three methods. The first of these which features extraction with Tris–CaCl2 has been used in our laboratory in a large-scale proteomics study of the substantial equivalence GM and non-GM wheat. The second and third extraction methods described have been used widely in the literature and can be applied to a wide range of tissues. 3.1. Extraction of Proteins

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White flour with Tris–CaCl2 buffer (based on (11))

1. To 2 g of white flour add 5 ml of chilled (4°C) extraction buffer. Stir for 30 m at 4°C. 2. Centrifuge at 10,000 × g for 30 m at 4°C to remove insoluble material. 3. Retain the supernatant and flash freeze aliquots in liquid nitrogen. Store at −80°C until use.

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L88-6 RRes

L88-6 LARS

B73-6-1 RRes

B73-6-1 LARS

Fig 1. Examples of 2D gel patterns from wheat white flour samples extracted with Tris–CaCl2 extraction buffer and run under the conditions described and stained with Sypro Ruby™. L88-6 is the parental non-GM control wheat line and B736-1 a transgenic wheat line derived from L88-6 expressing a high proportion of high molecular weight glutenin subunit 1Dx5 under the control of its own promoter. RRes refers to the Rothamsted Research site in Hertfordshire and LARS to Long Ashton Research Station, North Somerset; the two sites where the lines were grown in a field trial over 4 years.

4. Estimate protein concentration using the Bradford assay, with BSA as a standard. ●

Extraction of white flour, or green tissue following an acid/ acetone precipitation (based on (4))

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Flour: 1. In a 50-ml Falcon tube, suspend 5 g flour in 20 ml of solution I, vortex at maximum speed for 2 m and incubate at −20°C for 1 h.

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2. Centrifuge at 15,000 × g for 10 m at 4°C. 3. Discard supernatant and re-suspend the pellet by vortexing in 5 ml of solution II. 4. Centrifuge at 15,000 × g for 10 m at 4°C. 5. Repeat steps 3 and 4 twice more. 6. Discard supernatant, dry pellet under a stream of nitrogen in fume hood (ensuring no material is lost). 7. Re-suspend 40 mg of dried material in 1 ml of re-hydration buffer of appropriate pH. Vortex for 5 m and centrifuge at 13,000 × g at room temperature for 5 m. 8. Retain supernatant and aliquot. Flash freeze aliquots in liquid N2. 9. Store at −80°C. 10. Determine protein concentration of aliquots before use. ●

Green tissue:

1. Grind flash-frozen leaves with a pre-chilled mortar and pestle in liquid N2. 2. Suspend 2 g powdered leaf material in 20 ml of solution I, vortex at maximum speed for 2 m and precipitate at −20°C for 1 h. 3. Centrifuge at 15,000 × g for 10 m at 4°C and discard supernatant. 4. Re-suspend the pellet in 5 ml of solution II, by vortexing and centrifuge at 15,000 × g for 10 m at 4°C. 5. Repeat step 4 twice and finally dry the pellet under a stream of N2 in fume hood, ensuring no material is lost. 6. Re-suspend 20 mg of the dried pellet in 1 ml of rehydration buffer of appropriate pH. 7. Vortex for 5 m and centrifuge at 13,000 × g at room temperature for 5 m. 8. Retain the supernatant, flash freeze aliquots in liquid N2 and store at −80°C until use. 9. Determine protein concentration of aliquots before use. If preparations are contaminated with chlorophyll a PEG/ MgCl2precipitation may be tried (see Note 3). ● Osborne fractionation (based on (12)) 1. Stir 100 mg flour with 1 ml of water-saturated butan-1-ol at room temperature for 1 h. 2. Centrifuge at 5,000 × g for 10 m, at room temperature and discard supernatant. This is a ‘defatting’ step and is OPTIONAL.

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3. Extraction of ‘albumins/globulins’: stir the pellet (from 2) with 1 ml of 0.5 M NaCl for 1 h at room temperature. Centrifuge as described earlier, retain supernatant and repeat extraction. 4. Combine supernatants and dialyse extensively against 5% (v/v) acetic acid (at least four changes of 5 l) for 48 h at 4°C. 5. Extraction of ‘gliadins’: stir the pellet (from 3) with 1 ml of 70% (v/v) ethanol for 1 h at room temperature. Centrifuge, as described earlier. Retain supernatant and repeat extraction. 6. Combine supernatants and dialyse extensively against 5% (v/v) acetic acid (at least four changes of 5 l) for 48 h at 4°C. 7. Extraction of ‘glutenins’: stir the pellet (from 5) with 1 ml of 50% (v/v) propan-1-ol/2% (v/v) 2-mercaptoethanol/1% (v/v) acetic acid for 1 h at room temperature (N.B. DTT may be directly substituted for 2-mercaptoethanol if desired). Centrifuge, as described earlier. Retain supernatant and repeat extraction. Combine supernatants and dialyse extensively against 5% (v/v) acetic acid (at least four changes of 5 l) for 48 h at 4°C. 8. Lyophilize all solutions after dialysis. 9. The final pellet (from 7) may be extracted in 1 ml ‘total protein extraction buffer’ (IEF re-hydration buffer): mix and leave to extract for 1 h at room temperature. Centrifuge, as described earlier and retain supernatant, aliquot and flash freeze in liquid nitrogen and store at −80°C. 3.2. Isoelectric Focussing (IEF)

Immobilized pH Gradient (IPG) strips are available in a range of lengths, e.g. 7, 13, 18 and 24 cm and with a range of pH values. IPG strips are usually re-hydrated overnight (16 h) at 20°C (temperatures lower than 20°C will result in urea crystallization) in the appropriate amount of re-hydration buffer. This may be achieved either in a re-swelling tray or directly in the strip holder. Whichever method is chosen it is essential that all air bubbles are removed from between the strip and the base of the well. Once this is done, the strip should be covered with non-conducting oil. If re-hydration is conducted in a re-swelling tray it must be transferred to the strip holder for running. IEF parameters depend upon the length of the strip, but it is always carried out at 20°C. If the temperature exceeds 20°C protein modification may result, from for example, carbamylation by ammonium cyanate formed from urea. Once isoelectric focussing is complete the excess oil is drained and strips are stored at −80°C. The instructions assume the use of a GE Healthcare IPGphor.

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Strip re-hydration. 1. For 24-cm IPG strips: thaw 4 ml re-hydration buffer and add DTT and IPG ampholytes to the following concentrations: DTT 40 mM; ampholytes 0.5% (v/v). 2. For IEF between pH 3–10 add the appropriate amount of sample to the re-hydration buffer so that 400 µg protein is loaded per strip. Ideally between 50 and 100 µl of sample per strip should be used; and the total amount of buffer and protein should be 450 µl for a 24-cm strip. Ideally a ratio of not less than 1:3 of sample:buffer should be used. 3. Apply 450 µl of buffer and protein mixture to the wells of a 24 cm, 12-well re-swelling tray by pipetting down one side whether re-hydration is to be carried out in the IPG manifold or directly in the strip holders (see Note 4). 4. For focussing between pH 6–11 the total protein and buffer used is the same as for pH 3–10 except that ampholytes of pH 6–11 are used instead of pH 3–10, and the re-hydration buffer is dispensed down one side of a well, and the appropriate amount of sample is applied separately at the anode (+). 5. Place Immobiline DryStrips™, gel-side facing down, into the wells in the re-swelling tray and ensure that there are no air bubbles trapped under the strip. 6. Cover with DryStrip™ cover oil, seal with the lid and leave to re-hydrate for 16–18 h (overnight) at 20°C. 7. Remove strips (using forceps) and drain off excess oil. 8. Place strips, with gel-side facing up, into the 24 cm IPGPhor manifold (up to 12 strips). 9. Align strips into the middle of each well and cover with DryStrip™ cover oil. Fill all wells with oil even if they are not being used.

10. Wet electrode wicks and blot to remove excess water. 11. Place wicks at both ends of the strips, slightly touching the gel. 12. Secure the detachable electrodes to the 24 cm IPGPhor manifold. Place electrodes on the parts of the wicks that are contacting the gel. 13. Lock into position and close IPGPhor lid. 14. The conditions used for IEF must be determined empirically for different extracts, but the following parameters are appropriate for extracts made with flour/Tris–CaCl2: 500 V 1 h; 1,000 V 1 h; 8,000 V 8.20 h. Total Vh 65,500. 15. Following IEF, remove the strips, drain off excess oil and place the strips in 10-ml plastic pipettes (with tips removed, to allow strips to fit inside). Seal with parafilm and store at −80°C.

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3.3. SDS–PAGE (Second Dimension)

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Using a GE Healthcare Ettan Dalt II apparatus 1. Soak plates in detergent overnight, rinse with de-ionized water, followed by ethanol and RO water and air dry. 2. Assemble gel plates (cassettes) according to manufacturer’s instructions (see Note 5 regarding silicone sealant use. Ensure a complete seal around gel plates if non-self-sealing plates are used). 3. For casting multiple gels, pack the gel cassettes into the casting box with a re-usable plastic spacer between each gel cassette. Ensure that the cassettes are packed tightly together and a good seal is formed between the front panel and the casting box by greasing the rubber gasket with silicone grease. Tighten screws on face panel and level the casting box. 4. Attach the casting tube and funnel and hold upright using a clamp and stand. 5. If using displacement solution; after the funnel is in place, add 100 ml of the displacement solution to the balance chamber. If no displacement solution is used polymerized acrylamide may be removed from the tubing at the end of casting with copious amounts of hot water. 6. For twelve 10% gels, use 1 l of gel solution: 250 ml 1.5 M Tris–HCl, pH 8.8, 333 ml 30% (v/v) Duracryl 0.65% (w/v) bis, 407 ml water. Stir, place in a 2 l Buchner flask and degas for 15 m. 7. Add 10 ml of 10% (w/v) SDS and stir briefly. 8. Immediately before use add 2.5 ml 10% APS and 0.5 ml TEMED and stir.

9. Pour the acrylamide solution into the funnel, continually topping up (It is important not to allow the funnel to empty completely as air bubbles will form in the gel). Continue filling funnel until 1–2 cm from top of first front plate (~1 l). 10. Remove the funnel and allow the displacement solution to flow down into the v-shaped trough at the base of the caster. 11. Overlay the top of each of the gels with 1 ml of water-saturated butan-1-ol and leave to polymerize for 3–4 h. 12. Once polymerized, dismantle the casting box, remove the cassettes and rinse with de-ionized water to remove watersaturated butanol and any small pieces of polymerized Duracryl attached to plates. 13. Prepare a 1 in 4 dilution (in RO water) of 1.5 M Tris–HCl pH 8.8 for use as gel storage solution. 14. Place gels in a large plastic box and add enough gel storage solution to completely cover them.

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15. Allow gels to ‘mature’ for 10–14 days at 2–4°C. Gels can be used after this time, but must not be used beyond 1 month of the preparation date. ●

Gel electrophoresis

1. Allow 10 ml of pre-made frozen equilibration buffer for each IPG strip to thaw overnight in a refrigerator or cold room at 4°C. 2. Prepare 7 l of lower tank buffer: 25 mM Trizma base, 15 mM glacial acetic acid. 3. Place buffer in Ettan Dalt II tank, switch on pump and cool to 15°C. 4. Prepare top tank buffer: 3 l of 200 mM Tris base, 0.4% (w/v) SDS, 200 mM Tricine. 5. Rinse IPG strip with water to remove excess oil and equilibrate in 10 ml of equilibration buffer containing 1% (w/v) DTT for 15 m. 6. Remove equilibration buffer and add a further 10 ml of equilibration buffer containing 4% (w/v) iodoacetamide, and incubate for 15 m. Cover tubes with aluminium foil as iodoacetamide is light sensitive. 7. Remove second dimension gels from cold storage and wash gel tops three or four times with upper tank buffer. 8. Fill wells of cassettes with top tank buffer and load strips. By convention the acidic end of the strip is on the left-hand side of the gel. Ensure no air bubbles are trapped between the gel strip and the gel. 9. Moisten the edges of the gel cassettes, to ease passage through the rubber sealing tubes, and load the gels into the running tank. Use Perspex blanks if 12 gels are not being run. 10. Add top tank buffer, avoiding disturbing the cassette wells, close lid and run using the following parameters: step 1: constant power = 60 W (5 W/gel); time 0.15; temperature = 15°C; pump = AUTO. Step 2: constant power = 120 W (10 W/gel); time 2 h; constant power = 240 W (20 W/gel); time 6 h; temperature = 15°C; pump = AUTO. 11. Stop electrophoresis when the dye-front is approximately 0.5–1.0 cm from the bottom of the gel. Total run time approximately 8 h. 12. Lever open glass cassettes and cut one corner of each gel for orientation. Place gels in appropriate stain (see later). 3.4. Gel Staining

For proteomic analysis of protein spots from 2-D gels the stains used must be compatible with mass spectrometry and as sensitive

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as possible. Several stains are available that meet these criteria, for example Colloidal Coomassie, certain silver stains and various fluorescent stains, e.g. Sypro Ruby™, Orange, Deep Purple and Flamingo. Colloidal Coomassie is not as sensitive as the silver and fluorescent stains, but is easy to use and relatively inexpensive (see Note 6 regarding sensitivity). Silver staining is a multi-step process, but allows less abundant proteins to be visualized. However, abundant proteins may be over-stained as there is no end-point. Also, reproducibility may be poor with silver stain as the degree of staining is dependent on the user. All staining procedures should be carried out on a shaker or rocking table, providing gentle agitation throughout the staining/destaining process. ●

Colloidal Coomassie Brilliant Blue G250 1. Fix gels overnight in 50% (v/v) methanol, 10% (v/v) acetic acid (or TCA). 2. Stain in Colloidal Coomassie stain (diluted in methanol following manufacturer’s instructions) (Bio-rad; 4:1 stain: methanol). 3. Destain in 25% (v/v) methanol, 7% (v/v) acetic acid for 1–5 m. 4. Rinse in 25% (v/v) methanol overnight and store in water.

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Silver stain (13) 1. Fix gels in 40% (v/v) ethanol, 10% (v/v) acetic acid (or TCA) for 30 m. 2. Place in ‘sensitizing solution’; 30% (v/v) ethanol, 0.2% (w/v) sodium thiosulphate, 0.5 M sodium acetate for 30 m. 3. Wash for 3 × 5 m in distilled water. 4. Add silver solution; 0.25% (w/v) silver nitrate for 20 m. 5. Wash for 2 × 1 m in distilled water. 6. Place in ‘developing solution’; 0.24 M sodium carbonate, 200 µl of 37% formaldehyde in 1 l (0.0074% final concentration). 7. Add fresh developer if solution becomes cloudy or after 10 m. Continue development until required staining is obtained. 8. Stop reaction by addition of 0.04 M EDTA for 10 m. 9. Wash twice with distilled water. 10. Store in water.

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Sypro Ruby™ protein stain 1. Fix gels overnight (18 h) in 40% (v/v) methanol, 10% (v/v) TCA.

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2. Stain with Sypro Ruby™ in the dark overnight. Sypro Ruby™ is light sensitive, so gels must be handled in a darkened room and gel storage boxes covered with aluminium foil. 3. De-stain gels in 10% (v/v) methanol, 6% (v/v) TCA. (see Note 7 regarding use and disposal of Sypro Ruby™). 3.5. Gel Imaging and Analysis

3.6. Destaining Gel Pieces Prior to Enzyme Digestion (14)

Many imaging systems are available from different manufacturers. The scanner must have high resolution and be integrated with a spot picker if mass spectrometry of gel spots is to be carried out. Similarly, many software packages are available that permit gel matching and the production of composite (or reference) gels. Spot volumes, peak heights and peak areas may also be obtained from the various packages so that comparisons of treatments may be made. Further statistical analysis may be necessary, for example principal component analysis (PCA), so the data from the gel analysis package must be transferable to Excel spread sheets and compatible with statistical packages. ●

For Coomassie and Sypro Ruby™-stained gel pieces: 1. Incubate in water for 15 m, remove water and incubate in 50:50 water: acetonitrile for a further 15 m. 2. Repeat this washing step. 3. Remove all liquid and cover gel pieces in acetonitrile. Leave for 5 m, by which time the gel pieces should be opaque and dehydrated. 4. Remove acetonitrile and add 0.1 M NH4HCO3 to rehydrate the gel pieces. Leave for 5 m. 5. Add acetonitrile at a ratio of 1:1 acetonitrile: NH4HCO3. Incubate for 15 m. 6. Remove all liquid and speed-vac to dryness.

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For silver-stained gel pieces: 1. Place gel piece in ‘Farmer’s reducing agent’ (20% (w/v) sodium thiosulphate; 1% (w/v) potassium ferricyanide mixed 1:1:1 with water) until gel piece is completely clear. Rinse in water. 2. Remove all liquid and speed-vac to dryness.

3.7. Reduction and Alkylation (14)

1. Incubate gel pieces in 10 mM DTT, 0.1 M NH4HCO3 at 56°C for 45 m to swell. Remove excess liquid and quickly replace with 55 mM iodoacetamide, 0.1 M NH4HCO3 and incubate in the dark at room temperature for 30 m. 2. Remove iodoacetamide solution and wash gel pieces with 1:1 acetonitrile: 0.1 M NH4HCO3 for 15 m. 3. Remove all liquid and speed vac to dryness.

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3.8. Enzyme Digests

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In-gel digests with trypsin (14) 1. Add sufficient trypsin solution to just cover the gel pieces, incubate on ice for 45 m to swell. 2. Remove excess liquid and add 25 mM NH4HCO3/5 mMCaCl2 to just cover the gel pieces. 3. Incubate at 37°C overnight. 4. Peptide isolation: spin down digest and add a minimal volume of 25 mM NH4HCO3. Incubate for 15 m at room temperature. 5. Add an equal volume of acetonitrile, mix and incubate at room temperature for 15 m. 6. Collect the supernatant and repeat the extraction two further times with 5% (v/v) formic acid:acetonitrile (1:1). 7. Pool the supernatants and speed-vac to dryness.

3.9. Desalting and Concentration of Peptides for MS Using Zip Tips

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MALDI-MS: 1. Re-suspend digest in 10 µl 0.1% (v/v) TFA. 2. Wet zip tip with 10 µl 50:50 acetonitrile:water, repeat twice. 3. Equilibrate tip with 10 µl 0.1% TFA, repeat twice. 4. Apply sample to tip by pipetting up and down ×10. 5. Wash tip with 0.1% TFA. 6. Elute with 4 µl 50:50 acetonitrile:water with 0.1% TFA.

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ESI-MS: (15) 1. Re-suspend digest in 10 µl 1% (v/v) formic acid in 4% (v/v) methanol. 2. Wet tip with 10 µl 1% formic acid, repeat twice. 3. Apply protein digest, by pipetting up and down ×10. 4. Wash tip with 10 µl 0.1% (v/v) formic acid. 5. Elute with 4 µl 1% (v/v) formic acid in 70% (v/v) methanol.

3.10. MALDI Target Spotting

1. Mix 1 µl of digest with 1 µl of matrix. 2. Spot onto MALDI target, and allow to air dry at room temperature.

4. Notes 1. TCA should be used instead of acetic acid to fix cereal grain storage proteins as they may be soluble in acetic acid solutions.

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2. Reconstitute trypsin in 50 mM acetic acid, aliquot and store at −20°C. Remove an aliquot and add digest buffer to give the working concentration of approximately 0.125 µg/µl. 3. To precipitate Rubisco: (16) add 60% PEG (w/v) to give a final PEG concentration of 20% (w/v) (e.g. 3.75 ml of 60% PEG 4000 per 10 ml of extract). Add 1 M MgCl2 at a rate of 0.2 ml per 10 ml of extract. Mix gently and keep the mixture on ice for at least 10 m. The Rubisco can then be removed by centrifugation. 4. To re-hydrate dry strips directly in strip holders, place them face down in re-hydration solution (containing sample), ensuring that no air bubbles are present. Cover them with mineral oil and place on lid. Place strip holders on IPGPhor, and programme to re-hydrate for 16–18 h and then to start directly into IEF programme. At the end of the run, remove the strip, wipe off excess oil and either freeze at −80°C or apply to the second dimension gel after reduction and alkylation as described in Section 3.3 (Gel electrophoresis) 5. 5. Silicone sealant may be used to seal the plate edges after assembly. If silicone sealant is used 3–4 days will be required for the sealant to cure. After gel running the sealant must be removed with a razor blade prior to washing the plates. 6. Colloidal Coomassie is sensitive to about 100 ng level, while silver and the fluorescent stains can detect between 10 and 50 ng level (depending upon which literature you believe!). 7. Sypro Ruby™ may be used more than once if gel matching is not required. If gel matching is required it is best to use fresh Sypro Ruby™ as ‘speckling’ of gels may occur when reused. Disposal of Sypro Ruby™: there is no information on the toxicity of Sypro Ruby™ protein gel stain. The stain comprises an organic component and a heavy metal component (ruthenium). For disposal, solutions of Sypro Ruby™ should be acidified with a small amount of glacial acetic acid, poured through activated charcoal or other combustible material and then burned in a chemical incinerator equipped with suitable afterburner/scrubber system to remove the dye. All federal, state and local environmental regulations should be observed when disposing of the dyes.

Acknowledgements Rothamsted Research receives grant-aided support from the Biotechnology and Biological Sciences Research Council (BBSRC) of the UK. This work was supported by the Food Standards Agency under the GO2 programme.

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References 1. Safety Evaluation of Foods Derived Through Modern Biotechnology: Concepts and Principles (1993) OECD, Paris. 2. Kuiper, H. A., Kleter, G. A., Noteborn, H. P. J. M. and Kok, E. S. (2001) Assessment of the food safety issues related to genetically modified foods. Plant J. 27, 503–528. 3. Shewry, P. R., Baudo, M., Lovegrove, A., Napier, J., Ward, J., Baker, J. and Beale, M. (2007) Are GM and conventionally bred cereals really different? Trends Food Sci. Technol. 18, 201–209. 4. Skylas, D. J., Mackintosh, J. A., Cordwell, S. J., Basseal, D. J., Walsh, B. J., Harry, J., Blumenthal, C., Copeland, L., Wrigley, C. W. and Rathmell, W. (2000) Proteome approach to the characterisation of protein composition in the developing and mature wheat-grain endosperm. J. Cereal Sci. 32, 169–188. 5. Amiour, N., Merlino, M., Leroy, P. and Branlard, G. (2002) Proteomic analysis of amphiphilic proteins of hexaploid wheat. Proteomics 2, 632–641. 6. Salt, L. J., Robertson, J. A., Jenkins, J. A., Mulholland, F. and Mills, E. N. C. (2005) The identification of foam-forming soluble proteins from wheat (Triticum aestivum) dough. Proteomics 5, 1612–1623. 7. Vensel, W. H., Tanaka, C. K., Cari, N., Wong, J. H., Buchanan, B. B. and Hurkman, W. J. (2005) Developmental changes in the metabolic protein profiles of wheat endosperm. Proteomics 5, 1594–1611. 8. Ruebelt, M. C., Leimgruber, N. K., Lipp, M., Reynolds, T. L., Nemeth, M. A., Astwood, J. D., Engel, K.-H. and Jany, K.-D. (2006) Application of two-dimensional gel electrophoresis to interrogate alterations in the proteome of genetically modified crops. 1. Assessing analytical variation. J. Agr. Food Chem. 54, 2154–2161. 9. Ruebelt, M. C., Lipp, M., Reynolds, T. L., Astwood, J. D., Engell, K.-H. and Jany, K.-D.

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(2006) Application of two-dimensional gel electrophoresis to interrogate alterations in the proteome of genetically modified crops. 1. Assessing natural variability. J. Agr. Food Chem. 54, 2162–2168. Ruebelt, M. C., Lipp, M., Reynolds, T. L., Schmuke, J. J., Astwood, J. D., DellaPenna, D., Engel, K.-H. and Jany, K.-D. (2006) Application of two-dimensional gel electrophoresis to interrogate alterations in the proteome of genetically modified crops. 1. Assessing unintended effects. J. Agr. Food Chem. 54, 2169–2177. Finnie, C., Mechior, S., Roepstorff, P. and Svensson, B. (2002) Proteome analysis of grain filling and seed maturation in barley. Plant Physiol. 129, 1308–1319. Osborne, T. B. (1907) The Proteins of the Wheat Kernel. Publication No 84, Carnegie Institute. Yan, J. X., Wait, R., Berkelman, T., Harry, R. A., Westbrook, J. A., Wheeler, C. H. and Dunn, M. J. (2000) A modified silver staining protocol for visualization of proteins compatible with matrix-assisted laser desorption/ionization and electrospray ionization-mass spectrometry. Electrophoresis 21, 3666–3672. Jensen, O. N., Wilm, M., Shevchenko, A. and Mann, M. (1999) Sample preparation methods for mass spectrometric peptide mapping directly from 2-DE gels, in. 2-D Proteome Analysis Protocols, (Link, A., ed.), Methods in Molecular Biology Series, Vol. 112, Humana Press, Totowa, pp. 513–530. Kristensen, D. B., Imamura, K., Miyamoto, Y. and Yoshizato, K. (2000) Mass spectrometric approaches for the characterisation of proteins on a hybrid quadrupole time-offlight (Q-Tof) mass spectrometer. Electrophoresis 21, 430–439. Hall, N. P. and Tolbert, N. E. (1978) A rapid procedure for the isolation of Rubisco from spinach leaves. FEBS Lett. 96 167–169.

Chapter 17 Establishing Substantial Equivalence: Metabolomics Michael H. Beale, Jane L. Ward, and John M. Baker Abstract Modern ‘metabolomic’ methods allow us to compare levels of many structurally diverse compounds in an automated fashion across a large number of samples. This technology is ideally suited to screening of populations of plants, including trials where the aim is the determination of unintended effects introduced by GM. A number of metabolomic methods have been devised for the determination of substantial equivalence. We have developed a methodology, using [1H]-NMR fingerprinting, for metabolomic screening of plants and have applied it to the study of substantial equivalence of field-grown GM wheat. We describe here the principles and detail of that protocol as applied to the analysis of flour generated from field plots of wheat. Particular emphasis is given to the downstream data processing and comparison of spectra by multivariate analysis, from which conclusions regarding metabolome changes due to the GM can be assessed against the background of natural variation due to environment. Key words: Metabolomics, NMR spectroscopy, multivariate analysis.

1. Introduction Qualitative and quantitative analyses of metabolites have always played some role in selection of plant lines for desirable qualities. Recently, progress in the relatively new science of plant metabolomics has expanded massively our ability to carry out comprehensive metabolite analysis and to bring these data together with phenotypic, quality trait and other ‘omics’ data in a broader ‘systems’ approach. The technology of plant metabolomics has obvious application in the study of substantial equivalence of new crops, whether GM or not, as well as in other related aspects of plant science such as breeding, QTL analysis, pest and disease resistance and functional genomics. The issue of concern to Huw D. Jones and Peter R. Shewry (eds.), Methods in Molecular Biology, Transgenic Wheat, Barley and Oats, vol. 478 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-379-0_17

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regulatory authorities is the potential of GM to alter metabolism to give harmful compounds. The direct effects of action of the transgene on metabolism can be predicted and readily tested. There is however concern about indirect effects that could arise either from disruption of endogenous genes, or, in cases where new enzyme activities are introduced, new metabolites arising from exposure of the enzyme to a diverse pool of substrates. Metabolomic analytical methods utilise the classic chemical spectroscopies (Nuclear Magnetic Resonance, NMR; Mass Spectroscopy, MS) often coupled with gas, liquid or capillary electrophoretic chromatography systems. Most metabolomics laboratories deploy a number of techniques depending upon the biological application. These techniques can be broadly divided into ‘targeted’ and ‘untargeted’ methodologies (1), and all have been applied to the study of substantial equivalence of GM plants (2–6). Targeted analysis – for particular compounds or classes of compounds – has for a long time underpinned research in areas such as plant metabolic engineering and trait selection from natural collections or within breeding programmes. Targeted analysis has relatively low throughput and is usually preceded by a sample clean-up regime. Furthermore, it often involves specific derivitisation, chromatographic separation as well as use of detector systems selected for optimal response to the analytes of interest. In contrast, the coupling of high-sample-throughput data collection from modern robotic spectrometers with chemometric computational techniques has revolutionised our ability to carry out large-scale untargeted analysis without the purification towards particular compound classes. This technology is particularly suited to screening of large numbers of samples without bias towards particular metabolites, and thus is highly suited to the study of unintended effects introduced by genetic modification as well as for the determination of biomarkers for quality trait selection. Screening by [1H]-NMR and direct infusion electrospray-MS (2, 3, 7) can provide complementary fingerprints from unpurified extracts, and application of these techniques is a good starting point for the determination of substantial equivalence. If used as part of large-scale trials, such screens can be used to investigate differences against a background of environmental variation in the metabolome. Targeted analysis can then be used to follow up any differences highlighted in the primary screens. Here we describe a detailed protocol for [1H]-NMR fingerprinting that we have developed (7) and used for analysis of field-grown wheat (3) and also discuss the key issues of experimental design, sampling, biological and technical replication, as well as provide a detailed guide to processing and multivariate data analysis of NMR data. For a comprehensive guide to the application of multivariate statistics and other pattern matching techniques to NMR data, an excellent review is provided by Lindon et al. (8).

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2. Materials 1. Eppendorf polypropylene tubes, 1.5 ml (Eppendorf UK, Cambridge, UK). 2. [1H]-NMR extraction solvent, prepared in sufficient volume to add 1 ml to each sample in the whole batch, comprising (v/v) 80% deuterium oxide (D2O, 99.9%D, Goss Scientific, Great Baddow, Essex, UK); 20% deuteromethanol (CD3OD, 99.8%D Goss Scientific) and 0.05% (w/v) deuterotrimethysilylpropionate (d4-TSP, Goss Scientific). 3. Clean, dry, 5 mm thin wall NMR tubes. 4. Modern NMR spectrometer operating at [1H] frequency of 400 MHz minimum, with dedicated 1H probes and contemporary software, ideally with batch processing and spectral editing facility (here we utilise XWIN-NMR and AMIX from Bruker Biospin, Rheinstetten, Germany). 5. SIMCA-P multivariate statistical software (Umetrics, Umea, Sweden). Other similar software packages are Pirouette (Infometrix, Bothell, WA, USA) and Spotfire (Spotfire Inc., MA, USA).

3. Methods

3.1. Metabolite Extraction and NMR Sample Preparation

The method described here has been optimised for the analysis of milled white wheat flour, but can be applied with modifications to the analysis of whole grain flours, bran or other tissues such as leaf and root. Key considerations before embarking on a substantial equivalence metabolomics experiment, as with any field or greenhouse trial, include design of plant growth regimes (see Note 1), sampling of the tissue (see Note 2) and sample labelling (see Note 3). 1. From each biological replicate of white flour, weigh three replicate 30 mg (+/−0.03 mg) samples into separate labelled 1.5 ml Eppendorf tubes. Randomise the biological and technical replicates across the experiment (see Note 4). 2. Add 1.0 ml of the NMR extraction solvent (see above) and close tubes. 3. Vortex-mix the contents of the tubes, until the flour is completely dis-aggregated (usually approximately 30 s) (see Note 5). 4. Heat the tubes at 50 ± 1°C for exactly 10 min. This is easily accomplished by use of a polystyrene raft and a pre-heated

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water bath. The tubes should be positioned so that their contents are below the water level of the bath. 5. Immediately after removal from the water bath transfer the tubes to a micro-centrifuge and spin for 5 min. 6. Each tube, transfer 800 µl of the supernatant to a clean labelled 1.5-ml polypropylene tube. 7. Heat-shock the solutions (see Note 6) at 90 ± 2°C for 2 min, using a pre-heated water bath as before. 8. Immediately after removing the raft from the water bath, transfer tubes to a rack and transport to a cold room (4°C) and leave at this temperature for 45 min. 9. Micro-centrifuge while still cold at full speed for 5 min. 10. Transfer 0.70 ml of supernatant to a clean dry labelled 5-mm thin wall NMR tube and cap ready for analysis (see Note 7). 3.2. NMR Data Collection

1. Load NMR tubes into the NMR auto-sampler rack, recording their position in an instrument log book. 2. Ensure that the variable temperature unit within the NMR spectrometer is set to 300K. 3. Enter the sample details into the automation program’s sample list taking care to accurately enter the appropriate sample label. For each entered sample select D2O as the sample solvent, 1024 as the desired number of scans and WATERSUP as the parameter set (see Notes 8–10). 4. Start the automation sequence. The NMR software then automatically loads each sample into the NMR magnet, finds the D2O signal and locks onto it, optimises the intensity of this signal (via an automated shimming routine), sets the receiver gain and then collects the NMR data. At the end of the data collection, the NMR automation routine automatically processes the data and saves the file before proceeding to the next sample (see Note 11).

3.3. NMR Data: Visual Inspection and Quality Assurance

1. At the end of the whole experiment, open AMIX (Analysis of Mixtures, Bruker Biospin, Germany) software and select the File >Open from XWin NMR command. In the pop-up box enter the appropriate file location for the experiment data files and click OK. 2. In the pop-up box select all the entries for the full experiment and click OK. 3. All the NMR spectra should now be displayed in the main window. Inspect the files to ensure that the data have been collected satisfactorily (see Note 12). 4. Re-run any samples identified in step 3 as being unsatisfactory (see Note 13).

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5. Once data have been collected and assessed for quality (see Note 14), NMR samples are removed from the NMR tubes and transferred to screw cap glass vials. These vials are stored in a refrigerator in case future analyses are required. 3.4. Data Processing, Databasing and Spectral Bucketing

Prior to analysis of the data in statistical packages, some further processing of the data is required. In this step data are prepared for inclusion in the Bruker NMR spectrometer’s database (SBase) (part of the AMIX software). The rationale for this step is to ensure a high comparability of the datasets and to reduce the complexity in the data from 32k data-points to a matrix consisting of approximately 1k data-points. This ‘bucketing’ process also negates alignment problems that can sometimes arise from minor chemical shift differences in some signals due to small variation in pH of samples. 1. Open AMIX (Analysis of Mixtures, Bruker Biospin, Germany) software and select AMIX Tools >Prepare Data. 2. In the Prepare data window open files to be added to SBase by selecting File >Open from XWin NMR command. In the pop-up box enter the appropriate file location for the experiment data files and click OK. 3. In the pop-up box select all the entries for the full experiment and click OK. 4. All the NMR spectra should now be displayed in the main window. 5. Zoom in on the d4-TSP peak at δ0.00. This is the internal standard peak. Scale this peak to the maximum height (see Note 15). 6. Reset the zoom so that the whole spectrum width can be displayed. 7. Select the batch-processing function and in the window that pops up select the following options: i. Remove negative peaks ii. Noise reduction using user-defined region δ10.0–10.2 8. Enter the sample name of the first sample (see Note 16). 9. After saving this spectrum to the spectra database (SBase), close the active spectrum and add the sample name for the next spectrum in the list. 10. When all data have been saved to the SBase close the Sample Preparation window and open the ‘Buckets’ Window (AMIX Tools >Buckets). 11. Select ‘Create new bucket table’. In the pop-up box select ‘data from SBase’. 12. A further window will pop up. Locate the data to be bucketed.

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13. In the Bucketing options window select (see Note 17). i. The data range to be bucketed (δ9.5–0.5 ppm) ii. Bucket width (0.01 ppm) iii. Inclusive peaks (all positive peaks) iv. Scaling method (scale to reference region) v. Reference region (δ0.05 to −0.05) vi. Exclusions (none) 14. Open the bucket table using a spreadsheet package such as Microsoft Excel. 15. Add extra row labels to assist in future data analysis (e.g. line, treatment, timepoint). 16. Remove rows corresponding to residual water (δ4.775– 4.865) and methanol (δ3.285–3.335). 17. Save file as an Excel Worksheet. 3.5. Multivariate Analysis (see Note 18)

1. Open SIMCA-P software (other similar software packages can be used – specific operations will be different, but the principles remain the same) 2. Create New Project (File >New) and select the analysis file (created by the steps above) (see Note 19) 3. Transpose the dataset (Commands >Transpose dataset) such that the chemical shifts are the columns and the samples are the row data 4. Highlight the top row of chemical shifts and select this row as the Primary Variable ID 5. Highlight the column containing the sample names and select this column as the Primary Observation ID 6. Highlight any descriptive columns and assign these columns as Qualitative X data 7. Proceed to upload the data. When prompted, do not exclude any data values (see Note 20) 8. Double click on the data model and select the workset button 9. Select the scaling tab, highlight all the data and select scaling method ‘ctr’. This centres the data around zero by subtracting the mean 10. Exclude any qualitative values from the PCA modelling by selecting the variables tab and highlighting the descriptive values. Exclude these entries 11. Auto-fit and inspect the PCA model (see Note 21) 12. The PCA scores plot can now be analysed (see Note 22). Plots of various components should be analysed for different

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clustering patterns. PC1 vs. PC2 should always be examined as these components often explain the largest variance in the dataset 13. By colour coding the variables according to your descriptor information (line, treatment etc) it is easier to see trends in the dataset 14. For each scores plot, the two corresponding loadings plots (see Note 23) should be generated to describe the metabolites responsible for differences in clustering 15. The positive and negative peaks in loadings plots can be assigned in terms of metabolites changing between the clusters. This is achieved by comparison with a library of NMR spectra collected on the same spectrometer, under the same conditions (solvent, temperature, pulse program), from authenticated pure compounds and that has also been bucketed in AMIX under the same conditions (see Note 24) 16. If there is no clear clustering, a discriminant analysis can be performed. This involves assigning classes to the dataset prior to modelling and using this information to ‘force’ differences in the dataset. Corresponding scores and loadings plots can be examined as described earlier (see Note 25) 17. In the process of generating loadings plots, information is gathered on those data-points that are responsible for the differences. These give clues only. Examination of the original data should also be carried out in order to confirm metabolite assignments and changes. All regions of the loadings plot should be examined as in many cases changes in the intensity of very small peaks can be more significant than smaller changes in the very large peaks in the spectrum 3.6. Determining Substantial Equivalence

The scores plot from PCA (an unsupervised technique) allows rapid evaluation of the degree of similarity and differences between the samples. The loadings plot is used to identify those metabolites that contribute to differences. In most cases studied so far (2, 3) cultivar and environmental differences are larger than the effect of the transgene. However, in the case of introduced metabolic enzymes or their regulators the direct effect of the introduced gene on specific metabolites should be taken into account. Building of different PCA models to examine environmental versus GM/non-GM effects will also be required. If GM vs. non-GM scores plots do not form separate clusters, when environmental and variety differences are removed, then they can be regarded as substantially equivalent, within the limitations of the analytical technique. More advanced supervised modelling, e.g. PLS or neural networks (8), can be used to further probe the datasets for differences.

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4. Notes 1. In plant metabolomic experiments, particularly those involving field-grown material where there are strong environmental effects, the plot layout must provide adequate biological replicates and be randomised in such a way as to minimise micro-climatic and soil effects. Ideally the trial should be carried out at more than one geographical location and repeated over several growing seasons. 2. Careful sampling and recording of the plant material is of utmost importance. The metabolome is highly dynamic and all sampling should be carried out at the same time in the photoperiodic cycle and the tissue harvested into liquid nitrogen to arrest metabolism. Pooling of plants or plant parts is best carried out at the point of harvest. Tissue should then be stored at −80°C prior to processing. In this work flour milled from grain from single-field plots consisting of approximately 350 plants was used. 3. Labels of samples should reflect the biology and contain identifiers for line, plot number, treatment and biological and technical replicates. Careful consideration of this before starting the experiment facilitates subsequent data processing, particularly multivariate analysis. 4. The inclusion of analytical replicates and tracking samples, randomised across the experimental array, can help to quality assure the whole experiment. At the data analysis stage the technical replicates should cluster together as should the tracking samples. These samples can be used to assess the reproducibility of the extraction process. 5. Any material not suspended in the solvent at this stage will lead to a higher variability in the extraction process. 6. We have found the heat shock step to be particularly important for cereal grain and flour extracts. Even though the solvent is 20% methanol, hydrolytic enzymes such as α-amylase remain active. The result of this is a change in the carbohydrate profile in the extract with time. This becomes evident from analysis of the NMR spectra of the technical replicates which should be randomised across the sample array. We have demonstrated that the 90°C/2 min heat shock eliminates this problem and the NMR spectra remain stable. The problem is much less pronounced in freeze-dried green tissue, but it is wise to incorporate this heat shock into all metabolomic extraction protocols that include aqueous solvents. 7. The stability of the samples should be assessed prior to undertaking a large experiment. This is achieved by collecting an

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NMR spectrum of a freshly prepared sample and comparing it to a spectrum collected several days later. This important step confirms that the samples will remain stable during the time spent in the auto-sampler prior to data collection. For wheat flour extracts prepared by the described method, samples remain stable for many days. 8. The WATERSUP parameter set is a standard Bruker parameter set. The residual HOD signal is suppressed by pre-saturation using a relaxation delay of 5 s. The parameter set can be modified to set the desired number of scans and the usual solvent so that these settings do not need to be adjusted for each individual sample. 9. In this protocol the NMR spectrometer operated at 399.752 MHz and a 5 mm multinuclear, broad-band 5 mm probe was utilised. 10. Each spectrum consisted of 1,024 scans comprising 32K data points and a spectral width of 4,845 Hz. With higher field instruments the number of scans can be reduced. For example, at 600 MHz with a 5 mm inverse probe, these samples require 128 scans to achieve a similar signal to noise ratio. 11. The WATERSUP parameter set calls upon a separate routine for processing of the spectra. In automation the spectra are automatically Fourier-transformed after the application of an exponential window function using a line-broadening setting of 0.5 Hz. Phasing and baseline correction are also carried out within the automation routine. [1H]-NMR chemical shifts are referenced to TSP-d (at δ0.00) in the x-axis. Spectra are also scaled in the y-axis to the largest peak (TSP in this protocol). 12. Each NMR spectrum is overlaid on top of all the others. When data files from the whole experiment are compared it is easy to spot any files in which the data are problematic. Obvious problems are peak widths which are too wide or data which are extremely noisy with shifted peak positions. Reasons for data inconsistency are samples not locking, linewidth problems often caused by a poor quality NMR tube or interrupted data collection due to power spikes. It should be noted however that relatively few samples actually produce poor quality data. 13. In a large experiment it is wise to ‘QA’ the data as the experiment is in progress so that any samples which need to be re-run can be done so quickly and before the sample is removed from the NMR auto-sampler. To avoid confusion if samples are re-run, they should be done so using the same file name as the original dataset, thus over-writing the poor

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spectrum. This avoids poor quality spectra getting through to the next stage of the analysis and also simplifies the spectra selection process. 14. A typical NMR spectrum from a polar solvent extract of milled wheat grain is shown in Fig. 1. The spectrum is relatively simple compared to that of plant green tissue extracts and comprises predominantly carbohydrate and amino acid signals. To assess the reproducibility of the extraction protocol, spectra from analytical replicates of the same biological sample can be overlaid. Fig. 2 shows what is expected from good analytical reproducibility and as a contrast, spectra from samples with poor analytical reproducibility. 15. As the internal standard was added to the initial solvent mixture added to each tissue sample, scaling to this peak renders all spectra in the experiment comparable. 16. The active spectrum that will be saved to the spectra base is the one at the top of the list. Use the sample name corresponding to that spectrum. After saving the active spectrum it is important to close that spectrum before entering the next sample name in the list. This procedure allows all the datasets to be scaled in the same way with just the different sample names being required to be entered. This whole process can be automated if desired by writing a separate sub-routine which utilises the experiment sample lists used in NMR data collection. For small experimental sets this is not necessary. 17. In datasets where there is variability in the sample tissue (e.g. in moisture or starch content) it can sometimes be useful to scale to total intensity rather than scaling to the reference region. 18. The series of steps suggested for multivariate analysis are not exhaustive. The approach to statistical data analysis depends on the nature of the biological questions being asked. In many cases a series of data models need to be constructed. In all cases, findings from multivariate analysis can be checked by inspection of the original NMR datasets, using AMIX. 19. When opening up the data files, do not exclude the rows that ‘are blank or contain text’. These assist in classifying the samples and colour coding the generated plots. 20. If any regions of the dataset are removed it becomes difficult to compare the output to the libraries of standard compounds as the number of data-points would be different. 21. For a good model R2 and Q2 values should be as near to 1 as possible. When Q2 values start to decrease no further components should be generated.

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22. The scores generated in a PCA model are new variables summarising the X variables. The scores for PC1 and PC2 are orthogonal and completely independent of each other. The score for PC1 explains the largest variation of the X space, followed by PC2, etc. Hence the scatter plot of PC1 vs. PC2 is a window in the X space, displaying how the X observations are situated with respect to each other. Observations near each other are similar, observation far away from each other are dissimilar. This type of plot can show the possible presence of outliers, groups, similarities and other patterns in the data. A simple example of a PCA scores plot is shown in Fig. 3. The data in the example clearly cluster into three groups. Group 1 is well over to the left-hand side of the plot and thus has a low PC1 score. These samples are very different to those in Groups 2 and 3 which have a higher PC1 Score. Samples in Group 2 can be separated from those in Group 3 by virtue of PC2. Group 2 thus has a higher score in PC2 than Group 3. 23. Loadings plots describe the differences in chemical shifts responsible for the separation of clusters in the PCA scores plots. The loadings plots can be represented as two-dimensional scatter plots or as line plots. The line plot format is especially useful as this resembles the initial NMR spectrum having peaks in both the positive and negative directions. An example of two loadings plots from a wheat dataset is shown in Fig. 4. The loadings plot for PC1 describes the separation

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in the horizontal direction of the PCA scores plot. Peaks which are positive in the PC1 loadings plot represent chemical shift intensities which are elevated in those samples residing on the right-hand side of the scores plot. The loadings plot for PC2 describes the separation in the vertical direction of the PCA scores plot. Peaks which are positive in the PC2 loadings plot represent chemical shift intensities which are elevated in those samples residing at the top of the scores plot. This is summarised in Fig. 5. 24. A database of NMR spectra of natural compounds needs to be constructed to enable loadings plot assignment. Once this is in place it is relatively straightforward to compare spectra of standards against the loadings plot. The example shown in Fig. 6 shows a small region of a loadings plot and a matched carbohydrate metabolite. 25. When using discriminant analysis, the data should be divided into a training and validation set to test the robustness of the statistical model. In the scores plot, samples in the training set and in the validation set would be expected to cluster together.

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Loadings Plot Matched Metabolite Fig. 6. Illustration of the interpretation of metabolites in loadings plot using metabolite matching with a library of standards.

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Acknowledgements The authors would like to thank Professor Peter Shewry (Rothamsted Research) for the provision of samples of wheat flour. The work was funded by the Food Standards Agency under their G02 programme and the BBSRC under the GARNet and MeT-RO projects. Rothamsted Research receives grant-aided support from the BBSRC.

References 1. Sumner, L. W., Mendes, P. and Dixon, R. A. (2003) Plant metabolomics: large-scale phytochemistry in the functional genomics era. Phytochemistry 62, 817–836. 2. Catchpole, G. S., Beckmann, M., Enot, D. P., Mondhe, M., Zywicki, B., Taylor, J., Hardy, N., Smith, A., King, R. D., Kell, D. B., Fiehn, O. and Draper, J. (2005) Hierarchical metabolomics demonstrates substantial compositional similarity between genetically modified and conventional potato crops. P. N.A.S. USA 102, 14458–14462. 3. Baker, J. M., Hawkins, N. D., Ward, J. L., Lovegrove, A., Napier, J. A., Shewry, P. R. and Beale, M. H. (2006) A metabolomic study of substantial equivalence of fieldgrown genetically modified wheat. Plant Biotech. J. 4, 381–392. 4. Defernez, M., Gunning, Y. M., Parr, A. J., Shephard, L. V. T., Davies, H. V. and Colquhoun, I. J. (2004) NMR and HPLCUV profiling of potatoes with genetic modifications to metabolic pathways. J. Agric. Food Chem. 52, 6075–6085.

5. Manetti, C., Bianchetti, C., Casciani, L., Castro, C., Di Cocco, M. E., Miccheli, A., Motto, M., Conti, F. (2006) A metabonomic study of transgenic maize (Zea mays) seeds revealed variations in osmolytes and branched amino acids. J. Exp. Bot. 57, 2613–2625. 6. Le Gall, G., Colquhoun, I. J., Davis, A. L., Collins, G. J. and Verhoeyen, M. E. (2003) Metabolite profiling of tomato (Lycoperiscon esculentum) using 1H NMR spectroscopy as a tool to detect potential unintended effects following a genetic modification. J. Agric. Food Chem. 51, 2447–2456. 7. Ward, J. L., Harris, C., Lewis, J. and Beale, M. H. (2003) Assessment of 1H-NMR spectroscopy and multivariate analysis as a technique for metabolite fingerprinting of Arabidopsis thaliana. Phytochemistry 62, 949–957. 8. Lindon, J. C., Holmes, E. and Nicholson, J. K. (2001) Pattern recognition methods and applications in biomedical magnetic resonance. Prog. Nucl. Magn. Res. 39, 1–40.

Chapter 18 Design and Management of Field Trials of Transgenic Cereals Zoltán Bedo˝, Mariann Rakszegi, and László Láng Abstract The development of gene transformation systems has allowed the introgression of alien genes into plant genomes, thus providing a mechanism for broadening the genetic resources available to plant breeders. The design and the management of field trials vary according to the purpose for which transgenic cereals are developed. Breeders study the phenotypic and genotypic stability of transgenic plants, monitor the increase in homozygosity of transgenic genotypes under field conditions, and develop backcross generations to transfer the introduced genes into secondary transgenic cereal genotypes. For practical purposes, they may also multiply seed of the transgenic lines to produce sufficient amounts of grain for the detailed analysis of trait(s) of interest, to determine the field performance of transgenic lines, and to compare them with the non-transformed parental genotypes. Prior to variety registration, the Distinctness, Uniformity and Stability (DUS) tests and Value for Cultivation and Use (VCU) experiments are carried out in field trials. Field testing includes specific requirements for transgenic cereals to assess potential environmental risks. The capacity of the pollen to survive, establish and disseminate in the field test environment, the potential for gene transfer, the effects of products expressed by the introduced sequences and phenotypic and genotypic instability that might cause deleterious effects must all be specifically monitored, as required by EU Directives 2003/701/EC (1) on the release of genetically modified higher plants in the environment. Key words: Wheat, Triticum aestivum L., field experiment, transgenic crops, environmental risk assessment.

1. Introduction Over the last decade, genetic transformation has become routinely used to broaden genetic variation for plant breeding programmes. Developments in wheat transformation systems have made it possible for plant breeders to develop large numbers of transgenic genotypes. The production of transgenic plants with Huw D. Jones and Peter R. Shewry (eds.), Methods in Molecular Biology, Transgenic Wheat, Barley and Oats, vol. 478 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-379-0_18

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improved traits, such as increased resistance to biotic and abiotic stresses and improved quality characteristics, could contribute to the development of new germplasm for breeding. However, the effects of introduced genes on various phenotypic traits require studies in field experiments. Barro et al. (2) found small phenotypic differences in agronomic traits and yield components when a transgenic line was compared with its non-transgenic parent in field trials. In contrast, Rakszegi et al. (3) did not measure any change in yield performance when they studied two related lines, the transgenic line B73-6-1 and its non-transgenic parent L88-6. Sharp et al. (4) showed the Wsm1 gene provided the best source of resistance to wheat streak mosaic virus, but observed variation in yield loss from 11 to 28% in field trials. Transgenes are usually inherited in the T1 generation as a dominant trait, although insertions may occur at multiple loci and Mendelian segregation ratios were not always observed (5). This may account for the fact that heterozygotes and homozygotes were found up to the T5 generation when transgenic genotypes were analysed genetically (6). Breeders may use field experiments to develop backcross generations with the non-transgenic parents to eliminate phenotypic differences or to transfer the introduced gene into secondary transgenic cereal genotypes. The multiplication of transgenic lines to produce sufficient amounts of grain for the detailed analysis of trait(s) of interest is also carried out under field conditions. The methodology for field testing of transgenic genotypes has certain special requirements compared to protocols for conventional breeding including environmental risk assessment. The objective of the present work is to describe the management and design of field trials of transgenic wheat, taking these specific criteria into consideration.

2. Materials Field experiments can be used at several stages of a transgenic breeding programme. 2.1. Growth of Candidate Plants

Candidate transgenic plants that have been transformed but not yet proved to be transgenic (T0), are generally grown under contained greenhouse conditions, but when large numbers of plants are used in routine breeding trials, field experiments may be employed.

2.2. Growth of T1–Tn Transformed Genotypes

The growth of transformed plants of different generations (T) in order to achieve stable homozygotes is carried out in field nurseries. It may be necessary to determine the segregation of transgenes or to check for gene silencing in the transgenic plants, after which the progeny of confirmed transformants can best be grown in head row experiments. In exceptional cases, the single seed

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descent (SSD) method may be used for generation acceleration, but this technique is generally only employed under greenhouse conditions. The integration and inheritance of the transgene must be stable if the aim is to develop germplasm and to breed transgenic varieties. 2.3. Testing of Secondary Transformant Populations and Lines

The development of backcross (BC) generations to transfer the introduced gene into cereal genotypes is important for breeding when the backcross parent has poor regeneration capacity resulting in low transformation frequency. In such cases, a genotype with good regeneration ability but little breeding value is used as the target genome. The BC method can only be used to eliminate differences between the transgenic lines and their original, non-transformed parents (7).

2.4. Determination of Agronomic Performance

Developed transgenic lines can be tested to determine their agronomic performance and stress resistance, using homozygous (T4 or later) progeny populations. In comparison with the original, non-transformed parents, comparison of their phenotypic response in various environments can be carried out in order to test their adaptability and determine their resistance mechanisms. In this way, diverse disease resistance reactions of transgenic genotypes to scab were identified under field conditions where the pathogen pressure was stronger than in greenhouse experiments (8).

2.5. Testing of Homozygous Transgenic Lines for Registration as Commercial Cultivars

Their Distinctness, Uniformity and Stability (DUS) and Value for Cultivation and Use (VCU) need to be established in accordance with the UPOV (International Union for the Protection of New Varieties of Plants) regulations.

3. Methods 3.1. Geographical Location of Release Sites

The release site is a cultivated, fenced area in which the flora and fauna are identical to those of the surrounding cropland with no rare or protected plant or animal species. It is also essential to ensure that there are no officially recognized biotopes or protected areas in the wider vicinity of the release location. The area should also be flat, partly to reduce the danger of run-off from the area and partly to facilitate tillage, making it possible to set up the experiment accurately and reduce experimental errors.

3.2. Description of the Experimental Management

1. Preparation of the experimental field The experimental area should be prepared using the normal procedures for the given crop in the experimental location. A previous

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crop that can be removed in good time, leaving few residues, should be chosen to ensure a high-quality seedbed. Stalk and root residues on the soil surface tend to become entangled in the coulters of the plot-sized seed drills, making it difficult to set the experiment up accurately. A small-plot transgenic experiment should not be conducted on a site used previously for a conventional small-plot experiment. 2. Nutrient supply Fertilizer should be applied as required for normal tillage and crop production practice. The height of the transgenic plants should be taken into consideration when determining fertilizer rates, and tall exotic transgenic plants, which are subject to lodging, should be grown with less than the normal rate of nitrogen fertilizer. Organic manure should not be used on small-plot field experiments as this results in heterogeneity in soil composition. 3. Soil disinfection This is only justified if infestation with pests is likely, and then only if the treatment does not interfere with the aims of the experiment. 4. Experimental design and preparation The first step in setting up the experiment is the preparation of the experimental plan, deciding what genotypes are to be tested and their positions in the design, and specifying plot size, number of replications and experimental design. The seed is prepared on the basis of the sowing list which contains details of the experimental codes and entry numbers of each plot and the origins of the seed (which plot of which experiment). In addition, the name and the pedigree of the genotype, or other distinguishing data, are usually included. For some types of experiment the sowing list also denotes the number of rows to be sown in each plot and the quantity of seed to be sown. 5. Seed preparation Seed preparation depends on the generation of the transgenic plants, whether primary or secondary genotypes are included in the experiment, and the amount of seed that is available. Spikes selected in the previous generation are threshed with a single head thresher or by hand, and the seed is placed in special containers. The seed lots are then put in order, based on the sowing list, and cleaned if necessary, using experimental seed-cleaning equipment designed to exclude the mixing of genotypes. The seed can then be dressed, if required, and the quantity to be sown in each plot is dispensed into bags. The prepared seed lots are stored separately from other experimental materials until sowing. When threshing, cleaning, dressing and counting have been completed, the experimental equipment and rooms are thoroughly cleaned, and any waste seeds and other plant tissues are collected for destruction.

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6. Sowing If only a few seeds are available (e.g. in the T1 and F1 generations), they can be sown with wide spacing by hand or with an individual seed drill. In later generations (T2, Tn), or in segregating populations of secondary transgenic crosses, the grains from single spikes are sown in the next generation in separate head rows. The single seed descent (SSD) method is rarely used for generation acceleration in field experiments, and transgenic homozygous populations are most frequently sown in microplots using a special experimental plot planter or head-row planter. These machines are constructed in such a way that they sow the full quantity of seed with which they are filled (manually or automatically) over a set distance. The sowing density can be adjusted by changing the quantity of seed or the sowing distance. The seeds sown in consecutive plots are not mixed and the equipment does not require cleaning after sowing. The planters close and compact the soil behind the coulter, making further tillage after sowing unnecessary. 7. Separation of transgenic field trials The transgenic plants should be surrounded by a border of at least 2 m consisting of a conventional wheat variety, by at least 20 m of fallow land or a non-cereal crop. After the latest genotype in the experiment has flowered there is no longer any danger that the transgenic and conventional wheats will cross, and the 2 m protective border of wheat plants can be destroyed. 8. Disease spreader border row If the aim of the field trial is to determine the resistance of the transgenic lines to fungal pathogens, a border must be planted consisting of susceptible genotypes suitable for artificial inoculation. In the case of race-specific diseases or pests the border will consist of a mixture of genotypes. The direct inoculation of the transgenic plants with the experimental inoculant is another possibility (3). In this case, some of the transgenic plants in the experiment or one row in each plot are inoculated. 9. Weed control and crop protection A seedling emergence the ends of the plots is cut off straight to achieve the final plot size. This may be carried out by hand, using a rotavator, or using a wide spectrum herbicide. The plots are marked using plastic labels printed using a thermoprinter and attached to stakes. The labels should contain the plot identification data, and any other relevant information, in both visible and barcode forms. When large numbers of plots are harvested at the same time, the labels removed from the plots can be attached to the bags or sacks containing the grain yield, ensuring that the lots can be reliably identified throughout the course of processing. If the local conditions are suitable, it is advisable to apply a pre-emergence herbicide immediately after sowing, especially

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on widely spaced plots easily infested with weeds. Conventional weed control treatments should be applied to control grass and broad-leaved weeds, taking into consideration the dominant species in the local weed flora. Fungicide and insecticide treatments are permitted, depending on the aim of the experiments. The plots must be covered with nets before maturity to ensure that birds do not spread the seeds. 10. Irrigation Wheat usually requires no irrigation. The only exception is when the survival of the transgenic plants can only be ensured by irrigation due to the dry environment. Irrigation of a field experiment may also be required to promote infection when testing the disease resistance of transgenic plants. In some artificially inoculated experiments irrigation is used to ensure high humidity, e.g. in the case of inoculation with Fusarium head blight. Irrigated and non-irrigated plots may also be required for comparison when testing for the effects of drought and heat stress. 11. Observations Field books organized to reflect the experimental design are used to record the data collected in field experiments. As well as data on the identity of the samples, the books should contain any observations or measurements from earlier experiments that could be of relevance. The site should be visited at regular intervals during the growing season according to the experimental protocol, and phenological data should be recorded and compared to those for the non-transgenic standards. The phenotypic responses of transgenic wheat plants developed to have increased resistance to biotic stresses should be recorded as soon as infection is observed. 12. Harvest Depending on the purpose of the experiment, the plots should either be harvested using a small-plot experimental harvester, or spike selection should be carried out on the transgenic plants. The harvested grain should be placed in bags or sacks labelled to ensure the precise identification of the contents. The labels used to identify the plots can be used for this purpose. The bags or sacks containing the grain should be taken to the granary in closed boxes or containers to ensure that no seeds are scattered on the way. The grain of transgenic wheat should be stored separately to non-transgenic material and clearly labelled. An accurate record should be kept of the seed quantities produced and utilized. To prevent mechanical mixing, the harvesters should be thoroughly cleaned before leaving the experimental plot to ensure that no grain or spike fragments remain on the cutting table, or in the drum or sieves. The voluntary emergence of seeds scattered on the soil could be one source of mechanical mixing between transgenic and conventional wheats, so scattering losses

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should be minimized during harvesting and efforts must be made to ensure that spikes are not left in the field. 13. Post-harvest measurements The remaining plant material should be removed or ploughed into the soil. The timing of inversion and tillage will be determined by monitoring the germination of volunteer seeds. Subsequent volunteers should be killed by spraying with an appropriate herbicide or should be removed by ploughing. No crop should be sown the following year and the subsequent crop should be non-cereal or fallow. 14. Emergency measurements In the very unlikely event that any undesirable effects are observed during the trial or that the plants spread, the plant material should be immediately removed or destroyed, and any remaining material ploughed in. If necessary, the plot and an area of 2 m around the plot should be treated with a suitable herbicide. The area should also be monitored for re-growth. 3.3. Environmental Risk Assessment for Cereal Field Trials

Risk assessment is an important part of the experimental field management of transgenic crops. Of particular concern are gene transfer, either to other crop species via pollen or from the plants to soil bacteria, and the effects of the target or marker genes on other organisms. As wheat is a self-pollinating crop, the probability of gene transfer does not seem to be significant. Similarly, no adverse effects of products controlled by introduced sequences have so far been reported. Nevertheless, it is necessary to estimate the probability that potentially harmful effects occur, bearing in mind the environmental conditions. 1. Capacity to survive, establish and disseminate Wheat is an annual plant, so it can only survive via production of grain. Spring wheat is generally sown in March and harvested in July or August, while the sowing time of winter wheat varies from region to region, between September and late November, and the crop generally matures 10–15 days earlier than spring wheat. Early harvest will minimize the dispersal of shed seed or spikes by animals or birds. However, ripe grains that fall to the ground before or during harvest may survive the winter and germinate the next spring. After harvest the straw may be burnt and any plant waste remaining on the field ploughed into the soil. The proteins encoded by the introduced genes and other novel components resulting from their action, are rapidly decomposed by the metabolic processes of soil-borne microorganisms. Shed grains will emerge after the stubble stripping that follows harvesting, and these can then be destroyed by a further tilling operation. They will thus be unable to survive in the long term. The probability of establishment will be minimized by treating the plots with herbicide and removing any re-growth. It is

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recommended that the release location should be sprayed with a 3 l/ha concentration of glyphosate herbicide within 10 days of harvesting and the removal of plant waste. If necessary, this treatment can be repeated. The field should be regularly monitored for the following year, and if any germinated wheat is found it must be removed by hand and burnt. The soil temperature also influences the ability of seeds to survive. In the case of spring wheat, for example, winter frost considerably limits the ability of scattered seeds to survive. 2. Potential for gene transfer by pollen to other wheat plants Under natural conditions wheat is over 99% self-pollinating, so the likelihood that a transgenic line will cross with other wheat varieties is extremely small. The pollen grains are heavy, and under normal circumstances they are not borne by the wind for more than 5–10 m. They are also only viable for 1–3 min, so crosspollination will only take place under extreme weather conditions and within a distance of 10–20 m at the most. 3. Potential for gene transfer by pollen to other plant species There are some concerns that transformed crops will become uncontrollable weeds and that the transgenes will spread into related wild species, including weeds, leading to the development of highly persistent weeds (often called ‘superweeds’). It has even been suggested that all transgenic crops should be prohibited because of the possibility of the transgenes spreading into weeds, even though many crops have no interbreeding relatives in many areas where they are grown. There is a very small probability (approximately 1%) that wheat will be capable of crossing with related wild species (Aegilops, Agropyron, Secale, Hordeum, Elymus) if they flower at the same time. However, there have been no reports of any viable plants being produced from spontaneous crosses of wheat with wild species, and any such hybrids would probably be either sterile or have very low viability. The possibility of cross-pollination can also be minimized by ensuring that no plants of related cultivated species are grown within 50 m of the experimental area. In particular, cultivated rye, which may cross-hybridize with wheat, should not be grown in the close vicinity of the sites. 4. Potential for gene transfer to microorganisms or animals The risk of horizontal gene transfer from plants to soil or gut bacteria under natural conditions is very low and the risk to human health that this represents when compared with the natural occurrence of the genes in bacteria (for example ampicillin resistance) is negligible. The frequently used uidA gene encoding the β-glucuronidase (GUS) enzyme, which is often used as a marker gene for transformation, also occurs widely in microorganisms. The ampicillin resistance gene which is often present on plasmid vectors used for transformation will not be expressed in the

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transgenic plants as it is controlled by a prokaryotic promoter, but this gene is also present in soil bacteria. In the extremely unlikely event that horizontal gene transfer of the ampicillin resistance gene to soil bacteria did occur, it would only impart a selective advantage if the antibiotic were present in the soil environment. 5. Effects of products expressed by the introduced sequences The products of the selectable marker genes may often represent the greatest risk for gene transfer, but it is also necessary to consider the products of the ‘genes of interest’. Because the selective marker genes (e.g. herbicide and antibiotic resistance genes) may be undesirable in the transgenic plants, attention is currently focussed on the development of techniques that can be used to remove the selection marker genes from the transgenic plants. This can be achieved by normal segregation (9), but gene removal using homologous recombination is much more promising. The multiauto-transformation (MAT) vector system was developed for this purpose, allowing transgenic plants free of selectible marker genes to be obtained in a single step (10). However, there are currently no reports of potentially harmful products originating from introduced sequences in transgenic crops, either to the wheat plant or to other organisms in the environment. 6. Phenotypic and genotypic stability Tissue culture and transformation can result in phenotypic effects which are unrelated to the effects of the transgene. Furthermore, these could theoretically pose a risk in field trials. All the transgenic lines selected for field release should therefore be selected to be phenotypically similar to the control plants, apart from the effects of the transgene. Variation can also arise from instability of the transgenes. Where abnormal development of transgenic plants is observed, breeders carry out a BC programme with the parental genotype in order to select lines phenotypically similar to the transformed plant. All the lines selected for field trials should therefore be selected to be genetically stable over several generations. 7. Deleterious effects on non-target organisms Wheat is not generally pathogenic to humans or other species, and the transformed plants should not differ from the parental control plants in pathogenicity or in their effects on humans and other organisms. Susceptible individuals may suffer adverse reactions from consumption of wheat products (allergy or intolerances such as coeliac disease and dermatitis herpetiformis), while respiratory allergy (baker’s asthma) may result from inhalation of flour. Similarly, wheat flour may result in hay fever allergy. However, there is no evidence that these deleterious effects are greater with transgenic wheat. According to our experience, the genetically modified wheat plants behave the same as control, non-transgenic plants in all respects.

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4. Notes Legal conditions for release. The release of transgenic plants into the environment (field testing) requires a special licence. In most countries, particularly EU member countries, licences are only granted by the gene technology authorities to qualified, responsible organizations which have adequate infrastructure to carry out the release. The licence for release is granted for a specific organism (species, variety) and for specific gene insertion events, and is valid for a limited period under specified release conditions. These specific requirements are based on EU Directive 2003/701/EC (1) on the release of genetically modified higher plants in the environment. References 1. EU Directive 2003/701/EC. Commission decision of 29 September 2003. Establishing pursuant to Directive 2001/18/EC of the European Parliament and of the Council a format for presenting the results of the deliberate release into the environment of genetically modified higher plants for purposes other than placing on the market. (notified under document number C(2003) 3405). 2. Barro, F., Barcelo, P., Lazzeri, P. A., Shewry, P. R., Martin, A. and Ballesteros, J. (2003) Functional properties and agronomic performance of transgenic Tritordeum expressing high molecular weight glutenin subunit genes 1Ax1 and 1Dx5. J. Cereal Sci. 37, 65–70. 3. Rakszegi, M., Békés, F., Láng, L., Tamás, L., Shewry, P. R. and Bedö, Z. (2005) Technological quality of transgenic wheat expressing an increased amount of a HMW subunit of glutenin. J. Cereal Sci. 42, 15–23. 4. Sharp, G.L., Martin, J.M., Lanning, S.P., Blake, N.K., Brey, C.W., Sivamani, E., Qu,R. and Talbert,L.E. (2002) Field evaluation of transgenic and classical sources of wheat streak mosaic virus resistance. Crop Sci.42,105–110. 5. Lörz, H., Becker, D. and Lutticke, S. (1998) Molecular wheat breeding by direct gene transfer. Euphytica 100, 219–223.

6. Rooke, L., Steele, S. H., Barcelo, P., Shewry, P. R. and Lazzeri, P. A. (2003) Transgene inheritance, segregation and expression in bread wheat. Euphytica 129, 301–309. 7. Barro, F., Barcelo, P., Lazzeri, P. A., Shewry, P. R., Martin, A. and Ballesteros, J. (2002) Field evaluation and agronomic performance of transgenic wheat. Theor. Appl. Genet. 105, 980–984. 8. Anand, A., Zhou, T., Trick, H. N., Gill, B. S., Bockus, W. W. and Muthukrishnan, S. (2003) Greenhouse and field testing of transgenic wheat plants stably expressing genes for thaumatin-like protein, chitinase and glucanase against Fusarium graminearum. J. Exp. Bot. 54, 1101–1111. 9. Komari, T., Hiei, Y., Saito, Y., Murai, N. and Kumashiro, T. (1996) Vectors carrying two separate T-DNAs for co-transformation of higher plants mediated by Agrobacterium tumefaciens and segregation of transformants free from selection markers. Plant J. 10, 165–174. 10. Ebinuma , H., Sugita, K., Matsunaga, E. , Endo , S. , Yamada, K. and Komamine, A. (2001) Systems for the removal of a selection marker and their combination with a positive marker. Plant Cell Rep. 20, 383 –392 .

Chapter 19 GM Risk Assessment Penny A.C. Sparrow Abstract GM risk assessments play an important role in the decision-making process surrounding the regulation, notification and permission to handle Genetically Modified Organisms (GMOs). Ultimately the role of a GM risk assessment will be to ensure the safe handling and containment of the GMO; and to assess any potential impacts on the environment and human health. A risk assessment should answer all ‘what if’ scenarios, based on scientific evidence. This chapter sets out to provide researchers with helpful guidance notes on producing their own GM risk assessment. While reference will be made to UK and EU regulations, the underlying principles and points to consider are generic to most countries. Key words: Genetic modification, risk assessment, contained use, field release.

1. Introduction Writing a GMRA for the first time can often seem like a daunting task. Where do you go to find help? Will you understand the terminology? Do you know the biology of the crop and its compatibility with wild relatives? Fortunately there are a number of useful resources at hand to help, if you know where to look, where a lot of the hard work has already been done. This chapter provides a step-by-step guide on how to approach writing your own GMRA and highlights a number of the resources that are currently available plus some handy tips to make the task easier. The regulation of GMOs can initially be divided into two parts: (1) contained use and (2) deliberate environmental release (noncontained use). The questions asked in all GM risk assessments (GMRAs) will be similar for both contained and non-contained Huw D. Jones and Peter R. Shewry (eds.), Methods in Molecular Biology, Transgenic Wheat, Barley and Oats, vol. 478 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-379-0_19

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use; however, the depth of supporting information and details required will be far greater for the latter. First let us consider the contained use of GMOs.

2. Contained Use of GM Plants Contained use covers the initial production of a GMO in the laboratory, as well as the use, storage, transport and destruction of the GMO. Typically contained use covers work in laboratories, greenhouses and closed industrial production facilities (see Note 1). Contained use DOES NOT cover GMOs that are deliberately released into the environment for experimental purposes, or products commercially released onto the market. Under the European Community Directive on contained use of genetically modified micro-organisms (see Note 2), all organisations wanting to produce or handle GMOs must notify the relevant competent authorities (see Note 3) of their intention to use their premises for contained use activities for the first time. For example in the UK this would be the Health and Safety Executive (www. hse.gov.uk). As a statutory requirement any institute carrying out GM work should have a ‘Genetic Modification Safety Committee’ in place. If you are carrying out GM work at an organisation that already carries out this kind of research, contact a member of the GM safety committee and request a GM risk assessment form – this is a standard form (often written by the organisation) that you will be required to complete (see Section 2.1 onwards). It is then the responsibility of the Institute Committee to collectively agree that the risk assessment is suitable and sufficient, and the appropriate containment level is selected to minimise any potential risks to negligible or low. GMRA’s are often then made available on organisation intranets – and can provide a useful reference point (or even template) for seeing what level of information is required. For activities involving GM plants, there are no standard containment levels. Such activities are simply classified as ‘notifiable’ or ‘nonnotifiable’ according to the nature of the GM plant itself. Notifiable activity would cover GM plants which pose a greater risk to human health than its unmodified parental organism (see Note 4). This type of work would then require notification to the relevant competent authorities 45 days prior to start of work. Nonnotifiable activity therefore covers GM plants that pose no greater risk to human health than its unmodified parental organism. The degree of risk arising from contained use of the GMO is determined by consideration of the severity of the potential harmful effects on the environment (contained) or to human health along with the possibility of those effects actually occurring.

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The risk assessment will also consider the exposure of humans or the environment to GMOs during the operation of, or possible unintended release from, a contained use facility. A typical GM risk assessment for contained use would therefore cover the following points: 1. General information on the project 2. Risk Assessment for Environmental Protection 3. Risk Assessment for Human Health and Safety 2.1. General Information

1. Description of the work This should include a brief description of the aims of the project and an outline of procedures undertaken (both in making and assessing the GMO). Full information concerning the host organism, inserted genes and vectors used should be given (often this can be fairly generic and indeed it is worth writing a GMRA that will cover all work by a group, i.e. to cover all GM wheat work for example) (see Note 5). 2. Name, status and previous experience of workers The person responsible for the study should be identified, and the names of the staff working on the project and details of their experience provided (step 1 – Section 3.3, see Note 12).

2.2. Risk Assessment for Environmental Protection

1. Hazard identification This is achieved by comparing the GM plant to the non-modified parent and considering and addressing the following points: 1. What is the GM plant’s capacity to survive, establish, disseminate and compete with and/or displace other plants? A summary of the biology (see Note 6), likely interactions with the potential receiving environment and likely effect of modification on these characteristics should be made. For example, will the GM plant be more invasive in the natural environment? Will it exhibit altered survival characteristics in the agricultural environment, requiring different management control strategies? 2. What is the GM plant’s potential to cause harm to animals? For example, will the GM plant be toxic to target organisms (such as insect pests) or harmful to non-target species (such as insect predators)? 3. What is the GM plant’s potential to cause harm to beneficial micro-organisms? For example, will either root exudates or the products of plant decomposition have the potential to cause harm? 4. What is the GM plant’s potential to exhibit altered interactions with plant pathogens and the potential for harm which may arise. For example, will sequences derived from plant viruses be inserted and could these recombine

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with secondary infecting viruses to create a novel pathogen? Will the GM plant become a new host for a plant pathogen, thereby creating a new reservoir in which they can establish? 5. What is the GM plant’s potential to transfer genetic material to other organisms, thereby conferring hazards 1–4 on them? For example, does the GM plant have relatives in the receiving environment with which it can cross-pollinate? 2. Assessment of likelihood The likelihood of a hazard being manifested should be identified as ‘high’, ‘medium’, ‘low’ or ‘negligible’, assuming the basic level of containment is used, for each hazard identified, and giving justifications. 3. Assessment of consequences For each hazard, a description of the consequences to the receiving environment, should such an exposure happen, should be given using the terms ‘severe’, ‘medium’, ‘low’ or ‘negligible’, and justifications given. 4. Preliminary determination of risk to the environment For each hazard, the level of risk should be identified by combining the consequence (assuming exposure) with the likelihood (assuming Level A containment). Express risk as ‘high’, ‘medium’, ‘low’ or ‘effectively zero’. 5. Containment measures to control the risk to the environment If the preliminary risk identified in step 4 – Section 2.2 is ‘low’ or ‘effectively zero’, then the Level A containment measures are appropriate. If the risk is ‘medium’ or ‘high’, additional Level B containment measures should be used. A description of the facilities and the containment measures required to control the hazards should be given (e.g. if air filtration is required, specify the filtering efficiency). 6. Final determination of risk to the environment For each hazard, a reassessment is made of the level of risk assuming that the measures in step 5 – Section 2.2 are in place. The control measures should be such that the overall risk is thereby reduced to ‘low’ or ‘effectively zero’. 2.3. Risk Assessment for Human Health and Safety

1. Hazard identification The following points should be considered and any potential hazards identified, when comparing the GM plant to the parent plant and assuming humans will be exposed. What is the GM plant’s potential 1. To be more toxic to humans than the parent plant 2. To be more allergenic to humans than the parent plant

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3. To exhibit other potential hazards to humans when compared with the parent plant Based on these descriptions, any hazards identified should be itemised within the risk assessment. 2. Assessment of likelihood Assuming the containment measures already specified in step 5 – Section 2.2 are in place, the likelihood of a hazard being manifested should be expressed as ‘high’, ‘medium’, ‘low’ or ‘negligible’ with justifications. 3. Assessment of consequences For each hazard, a description of the consequences needs to be made, assuming that humans will be exposed. The severity of the consequences should be expressed using the terms ‘severe’, ‘medium’, ‘low’ or ‘negligible’. 4. Determination of risk For each hazard, identify the level of risk by combining consequence (assuming exposure) with likelihood (assuming measures listed in step 5 – Section 2.2). Express risk as ‘high’, ‘medium’, ‘low’ or ‘effectively zero’. 5. Containment measures to control the risk to human health If the risk in step 4 – Section 2.3 is ‘low’ or ‘effectively zero’, then the containment measures described in step 5 – Section 2.2 are sufficient. If not, additional containment measures should be used to reduce the risk to ‘low’ or ‘effectively zero’. A description of the proposed facilities and containment measures should be given and how this impacts on the determination of risk.

3. Deliberate Release into the Environment

Genetically modified plants may not be released into the environment without prior approval. The deliberate release into the environment of GM plants, within Europe, is governed by the 2001/18 EC directive (see Note 7). This directive covers two types of environmental release; experimental release (Part B) and commercial release for placing on the market (Part C) (see Note 8). For every authorised Part B and C release, the national authority provides the European Commission with a summary of the key information in the application (SNIF, summary notification information format). The SNIF document is then made public (http://gmoinfo.jrc.it/) for comment (see Note 9). The Public Research & Regulation Initiative (PRRI www. pubresreg.org) has produced an extensive guide to producing a GM risk assessment (see Note 10). Parts of this guidance document are summaries below (Sections 3.1–3.5) – but it is highly

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recommended that this guidance document also be consulted as it provides excellent templates and information to help with writing a GMRA (see Note 10). 3.1. Part B: Experimental Release (Field Trials)

An application for an experimental field trial (Part B) must be submitted to the competent national authority for the Member State in which the trial is set to take place (see Note 11). The accompanying documents (discussed later) must demonstrate that the field trial does not threaten the environment and the surrounding ecosystem. The notification must include a technical dossier supplying information necessary for evaluating foreseeable risks, whether immediate or delayed, which the GMO(s) may pose to human health and the environment (see Note 11). A notification document requires information on the following: 1. The GMO (e.g. recipient organism, details of modification and novel trait) 2. Proposed release and the receiving environment 3. Interactions between the GMO and the environment 4. Monitoring regime and procedures for controlling the release 5. A statement evaluating the impacts and risks posed by the GMO(s) to human health and the environment from the uses envisaged The competent authority of the Member State that receives the notification examines it for compliance with the Directive and evaluates the risks posed by the release. The information provided should be sufficient to enable a decision to be made, but further information and clarification can be sought at any time. A decision must be made within 90 days of receiving the notification, during which period comments may be received from other Member States who are informed of the notification soon after its receipt. The decision on whether or not to grant consent is made on the basis of safety to human health and the environment. No other criteria are considered in the decision-making process. The notifier may proceed with the release only after receipt of written consent of the competent authority and in conformity with any conditions required in the consent.

3.2. Part C: Marketing GMOs in the EU

A Part C notification for placing a GMO on the EU market must be submitted to the competent authority of the Member State where the product is to be placed on the market first. Notifications for placing GMOs on the EU market should, in addition to the information required for a Part B release, provide the following extra information: 1. Extended information taking into account the diversity of sites of use of the product, including information gained from research and development releases carried out under Part B consents

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2. Information concerning the ecosystems that could be affected by the use of the product and an assessment of the risks posed to human health and the environment 3. Conditions for placing the GMO product on the market, including conditions for use and handling and a proposal for labelling and packaging The competent authority then reviews the notification and forms an opinion, which must be within 90 days of receipt. If that opinion is favourable, the lead competent authority will forward the dossier to the European Commission. The Commission circulates the dossier to the other Member States who are given 60 days to evaluate the application in detail, taking into account the particular health and environmental safety issues unique to their territories. If no objections are made, the lead competent authority issues the marketing consent, which applies throughout the European Community. 3.3. General Information in Notifications (both Experimental and Commercial Releases)

General information requirements are necessary for the administrative processing of the notification, as well as information that provides the context of the notification (Note 12). In short, the information needs to address the following questions: 1. Who submits the notification? In most legal systems, permits can be given to legal persons (see Note 13) and sometimes also to natural persons (i.e. individuals). Usually the legal system will require that the ‘notifier/applicant’ be the legal person who carries responsibility and liability for the release, not the researcher. Notifying on behalf of an ad hoc collaborative group of researchers without a legal status is in most cases not possible. It is therefore advisable that the notification be submitted on behalf of a legal entity, such as a department of a university, with an assigned contact person. This may be the responsible researcher or another designated responsible person. 2. What, where and for how long? It is important to make clear in the title what the intended activity is, e.g. a confined field trial with herbicide-resistant maize. Notifications often seek permission to carry out trials for several years, to cover any unexpected hold ups which may delay the trial, such as continued bad weather. Similarly, permits for field trials are often requested for a number of sites on different locations, to allow flexibility in choice of field sites. In those cases it is usually required that for each year the exact locations are notified to the competent authority. 3. Purpose of the activity Notification requirements often include questions referring to ‘purpose of the activity’, which can refer to the purpose of the genetic modification (e.g. insect resistance) as well as to the pur-

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pose of the activity (e.g. performance testing) which is notified. It is useful for decision makers to know what the purpose of the genetic modification is (e.g. insect resistance with expected increases in yield and reduction of pesticide use), to understand the broader context of the request at hand when they are preparing their final decision. 3.4. Technical Information in Notifications

1. Technical information in relation to the risk assessment The technical information is primarily the information that is necessary for the risk assessment and for decisions regarding risk management. Only the information that is directly relevant to the risk assessment and management decision should be provided or cited. The level of detail required should be appropriate to the nature of the activity being considered. Early in development of a GMO, when less information is available, information on risks is limited and so confinement is higher to reduce exposure. Thus, small-scale field trials that determine the efficacy of a gene will require less safety information than commercial releases, but will have a higher level of risk management. Furthermore, because field trials are generally small scale, represent a temporary shortterm exposure to the environment and are carried out under confinement conditions, the likelihood and consequences of risk components can be considered ‘unlikely’ or ‘highly unlikely’, and ‘minor’ or ‘marginal’, respectively. Therefore while risk assessment for confined field trials can be simpler, risk management takes on a more important role. 2. Characteristics of the recipient organism Include a brief summary (one or two paragraphs) in the notification itself with more detailed relevant information in an annex (few pages), with references to existing documentation, databases, etc. Again, the information in the notification and in the annexes should be pertinent to the notification. The annexes would address topics such as: 1. Origin and taxonomy of the recipient plant 2. The use as a crop 3. The genetics 4. Weedy characteristics, including survival, dispersal, volunteers and dormancy 5. Potential for outcrossing – gene transfer 6. Further references, literature and databases cited See again Note 10. 3. Characteristics of the inserted genes and sequences and related information about the donor(s) and the transformation system

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The first step in this process is to specify what has been inserted and incorporated in the genetic material of the recipient plant. There are two approaches: 1. A full molecular characterisation of the transformed plants has been carried out, identifying which part or parts have actually been inserted and integrated in the plant’s genome (see Note 14). 2. A full molecular characterisation has not yet been carried out. In this case it is assumed that the entire construct may have been integrated into the recipient plant, and the risk assessment is conducted on that basis. The second option is particularly important for public research in cases where there are many transformants to test and the release is a small-scale, confined field trial. This approach also allows researchers to submit notifications well in advance and even before the actual transformants are produced. A more detailed characterisation is usually requested when moving to larger scale or less confined field trials, leading to a full characterisation for unconfined (commercial) release. All inserted functional genes are, in principle, relevant to the risk assessment, regardless of whether they are the ‘genes of interest’ or genes that have ‘travelled along’ in the process, such as selectable markers. A gene with a prokaryotic promoter (i.e. which will not be expressed in a plant cell) will also be considered in the risk assessment. Once the relevant inserted sequences are identified, the process continues by listing the following for each inserted gene (see Note 10): 1. The name and abbreviation 2. Origin 3. Resulting new or changed traits (phenotype) and related traits of the donor organism 4. The resulting gene product and its mode of action 4. The resulting GMO In some cases, data about the resulting GMO are available from growing the GMO in growth chambers, greenhouses and/or earlier field trials. Those data may contain useful information for the notification at hand, in particular, data that show whether and to what extent the resulting GMO behaves differently to the nonmodified host plant. However, one should always remain aware that plants can behave differently in contained situations such as greenhouses compared to those growing in the open air. 5. Suggested detection and identification methods and their specificity, sensitivity and reliability

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Detection and identification are important for reasons of monitoring and enforcement; this could focus on the inserted DNA, the resulting proteins or both (see Note 14). 6.The intended use (e.g. field trial or commercial use) The major distinction between placing on the market and field trials is that with field trials, the GMOs involved are still under various degrees of control, whereas after placing on the market for commercial production of a GMO, its use is in principle unrestricted except of course for specific product-use conditions, such as labelling or monitoring. 7. The receiving environment The characteristics of the receiving environment are crucial for the risk assessment. Relevant characteristics for field trials include the following: 1. Comparison between the normal growing environment and proposed field trial environment 2. Specific environmental factors influencing survival and distribution (e.g. climate, soil conditions) 3. Presence of sexually compatible crops and compatible wild relatives 3.5. The Environmental Risk Assessment

The risk assessment methodology typically follows a number of steps (similar to the contained GMRA), i.e. hazard identification; likelihood estimation; consequence evaluation, including a baseline assessment; risk estimation; risk management and consideration of overall risk. These steps are part of a phased approach: Phase 1: Consideration of each of the inserted genes and sequences individually Phase 2: Consideration of the whole plant, including possible synergies and insertion effects and including available empirical information on the resulting GMO Phase 3: Consideration of risk management and overall risk. Phase 1: Consideration of the inserted genes and sequences individually Step 1. Hazard identification After the basic information about inserted sequences is collected the actual risk assessment starts with identifying any potential hazards or potential adverse effects for each of the genes selected for consideration in the risk assessment. It is important to consider the types of potential adverse effects which are scientifically conceivable, given the characteristics of the gene involved and regardless of the likelihood that such effects will actually occur in the proposed release. The question of likelihood will be addressed in the next stage of the risk assessment!

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Examples of the types of potential adverse effects that, depending on the case, are considered in risk assessments for GMOs are: 1. Toxicity: This focuses on the question of whether the expressed product of the inserted gene/sequence can result in toxic effects in the recipient plant in case of incidental consumption by humans or animals. 2. Allergenicity: Whether the expressed products of the inserted gene/sequence can result in allergenic effects arising from cases of incidental consumption of the GMO by humans or animals, or in case of exposure to parts of the plants, such as pollen. 3. Weediness: Can the inserted gene/sequence cause changes in the weedy characteristics of the recipient plant, i.e. can the recipient – due to the genetic modification – become more persistent in agricultural habitats or more invasive in natural habitats? 4. Susceptibility to pathogens: Can the inserted gene/sequence cause changes in the susceptibility to pathogens, which in turn can cause the dissemination of infectious diseases and/or create new reservoirs of pathogens or vectors? 5. Effects on non-target organisms: Can the inserted gene/ sequence cause adverse effects on populations of non-target organisms, for example by indirect effects on the populations level of other insects than the target insects or, where applicable, predators, competitors, herbivores, pollinators, symbionts, parasites and pathogens? 6. Unintended effects on the target organisms: Can the inserted gene/sequence cause unintended adverse effects on the target organisms, such as resistance development? Resistance development is not an adverse effect in itself, unless it impairs other types of treatments such as spraying with microbial pesticides. 7. Can the inserted gene/sequence result in a change in management of the genetically modified crop plant that has a negative impact on the environment? 8. Can the inserted gene/sequence cause adverse changes in biogeochemical processes, such as changes in the nitrogen cycle? 9. Can the inserted gene/sequence cause other unintended adverse effects, such as: 1. reduced effectiveness of an antibiotic used in medicine as result of horizontal transfer of antibiotic-resistance genes?

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2. the development of new virus strains due to the introduction of viral sequences into a plant genome and possible recombination of genetic material? Step 2. Estimation of likelihood The next step in the risk assessment is an estimation of the likelihood that the events will actually occur in the particular case that is being examined. Here an estimation of likelihood is made for each potential adverse effect identified for each of the inserted genes or sequences. Here the term ‘estimation’ is chosen, because exact values for the frequency with which something will happen in nature cannot always be given. While this may be possible in certain risk calculations such as non-target risks, more frequently the risk finding is qualitative on the basis of a ‘weight-of-evidence’ analysis. In the risk assessment, therefore, terms such as ‘highly likely’, ‘likely’, ‘unlikely’, ‘highly unlikely’, ‘negligible’ or ‘effectively zero’ are frequently used to convey the weight-of-evidence finding. The likelihood of a certain inserted gene or sequence actually having a potential adverse effect is influenced by many different factors, such as: 1. The characteristics of the inserted gene: for example, a gene that is not involved in toxicity of the donor organism is very unlikely to cause the recipient organism to be toxic. On the other hand, it is likely that a gene product that is known to be toxic for one insect, such as the endotoxins produced by Bacillus thurigiensis, will also be toxic to other closely related insects as such data describing its toxicity or toxic effects in the GMO will be required. 2. The characteristics of the recipient organism: for example, the potential for outcrossing with wild relatives is negligible for sterile plants or in regions where no cross-compatible relatives exist, but is likely with fertile plants in an environment where cross-compatible wild relatives are present. 3. The characteristics or the scale of the activity: for example, the likelihood of a genetically modified plant with a certain ‘built-in’ pesticide resulting in significant impacts on insects or other organisms other than the target pest is negligible in a small-scale confined field trial, but may be likely in widespread commercial use. In cases where the estimation of likelihood does not result in a clear conclusion, it is sometimes advisable to proceed to the next step of the assessment, by assuming as a ‘worst-case scenario’ that a certain event will occur. For example, rather than spending much time and effort to determine the exact frequency of outcrossing of a certain variety, it can be assumed that if the plant can outcross, then it will outcross. Attention is then focused on the next step in the risk assessment, i.e. what are the potential consequences of such outcrossing?

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Step 3. Evaluation of the consequences The next step in the process is an evaluation of the consequences should these adverse effects be realised. This step is different from the first step, because it evaluates the severity of a certain effect in a particular situation and environment: something that may be of no significant consequence in one environment may be of significant consequence in another. Terms often used in this step of the risk assessment are ‘major’, ‘intermediate’, ‘minor’ and ‘marginal’. Step 4. Estimation of risk The next step in the risk assessment is the evaluation of risk (see Note 11), for each of the identified potential adverse effects. A certain event may be of a very low likelihood, but the consequences could be so severe that the risk is still high. In the absence of quantitative descriptions of likelihood, terms often used in this step of the risk assessment are: high, moderate, low, negligible. Phase 2: Consideration of the GM plant ‘as a whole’ After the systematic ‘gene by gene’ approach, the risk assessment moves to a more ‘holistic’ phase by looking at the plant ‘as a whole’. In this phase, the risk assessment considers: (1) potential synergistic effects of the inserted genes and (2) available data of the GMO itself, including data on insertion effects. 1. Potential synergistic effects Do the introduced traits confer characteristics that may enhance or reduce the effects of the GM plant in the environment? Certain combinations of traits may enhance the potential for adverse effects, whereas other combinations may reduce the likelihood of adverse effects. The use of two different Bt genes, for example, is sometimes applied to reduce the likelihood of resistance development in the target organism. In some instances data for the individual genes and expressed products will be sufficient to understand the potential for synergistic effects, whereas in other cases additional research may be required. 2. Data on the resulting GMO In some cases, data about the resulting GMO are available from growing the GMO in growth chambers, greenhouses and/or earlier field trials. Those data may contain useful information for the notification at hand, in particular, data that show whether – and to what extent – the resulting GMO behaves differently than the non-modified host plant. 3. Possible insertion effects In considering possible effects as a result of genetic modification, some systems – in particular food safety systems – also consider possible effects as a result of insertion of a sequence within a gene, which could interfere with the development or metabolism within the plant. These are often referred to as ‘insertion effects’. Some genes are only usually expressed during stress, and

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changes to these genes – due to disruption of plant genes during the insertion process – may not be noted until a specific stress is experienced. Such effects cannot be predicted and can only be verified, although with a certain degree of uncertainty, by looking at the GMO as a whole. Phase 3: Consideration of risk management and a determination of overall risk. Finally, in cases where the risks involved are not deemed to be ‘negligible’ or ‘marginal’, the risk assessment continues with the next phase, which is a consideration of whether the identified risk is manageable or acceptable. 1. Risk management In this phase of risk assessment, the question to address is whether the identified risks require specific risk management measures. If the answer is yes, then a risk management strategy is defined. For cases where a risk management strategy has been defined, the risk assessment ‘loops back’ to the earlier steps in the risk assessment to determine whether the proposed risk management strategies sufficiently reduce the likelihood or the consequence. Many different strategies are suitable for the risk management of genetically modified plants including: reproductive isolation (removal of flowers, use of isolation distances or border rows), temporal isolation (reduction of the size or duration of an application) and/or special design features such as male sterility. 2. Determination of overall risk In cases where the level of risk is intrinsically not negligible or where the application of risk management would be very costly or difficult, the question arises whether any identified risks are acceptable. Risks can be considered acceptable in two different contexts: in the context of the overall risk to human health and the environment, and in the context of other socio-economic considerations. This is why any identified risk for the environment or human health related to the genetic modification is compared with the risks associated with use of the non-modified recipient. For example, the aforementioned introduced insect resistance may have a potential impact on some non-target insects, but comparison with the practice of spraying synthetic pesticides on the non-modified crop may indicate that the impact on non-target organisms of the spraying practice is far more severe. 3.6. Conclusion

We often hear the phrase ‘case by case’ when GM risk assessments are considered. While it is true that the level of detail and topics addressed in completing a risk assessment for GMOs will be case dependent, it is still possible to follow the scientifically sound, systematic and transparent approach outlined within this chapter. By making use of the resources available, this will make the task of producing your own GMRA both easier and more thorough.

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4. Notes 1. Poly-tunnels, and/or greenhouses where plants are directly planted into ground soil, would potentially be considered a deliberate environmental release and as such would require notification to the relevant competent authorities. 2. Directive 90/219/EEC as amended by Directive 98/81/EC. 3. In the UK the competent authority would be the Health and Safety Executive (http://www.hse.gov.uk). However, the Department for Environment, Food and Rural Affairs (Defra), the Scottish Executive, and the National Assembly for Wales are also involved in the scrutiny of notifications. 4. For example, for a GM plant containing virus sequences, under such circumstances a Plant Health licence may be required and a higher level of containment would be needed. N.B. ubiquitous virus sequences such as the 35S promoter from CaMV would not warrant notification. 5. For example, ‘work will include the transformation of genotypes/subspecies X, Y, Z using disarmed strains of Agrobacterium (strains 1, 2 or 3), containing the gene of interest (plant or bacterial source), with selectable markers (nptII, hpt, etc.)’. 6. See Note 9. 7. Many competent authorities have excellent websites with information on notification and permit or approval procedures. However, in cases where a notification is made for the first time, it is always advisable to contact the competent authority, to explain the intended activities and to seek guidance about the procedure of notifying. 8. The key difference between Parts B and C is that for research and development releases, decisions are made by individual Member States, whereas for placing GMO products on the market, decisions are made by all Member States, which often necessitates a voting procedure to address differences in opinion on risk. The European Food Standard Agency (EFSA) also oversees this stage in Europe. EFSA oversees all Food and Feed applications (under directive 1829/2003/ EC); as well as non-food and feed cases, where agreement has not been met by all member states. 9. SNIF reports are awlso a useful source of information for writing risk assessments, e.g. a maize SNIF report should provide a good description of the biology of the crop. 10. PRRI Guide on notification and risk assessment for releases into the environment of GMOs-2nd edition. http://pubresreg.org/index.php?option=com_content&task=blogcat egory&id=29<emid=40 Annex I of this guide provides

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useful ‘biology of the crop’ summaries for a number of major crops, while Annex II provides examples of information on a number of frequently used genes. This database contains information on the identities and functions of the genes as well as information on environmental risk assessments that have been carried out earlier in relation to these genes. 11. For example, within the UK this would be the Department for Environment Food and Rural Affairs (Defra). (http:// www.defra.gov.uk/environment/gm/regulation). 12. The detailed information required in notifications is set out in an Annex II of the 2001/18/EC Directive, for the EU, or Annex III of The Cartagena Protocol on Biosafety (CPB). The CPB is the internationally recognised protocol for biosafety (http://www.biodiv.org/biosafety/). 13. A legal person is an artificial entity through which the law allows a group of individuals to act as if it were one person for certain purposes. 14. Standard molecular techniques and checklists are available for molecular characterisation. See for example: http://www. aphis.usda.gov/brs/international_coord.html. 15. Examples of protein-based testing methods include: western blot; ELISA lateral flow strip; magnetic particles; protein chips. DNA based testing methods include: Southern blot; qualitative PCR; quantitative end-point PCR; quantitative real-time PCR.

Acknowledgements In compiling this chapter I have consulted a number of publicly available internet resources and guides. I would like to acknowledge that these resources been used extensively to write this chapter, with certain sections being précised or duplicated in full from these guides. Therefore particular thanks goes to the many unknown people who contributed to the guidance notes available via the Public Research and Regulation Initiative (PRRI www.pubresreg.org); the Health and Safety Executive (HSE www.hse.gov. uk); and the Advisory Committee on Releases to the Environment (ACRE - www.defra.gov.uk/Environment/acre/) for the useful information provided. It is recommended for a more in-depth and complete 'guide for notifications and risk assessment of genetically modified organisms' readers should refer to the PRRI website and follow the Working Group - Risk Assessment link where a pdf of this guide can be found. The PRRI guide is a working document and is continually being updated.

Chapter 20 Transgenic Wheat, Barley and Oats: Future Prospects Jim M. Dunwell Abstract Following the success of transgenic maize and rice, methods have now been developed for the efficient introduction of genes into wheat, barley and oats. This review summarizes the present position in relation to these three species, and also uses information from field trial databases and the patent literature to assess the future trends in the exploitation of transgenic material. This analysis includes agronomic traits and also discusses opportunities in expanding areas such as biofuels and biopharming. Key words: Transgenic, transformation, biofuels, biopharming.

1. Introduction The general prospects for transgenic crops have been reviewed extensively in the last few years (1, 2), and this information will not be repeated in the present summary. Instead the focus will be on the latest progress available in both conventional scientific literature, and also that published in granted patents and in patent applications (3, 4). Such information is freely available on-line from a range of international sites (http://www.uspto.gov/ patft/index.html;http://www.bios.net/daisy/bios/50;http:// www.surfip.gov.sg/) and is an excellent means of assessing future commercial trends.

Huw D. Jones and Peter R. Shewry (eds.), Methods in Molecular Biology, Transgenic Wheat, Barley and Oats, vol. 478 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-379-0_20

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2. Data from Field Trial Databases

Information on the status of field trial applications for transgenic crops provides a means of estimating the time course of future commercial priorities and longer term trends. This information is available on-line for each of the main countries where such tests are undertaken. For the USA, access to the USDA APHIS data is most easily accessed through the Information Systems for Biotechnology (ISB) web site (http://www.nbiap.vt.edu/cfdocs/ fieldtests1.cfm). Inspection of these data in March 2007 shows a total of 420 applications for field trials of wheat, 72 for barley and one for oats (from 1998). Data for wheat from January 2006 to March 2007 show 28 applications of which the most recent are given in Table 1. Corresponding data for barley are given in Table 2. Data for the EU are available from http://biotech. jrc.it/deliberate/gmo.asp for trials conducted under Directive 90/220/EEC and http://gmoinfo.jrc.it/for those conducted more recently under directive 2001/18/EC. Data up to 1999 are available from the OECD (http://webdomino1.oecd.org/ ehs/biotrack.nsf/by%20organism?opendatabase).

3. Transformation Methods There are now relatively well-established methods for transformation for wheat (5, 6), barley (7) and oats to a lesser extent (8). Amongst the current themes is the increasing efficiency of Agrobacterium-based methods (9) for both wheat (10, 11) and barley (12, 13) and the use of less sophisticated tissue culture techniques. Additionally there is continuing progress with methods that ensure simpler integration patterns (14–17) and improved co-transformation (18). It can be expected that all these trends will continue, together with progress on plastid transformation (US Patent: 7,186,560) (Table 1). In addition to the more ‘traditional’ methods for nuclear transformation other novel techniques have been developed. The most recently described include laser micropuncture (19) and electroporation (20).

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Table 1 Recent US field trial applications for wheat Number

Applicant

Gene(s)

Trait

07-057-105

Kansas State

?

Scab resistant

07-057-102

Montana State

?

Grain hardness

07-050-104

Rutgers University

?

Fusarium blight

07-038-106

University of Minnesota

Lipid trans. Prot.

Fungal resistant

Trichothecene

Fungal resistant

Glucanase

Fungal resistant

Chitinase and RIP

Fungal resistant

Chitinase and thaumatin

Fungal resistant

Barley RIP

Fungal resistant

07-029-102

Oklahoma State

?

Drought tolerant

06-318-101

USDA ARS

Glutenin

Protein content

06-291-103

USDA ARS

Glutenin

Protein content

06-087-02

Biogemma

?

Starch metabolism

06-073-07

Montana State

ADPG pyrophosphorylase

Yield

06-073-09

Montana State

GUS

Marker

06-073-08

Montana State

Puroindoline

Feed quality

06-065-09

Montana State

Cinn. alc. Dehydrogenase

Lignin

06-046-04

Syngenta

?

Fusarium blight

06-062-05

University of Nebraska

Ribosome inact. Protein

Fusarium blight

06-053-01

Kansas State

Non-express. PR protein

Scab

06-062-06

University of Nebraska

Apoptosis gene inhibitor

Fusarium

4. Agronomic Traits Many of the traits tested in the recent field trials (Tables 1 and 2) comprise agronomic traits for which the full details are unknown, often for reasons of commercial confidentiality. The summaries below provide supporting evidence where available. 4.1. Herbicide Tolerance

Probably the most advanced example of herbicide tolerance is that of wheat resistant to glyphosate. This material has been tested in many locations, and in one recent study (21) it was shown that weed control with glyphosate tended to be better than with

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Table 2 Recent US field trial applications for barley Number

Applicant

Gene(s)

Trait

07-057-113

Montana State

?

Soft endosperm

07-057-112

Montana State

?

Soft endosperm

06-352-102

Washington State

?

Protein

06-111-102

Washington State

Hordein

Protein

06-114-104

Washington State

Trichoderma protein

Rhizoctonia resistant

06-073-11

Montana State

Puroindoline

Feed property

05-340-01

Washington State

Lactoferrin, lysozyme

Protein

06-059-10

University of Minnesota

Trans. initiat. Factor 5A

Stem rust

06-054-11

University of Idaho

Trichoecene acetyltrans

Fusarium

Endochitinase Thaumatin-related prot. Trichothecene efflux pump 06-040-02

USDA ARS

Trichoecene acetyltrans.

Fusarium

Trichothecene efflux pump Endochitinase 06-039-01

USDA ARS

Ac transposable elem.

Marker

05-090-01

Washington State

Lactoferrin, lysozyme

Protein

05-073-07

University of Idaho

Thaumatin-related prot.

Fusarium

Drug-resistant protein Trichoecene acetyltrans. Endochitinase

conventional herbicides, and wheat treated with glyphosate produced approximately 10% more grain than wheat treated with conventional herbicide tank mixes. Such lines, and equivalent ones from durum wheat, have recently received regulatory approval in Canada: ‘…unconfined release into the environment of the CLEARFIELD™ durum wheat events DW2, DW6, and DW12 and use as livestock feed of durum wheat event DW12 are authorized by the Plant Biosafety Office (PBO) of the Plant Products Directorate and the Feed Section of the Animal Health and Production Division as of 4 January 2007. Any wheat lines derived from events DW2, DW6, and DW12 may also be released into the environment and any wheat lines derived from event DW12 may be used as livestock feed…’. Similarly, ‘unconfined release into the

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environment and use as livestock feed of the wheat events BW2552 and BW238-3 are therefore authorized as of 22 June 2006’. Despite these approvals, full commercialization is uncertain. 4.2. Pest Tolerance

4.3. Pathogen Tolerance

Recent relevant studies include those showing the use of cowpea trypsin inhibitor to protect against the grain moth (Sitotroga cerealella Olivier) (22), and potato proteinase inhibitor (PIN2) to protect against nematodes (23). 1. Fungi Most of the recent field trials of transgenic wheat and barley involve tests of fungal resistance (Tables 1 and 2). One such application is that (Application 06-SBI1-295-WHT01-0871MB007-01) from Syngenta for growth of Fusarium-resistant material in Manitoba, Canada. Other published studies include either general resistance (24) or resistance to the following pathogens: stinking smut (Tilletia caries) (25), powdery mildew (Blumeria graminis) (26, 27), Fusarium head blight (Fusarium graminearum) 28–31), wheat leaf rust (Puccinia triticina) (32). Similarly, the expression of barley hordothionin in transgenic oat seeds is claimed to provide resistance to Fusarium (33). In addition to these investigations of specific effect genes, several other studies are designed to optimize the promoter in order to direct expression to the appropriate organ of the plant (Tables 3 and 4). For example, the Lem2 gene promoter of barley directs cell-

Table 3 Selection of recently granted US patents Date

Number

Subject

Assignee

6 March 2007

7,186,892

Sedoheptulose bisphosphatase

Monsanto

6 March 2007

7,186,890

Serine O-acetyltransferase

Du Pont

6 March 2007

7,186,884

Amyrin synthase

Plant Biosci.

6 March 2007

7,186,563

Transcription factor

Purdue RF

6 March 2007

7,186,561

Salt tolerance

BASF

6 March 2007

7,186,560

Plastid transformation

Rutgers

27 February 2007

7,183,457

Mildew Resistance

Du Pont

27 February 2007

7,183,398

Salt tolerance

Sapporo Brew.

27 February 2007

7,183,109

Embryo-specific promoter

Appl. Bio. Inst.

20 February 2007

7,179,964

Increased thioredoxin

University of California

20 February 2007

7,179,963

CLAVATA protein

Pioneer

20 February 2007

7,179,962

Stress tolerance

BASF

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Table 4 Selection of recent US patent applications Date

Number

Subject

Assignee

1 March 2007

20,070,060,869

Embryo sac-specific genes

Advanta1

1 March 2007

20,070,050,864

Rubisco activase

Pioneer

22 February 2007

20,070,044,180

Brachytic2 (Br2)promoter

Pioneer

22 February 2007

20,070,044,176

Nitrate transport

Du Pont

22 February 2007

20,070,044,172

Nitrogen use efficiency

?

22 February 2007

20,070,044,171

Plant improvement

Monsanto

22 February 2007

20,070,044,169

Male/female sterility

Syngenta

15 February 2007

20,070,039,074

Gene targeting

N. Carolina

8 February 2007

20,070,033,677

Bi-directional promoter

Taiwan University

8 February 2007

20,070,033,673

Annexin promoters

Du Pont

8 February 2007

20,070,033,670

Herbicide-resistant wheat

BASF

1 February 2007

20,070,028,327

Endosperm-specific promoters

Biogemma

1 February 2007

20,070,028,320

Plant artificial chromosomes

Chicago University

and development-specific expression tissues of developing seed spikes of cereal grains, especially lemmas and epicarps (34). 2. Viruses Two recent relevant examples are those claiming resistance of transgenic wheat to barley yellow dwarf virus by expression of a composite hpRNA with the dsRNA stem of BYDV GPV replicase gene and the antisense RNA loop homologous of coat protein gene (35) or the pac1 gene from fission yeast (Schizosaccharomyces pombe) (36). 3. Abiotic stress tolerance Compared with pathogen resistance there are fewer studies on abiotic stress. The published research includes the tolerance of mannitol-accumulating transgenic wheat to water stress and salinity (37), and the use of the barley HVA1 gene to confer salt tolerance in transgenic oat (38). Most recently, the HPLC-6 alpha-helical antifreeze protein from winter flounder was rationally redesigned, and a transgenic wheat line that exhibited the highest levels of antifreeze activity was shown to have significant freezing protection even at temperatures as low as −7°C (37). One particularly interesting recent field trial application for transgenic wheat is from Australia (DIR 071/2006) (http://www. ogtr.gov.au/rtf/ir/dir071appsum.rtf). The details are as follows:

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‘Each GM wheat line contains one of six genes encoding proteins expected to enhance drought tolerance. The genes are derived from the plants Arabidopsis thaliana and Zea mays, the moss Physcomitrella patens and the yeast Saccharomyces cerevisiae. For each gene there are two constructs, one driven by a stress inducible promoter and one by a constitutive promoter, giving a total of 12 different gene constructs used to produce the 30 GM wheat lines’. General biotic and abiotic stress resistance is claimed in a barley line with ectopic expression of a small RAC/ROP-family G protein (40), and barley plants overexpressing a 13-lipoxygenase to modify oxylipin signature were shown to have modified senescence (41).

5. Quality Traits 5.1. Protein/Starch

There have been many studies on transgenic cereals with modified seed protein composition. For example, changes in high molecular weight (HMW) glutenin subunit composition can be genetically engineered without affecting wheat agronomic performance (42), and most recently, transgenic wheats with elevated levels of HMW subunit 1Dx5 and/or 1Dy10 were shown to yield doughs with increased mixing strength and tolerance (43). A related study (44) examined the associated chromatin decondensation in two transgenic wheat lines containing about 20 and 50 copies each of the HMW glutenin genes together with their promoters. The degradation of glutenin has also been examined in lines expressing an antisense construct directed against thioredoxin accumulation (45). Another approach to modifying protein quality is exemplified by complementation of a null Pina allele with the wild-type Pina gene sequence as a method to restore a soft phenotype in transgenic wheat (46). There is less information concerning modification of starch quality. In one example, RNA silencing of the Waxy gene results in low levels of amylose in the seeds of transgenic wheat (47). In a more practically advanced example, field evaluation was conducted on transgenic wheat expressing a modified maize ADP-glucose pyrophosphorylase large subunit (Sh2r6hs). Transgenics were field tested over four growing years, in three locations, with varying planting density and irrigation, and the results indicated that significant yield increases were more likely to occur in space-planted, irrigated environments than densely planted, rainfed environments. It was concluded that limited abiotic resources may subsequently limit Sh2r6hs-associated yield enhancement (48). Other associated advances have been made in the qualitative or quantitative production of enzymes in transgenic cereals

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(see also Section 5.2). Examples include production of heat-stable phytases in transgenic wheat (49), and increased expression of the transcription factor HvGAMYB in transgenic barley as a means of increasing hydrolytic enzyme production by aleurone cells and thereby improving malting quality (50). As with other transgenic studies, the success of these projects depends upon the availability of appropriate promoters; one example of recent improvements is the development of a novel cisacting element, ESP, that contributes to high-level endospermspecific expression in an oat globulin promoter (51). 5.2. Biofuels

With the present emphasis on mitigating any effects of global warming, there is extensive international interest in increasing the proportion of energy derived from plant material rather than fossil fuels. 1. Background Biofuels mean either bioethanol (or other alcohols) produced from starch or other carbohydrate polymers (notably cellulose), or biodiesel produced from oils. They are presently used as major fuels in countries such as Brazil, or as additives at low concentration in many other countries. Present international trends, including the latest UN announcement of collaborative programmes, are available from the ISAAA Biofuels Supplement (http://www. isaaa.org/kc/cropbiotechupdate/biofuels/news/2007/03/09. html) and at Biofuel Review (http://www.biofuelreview.com/ content/view/799/2/). It is estimated that the global market is worth around $22bn and is expected to be worth $35bn by 2020, with corn (maize) leading the growth due to expansion of genetically modified (GM) traits and bioethanol demand in the USA and elsewhere (52). Within Europe, the Biofuels Directive issued in 2003 on the promotion of the use of biofuels and other renewable fuels for transport set out indicative targets for Member States. To help meet the 2010 target – a 5.75% market share for biofuels in the overall transport fuel supply – the European Commission has adopted an EU Strategy for Biofuels (http:// ec.europa.eu/energy/res/biomass_action_plan/doc/2006_ 02_08_comm_eu_strategy_en.pdf), and further research is summarized in the EPOBIO consortium (http://www.epobio. net/). The UK industry position is summarized at http://www. bcsbioscience.co.uk/BCS/BIOCMS/BIOcms.nsf/ID/BUNB68BGU4?OpenDocument. Possibly the most advanced of the transgenic lines designed for bioethanol production is Syngenta corn Line 3272. This has been modified to contain corn amylase, which improves the fermentation of the corn starch to sugar in the bioprocessing stages of ethanol production. This product is undergoing elite breeding, field trials and regulatory filings, before a target launch in 2008. Syngenta estimates that the proportion of US corn planting that

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is dedicated to ethanol production will increase from around 10% in 2003 and 2004 to 30% in 2008 and even higher thereafter, as ethanol production ramps up from under 5bn gal/year to close to 15bn gal/year by 2011. Central to the science of novel or GM bioethanol is the utilization of more efficient enzymes to release fermentable sugars from starch or cellulosic biomass. Like many other enzymes used at present in biotechnology and food production, it can be expected that many of these enzymes will be recombinant (GM). Many companies are making significant investments in such technology. In an analysis of economic trends in this area (http://www.purchasing.com/index.asp?layout = articlePrint&articleID = CA6419082) it is stated: ‘Genencor is working on commercial development of enzymes that will be suitable for future cellulosic biofuels production. Similar projects underway at most of the other big players in enzymes, including Syngenta, Diversa, DuPont, Novozymes, and Codexis’. As well as the 3272 corn line, Syngenta hopes to make the necessary enzymes for cellulose-to-biofuels in green plants. Plant-expressed enzymes, it is claimed, ‘can provide the lowest cost capability to make enzymes’ for biofuels, compared with traditional production methods such as fermentation. Diversa, which is collaborating with Syngenta on its biofuels programme, is looking for genes that code for cellulose-degrading enzymes in a variety of sources including the stomachs of cows, which naturally digest cellulose. The most recent announcement of business investment reported on 8 January 2007 (http://www.seedquest. com/News/releases/2007/january/18020.htm), concerned the particular joint venture between Syngenta and Diversa. This new agreement allows Diversa to independently develop and commercialize fermentation-based enzyme combinations from its proprietary platform. Syngenta will have exclusive access to enzymes from Diversa’s platform to express in plants for enhanced cost-effective production. It is likely that much of this technology will be appropriate for use on any biomass whether from corn or other cereals. Issues relating to Identity Preservation in the USA, particularly with respect to corn (including for biofuels) are summarized in a recent USDA report (53). Opposition to the use of GM plants as a source of material for biofuels (Frankenstein fuels) has, not unexpectedly, generated considerable opposition in some parts of the press (http://www. newstatesman.com/200608070031), as well as other campaigning organizations (http://www.i-sis.org.uk/NBR.php). As well as overt political opposition, there is also opposition on the grounds of economic misjudgement (54, 55). In summary it is claimed that the production of maize-derived fuel requires 29% more fuel than it produces; soybean biodiesel requires 27% more fuel than it produces; switch grass uses 45% more fuel than it yields; wood biomass requires 57% more fuel than it produces;

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and sunflower plants require 118% more fuel to produce than they create. General concern has been also expressed about taking 18% of U.S. corn, and more in the future, to produce ethanol for burning in automobiles instead of using the corn as food. These issues are just as relevant to the position in Europe where wheat and possibly barley are potential feedstocks for biofuels. 5.3. Biopharming

Production of pharmaceutically active proteins in plants rather than microbes has received much attention over the last decade (56), although there are few commercial successes to date (57). Amongst the companies involved is Ventria, whose production platform is based on transgene expression in the seed of rice and barley crops. Several US field trials of GM versions of these crops for the production of lactoferrin and other compounds have taken place. The most recent relevant research reports include the expression of the glycosylated F4 (K88) fimbrial adhesin FaeG in barley endosperm (58). This protein can be used to orally immunize piglets against the F4-positive enterotoxigenic Escherichia coli (ETEC) strains that are a frequent cause of porcine postweaning diarrhoea. In another example, single-chain antibody fragments for use in eye drops have been produced at high levels in a commercial wheat variety (59).

6. Concluding Comments There is now extensive experience with transgenic wheat and barley in both conventional agriculture systems and for use in higher value industries. Whether this undoubted potential (60) is successfully realized depends more on regulatory and public acceptance than on any underlying scientific uncertainty. References 1. Dunwell, J. M. (2002) Future prospects for transgenic crops. Phytochem. Rev. 1, 1–12. 2. Dunwell, J. M. (2005) Transgenic Crops: Current and Future Generations. Methods in Molecular Biology, Vol. 286, in Transgenic Plants: Methods and Protocols (Peña, L, ed.), Humana, Totowa, NJ, pp. 377–397. 3. Dunwell, J. M. (2005) Review: Intellectual property aspects of plant transformation. Plant Biotech. J. 3, 371–384. 4. Dunwell, J. M. (2006) Patents and transgenic plants. Acta Hortic. 725, 719–732. 5. Vasil, I. K. and Vasil, V. (2006) Transformation of wheat via particle bombardment. Methods Mol. Biol. 318, 273–283.

6. Wan, Y. and Layton, J. (2006) Wheat (Triticum aestivum L.). Methods Mol. Biol. 343, 245–53. 7. Jacobsen, J., Venables, I., Wang, M. B., Matthews, P., Ayliffe, M. and Gubler, F. (2006) Barley (Hordeum vulgare L.). Methods Mol. Biol. 343, 171–183. 8. Perret, S. J., Valentine, J., Leggett, J. M. and Morris, P. (2003) Integration, expression and inheritance of transgenes in hexaploid oat (Avena sativa L.). Plant Physiol. 160, 931–943. 9. Shrawat, A. K. and Lörz, H. (2006) Agrobacterium-mediated transformation of cereals: a promising approach crossing barriers. Plant Biotechnol. J. 4, 575–603.

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INDEX A Acetosyringone ................... 96–97, 107–108, 111, 117, 120 Act1 (actin promoter) ........................ 24–25, 149, 152, 155, 173, 177–178 AGL0......... ................................................................... 138 AGL1......... .................93–94, 101, 106, 111, 138–139, 141 Allergenicity................................................................... 325 Aminoglycoside antibiotics.............................................. 84 Anthocyanin (C1 + R/B genes)......... ......39, 43–47, 56–57, 105, 111, 112 Antibiotic resistance............ ..... 7–8, 23, 217–218, 313, 325 Antisense......... ............... 186–187, 190, 194, 197, 338–339 Auxin......... .............................................................7, 33, 84 Avidin......... ....................................................229, 232, 238

B Backbone sequences.................... 9, 204, 216–217, 224, 248 Bar (pat) (phosphinothricin acetyltransferase, Basta resistance)................................................... 30 Barley stripe mosaic virus (BSMV)......... ......188, 190–191, 195–197 Binary vector................................... 7–9, 101, 102, 194, 204 Biofuels...........................................................333, 340–342 Biolistics......... ...............3, 9–10, 23–26, 31, 32, 71–89, 125 Biopharming.......................................................... 333, 342 Biotin........... 95, 98, 140, 227–229, 232, 234, 237–238, 262 Brome mosaic virus (BMV).......... ................................ 189

C C1 + R/B genes (Anthocyanin)......... .................. 39, 43–44 Cah (cayanamide hydratase)......... ............................. 24, 30 Callus induction medium.......... 26–27, 127, 130, 132, 135, 142–144, 146, 147 CaMV 35S (cauliflower mosaic virus 35S promoter)......... ......... 24–25, 31, 138, 172, 174, 178 cDNA Microarray Data Analysis......... ................. 257–258 cDNA Synthesis, 249–250, 256, 262 CFP (cyan fluorescent protein).................................. 43, 59 Chromosome......... ................................210, 213, 227–231, 233–238, 240–244, 261, 338 CmYLCV (cestrum yellow leaf curling promoter)....................................... 172, 177 Complex insertions......... ............................................... 204 Constitutive expression...................................149, 173, 178 Contained use......... ............................................... 315–317

Containment glasshouse......... ................28–29, 79, 82, 249 Copy number......... ...................... 9, 93, 105, 107, 109–110, 138–139, 221, 224, 225 Counterstaining......... .....................................232, 235, 240 Cytogenetics......... ..........................................227–228, 230 Cytokinins ......................................................................... 7

D DAPI (2′, 6-diamidino-2-phenylindole), 227–230, 232, 234, 236, 240, 241, 244 Dicer......... ............................................................. 186, 187 Digoxigenin......... ........... 227, 229–230, 232, 234, 237–238 Donor plants...................................................6, 72–73, 138 2-D PAGE (Two dimentional Polyacrylamide Gel Electrophoresis).................................. 273, 277 dsRNA....................................................185–188, 193, 338

E EHA105......... ............................................................... 116 Electroporation.........................................4–5, 53, 106, 334 Environmental risk assessment......... .....305, 306, 311–313, 317–318, 324–328, 330 Epsps (glufosinate resistance)......... ...... 23, 24, 30, 126, 138 Escapes......... ..............................................28, 33, 102, 145

F Field release......... .................................................. 313, 315 Field trial applications......... ...........................320, 334–338 Field trials...................................... 278, 305–314, 320–324, 326, 327, 333–338, 340–342 FISH (Fluorescent in situ hybridization)......... 82, 227–244 Floral transformation............................................. 105–111 Fluorescent microscopy...................................... 55–56, 227

G G418.................. 7, 23–25, 72, 75, 77, 84, 85, 112, 117, 118 Gene silencing......... ................. 58, 185–196, 211, 222, 302 Gene transfer......... 4–5, 7, 93, 105, 305–307, 311–313, 322 GeneChip......... .............................. 248, 251, 261–264, 270 Genotypic dependence......... ..................................... 10–12 Germ line......... ....................................10–11, 71, 105–106 GFP (green fluorescent protein)......... ..... 15, 25, 39, 41–43, 45–46, 56, 58, 66, 82, 85, 133, 172–177, 208 Glufosinate......... ..............23–28, 32, 33, 75, 77, 83–85, 98, 126, 138, 153, 160, 164, 208

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Gluten proteins...................................................... 247, 277 Glyphosate.................... 23–24, 30, 208, 311–312, 335–336 GM risk assessment............................................... 315–329 Gold particles......... ......................... 72, 77–81, 84–88, 125, 128, 130–132, 135, 155 Golden promise......... .............................126, 138, 173–176 Gus (UidA, beta-glucuronidase)................................... 5, 9, 39–41, 46–50, 53, 56, 59–62, 79, 82, 88, 102, 133, 134, 139, 145, 152, 154, 155, 160, 172, 177–178, 204, 205, 208–214, 216–224, 312–313, 335

H Hairpin RNA......... ................ 186, 188, 190, 193–195, 338 Hazard......... .............................................62, 317–319, 324 Helper plasmid......... ................................................... 8, 94 Herbicide resistance........................ 7, 23, 85, 106, 133, 149 Herbicide Tolerance............................................... 335–337 Hormones......................................... 31, 73, 75, 82, 83, 153 Hpt (hph, aph-IV, hygromycin resistance)................ 23–25, 30, 138, 139, 204, 208, 217–221, 224, 607 hva1......... ........................149, 151, 152, 157–162, 165, 338 Hybridization......... ........................... 58, 82, 159, 192, 196, 214–218, 221, 223–225, 227–243, 250, 251, 256–257, 261, 262, 265

Microsalts......... ........................................26, 73–74, 76, 97 MUG (4-methyl-umbelliferyl-â-D-glucuronide)........... 40, 47–49, 61 Multivariate analysis......... ..............289–290, 294–296, 298

N Negative selection......... ......................................15, 30, 194 NMR spectroscopy......... ................................289–298, 302 Non-target organisms......... ............................313, 325, 328 Northern blot analysis......... .................................. 158–159 Npt II (kanamycin resistance).........7, 23, 30, 72, 82, 84, 85, 107, 110–112, 116, 208, 217, 219–221, 329

O Osmolarity......... ...................................................26, 76, 84

P

Immature inflorescences......... ...6, 72, 78, 85, 174, 175, 177 In Planta Inoculation............................................. 115–122 Inducible gene expression......... ..................................... 179 Induction Medium......... 26, 27, 31–33, 78, 81, 95–97, 100, 126–127, 130, 132, 135, 142–144, 146, 147 Inflorescence......... ...............6, 11, 72, 78, 85, 174, 175, 177 Inheritance......................137–138, 149, 151, 157, 203–206, 209–214, 222, 223, 307 Inoculation.............. 8, 93–99, 101, 102, 115–123, 142, 143, 185, 195, 309–310 Isoelectric focussing (IEF)......... .....................275, 280–282 Isolation distances.......................................................... 328

Paromomycin.........................25, 84, 85, 105, 107, 110, 112 Part C notification......... ........................................ 320–321 pat (Bar) (phosphinothricin acetyltransferase, Basta resistance) .....................................23, 25–26, 30, 31 Patents......... ........................... 110, 115, 333, 334, 337–338 Pathogen tolerance......... ............................................... 337 pBRACT......... ............................... 128, 138, 139, 141, 145 PCR Analysis......... .................... 28–29, 122, 123, 157, 214 PDS-1000/He......... .................................................... 5, 80 Pest tolerance......... ........................................................ 337 Physical mapping........................................................... 227 Picloram......... .......................................... 27, 33, 84, 96, 97 pmi (phosphomannose isomerase)......... .............. 24–25, 30 Positive selection...................................................15, 24, 30 Post-transcriptional gene silencing (PTGS)......... . 185–186 Probe labelling......... .......................................231, 237, 242 Promoters............... 3, 7, 12–13, 24, 41–42, 45, 47, 58, 138, 171–179, 193, 247, 248, 338–340 Proteomics......... .................................................... 273–286 Protoplast.................................4–5, 7–8, 48, 52, 60, 61, 203 pSTARLING vectors......... ............................190, 193, 194

L

Q

LBA 4404......... ............................................................. 138 LUC (lucifierase)......... ............................ 15, 25, 39–42, 46, 47, 50, 52–56, 60, 63, 64, 66

Quality traits...........................................289–290, 339–342 Quantitative Real-Time PCR......... ...............191, 196, 330

I

M Macrocarriers........... 40, 80, 87, 88, 128, 131, 135, 156, 164 Macrosalts........................................... 26, 73–74, 76, 96, 97 Marker genes......... ...............5, 7, 15, 23–25, 39, 72, 82, 85, 172, 204, 208, 247–248, 251–252, 311–313 Mass spectrometry......... ........... 57, 273, 276, 283–285, 290 Mendelian inheritance............ 149, 209, 210, 212, 214, 223 Metabolite extraction............................................. 291–292 Metabolomics......... ............................................... 289–303

R Real Time PCR......... .............................191, 196, 264, 330 Regeneration...........................3, 5, 6, 10–12, 26–28, 31–33, 58, 71, 72, 76–77, 79–82, 85–89 Regeneration medium.........26–28, 33, 81, 82, 88, 100, 122, 132, 133, 144, 153 Regulatory elements......... ..........................12–13, 171, 178 Reporter genes......... ...................... 3, 12–13, 15, 39–58, 64, 82, 85, 171–174, 176–179, 204, 208, 210, 212 Rifampicin......... .............................................106, 107, 141

TRANSGENIC WHEAT, BARLEY AND OATS 349 Index RNA extraction......... .....................249, 253–256, 261–262 RNA interference......... ..................185–190, 192–195, 197 RNA-induced silencing complex (RISC)......... ..... 186–187 RTBV (rice tungro bacilliform virus promoter)............. 172

S Scutellum..................................... 5, 6, 72, 78, 98–100, 102, 121, 130, 131, 134–135, 143, 146, 175, 176 SDS-PAGE (SDS-Polyacrylamide Gel Electrophoresis)......... ......... 249, 253, 275, 282–283 Segregation......... ....157, 158, 203, 207–209, 211, 222–223, 227–229, 305–307, 313 Selectable marker genes......... ....... 23–25, 72, 203–204, 313 Shoot multiplication medium......... ....................... 152, 153 Shoot tip cultures................................................... 154–156 Silicon carbide fibres...................................................... 4–5 Somatic embryogenesis.........................................33, 71, 88 Southern analysis......... ..............................32, 82, 100–101, 123, 203, 214–220, 223–224 Spectinomycin......... .............................................. 106–107 Spermidine......... ................................. 77, 79, 84, 128, 130, 135, 155, 249, 267 Substantial Equivalence......... ....................13–15, 247–270, 273–287, 289–302

T TaqMan......... ........................................................ 264, 270 Ti plasmids......... ......................................................... 7, 93 Tissue culture......... ......................... 3, 8, 10–12, 23, 26, 27, 31, 33, 34, 72–73, 82, 84, 86, 87, 89, 95, 105, 117–118, 121–122, 126–127, 129, 132, 134, 140–141, 144–146, 151, 204, 227, 313, 334 Tissue-specific expression.......................174–176, 178–179 Toxicity......... ................................14, 39, 44, 58, 64, 66, 68, 267, 287, 317, 318, 325–326 Transcriptomics......... ............................................ 247–270

Transgene expression......... ..........................8, 13, 171–179, 208, 222–223, 247, 342 Transformation efficiency......... .......................6, 11–12, 88, 100–101, 123, 146 Transgene copy number......... ............. 9, 138, 203, 221, 224 Transgene inheritance............................................ 157–158 Transgene loci........................................203–204, 207–211, 213–216, 218, 222–224 Transgenic locus......... ...........................203, 208–210, 213, 216–218, 220–224 Transient expression......... .....................5–6, 41–42, 44–46, 56–57, 72, 86, 134–135 Transition medium......... ................ 132, 134, 143–144, 147

U Ubi1 (ubiquitin promoter)......... ..................24–25, 31, 128, 138–139, 173–174, 177–178

V Vernalization.........................................73, 94–95, 100, 101 VIGS (virus-induced gene silencing,virus-induced gene silencing)......... ...................185–191, 195, 197 Vitamins......... .....73–74, 76–77, 96–97, 116–118, 152, 205

W Weediness......... ............................................................. 325 Western Blot Analysis......... .................................. 158–160

X X-gluc (5-bromo-4-chloro-3-indolyl-âD-glucuronide)......... .........................40, 47, 49–50, 154, 160, 205

Y YFP (yellow fluorescent protein)......... ...........43, 46, 59, 65