Plant Breeding Reviews (Volume 31)

PLANT BREEDING REVIEWS Volume 31

edited by

Jules Janick Purdue University

PLANT BREEDING REVIEWS Volume 31

Plant Breeding Reviews is sponsored by: American Society of Horticultural Science International Society for Horticultural Science

Editorial Board, Volume 31 I. L. Goldman C. H. Michler Rodomiro Ortiz

PLANT BREEDING REVIEWS Volume 31

edited by

Jules Janick Purdue University

Copyright # 2009 by John Wiley & Sons, Inc. All rights reserved. Wiley-Blackwell is an imprint of John Wiley & Sons, Inc., formed by the merger of Wiley’s global Scientific, Technical, and Medical business with Blackwell Publishing. Published by John Wiley & Sons, Inc., Hoboken, New Jersey Published simultaneously in Canada No part of this publication may be reproduced, stored in a retrieval system, or transmitted in any form or by any means, electronic, mechanical, photocopying, recording, scanning, or otherwise, except as permitted under Section 107 or 108 of the 1976 United States Copyright Act, without either the prior written permission of the Publisher, or authorization through payment of the appropriate per-copy fee to the Copyright Clearance Center, Inc., 222 Rosewood Drive, Danvers, MA 01923, 978-750-8400, fax 978-750-4470, or on the web at www.copyright.com. Requests to the Publisher for permission should be addressed to the Permissions Department, John Wiley & Sons, Inc., 111 River Street, Hoboken, NJ 07030, 201-748-6011, fax 201-748-6008, or online at http://www.wiley.com/ go/permission. Limit of Liability/Disclaimer of Warranty: While the publisher and author have used their best efforts in preparing this book, they make no representations or warranties with respect to the accuracy or completeness of the contents of this book and specifically disclaim any implied warranties of merchantability or fitness for a particular purpose. No warranty may be created or extended by sales representatives or written sales materials. The advice and strategies contained herein may not be suitable for your situation. You should consult with a professional where appropriate. Neither the publisher nor author shall be liable for any loss of profit or any other commercial damages, including but not limited to special, incidental, consequential, or other damages. For general information on our other products and services or for technical upport, please contact our Customer Care Department within the United States at 877-762-2974, outside the United States at 317-572-3993 or fax 317- 572-4002. Wiley also publishes its books in a variety of electronic formats. Some content that appears in print may not be available in electronic formats. For more information about Wiley products, visit our web site at www.wiley.com. Library of Congress Cataloging-in-Publication Data: ISBN 978-0-470-38762-7 (cloth) ISSN: 0730-2207 Printed in the United States of America 10 9 8 7 6 5 4 3 2 1

Contents

Contributors 1. Dedication: Anthony H. D. Brown Conservation Geneticist

ix 1

Reid G. Palmer and Jeff J. Doyle I. Biographical Sketch II. Research Accomplishments III. The Man IV. Honors and Awards Selected Publications of Anthony H. D. Brown

2 6 10 12 12

2. Brassica and Its Close Allies: Cytogenetics and Evolution

21

Shyam Prakash, S. R. Bhat, C. F. Quiros, P. B. Kirti, and V. L. Chopra I. II. III. IV. V. VI.

Introduction Cytogenetics Genome Manipulation Wide Hybridization Cytoplasmic Substitution and Male Sterility Genome Dissection and Development of Chromosome Addition Lines VII. Mitochondrial Genome VIII. Plastid Genome IX. Potential Role of Arabidopsis thaliana in Brassica Improvement X. Chloroplast Genomes and their Phylogenetic Implications XI. Evolution of Morphological Characters XII. Concluding Remarks Literature Cited

24 26 56 71 95 104 110 113 114 123 137 142 146 v

vi

CONTENTS

3. Genetic Enhancement for Drought Tolerance in Sorghum

189

Belum V. S. Reddy, S. Ramesh, P. Sanjana Reddy, and A. Ashok Kumar I. Introduction II. Breeding for Drought Tolerance III. Selection among Cultivars and Landraces IV. Breeding for Drought Escape V. Growth Stage–Specific Screening Techniques VI. Physiological Response Traits for Drought Tolerance VII. Marker-Assisted Breeding for Drought Tolerance VIII. Outlook Literature Cited

4. Breeding for Resistance to Stenocarpella Ear Rot in Maize

189 190 194 197 199 207 210 212 214

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Johannes D. Rossouw, Z. A. Pretorius, H. D. Silva, and K. R. Lamkey I. Introduction II. Distribution and Importance III. Pathogen IV. Epidemiology V. Disease Management VI. Summary and Conclusion Literature Cited

5. Cassava Genetic Resources: Manipulation for Crop Improvement

224 225 229 232 233 240 241

247

Nagib M. A. Nassar and Rodomiro Ortiz I. II. III. IV.

Introduction Wild Manihot Species: A Botanical Review Interspecific Hybrids Cassava Diversity as Revealed by DNA Markers and Genetics V. Trait Transfer VI. Outlook Literature Cited

248 252 253 257 262 267 268

CONTENTS

6. Breeding Roses for Disease Resistance

vii

277

Vance M. Whitaker and Stan C. Hokanson I. Introduction II. Causal Pathogens III. Resistance Screening IV. Breeding V. Molecular Tools VI. Future Prospects Literature Cited

7. Plant Breeding for Human Nutritional Quality

277 279 288 298 305 313 316

325

Philipp W. Simon, Linda M. Pollak, Beverly A. Clevidence, Joannne M. Holden, and David B. Haytowitz I. Introduction II. Sources of Nutrients III. Progress in Breeding for Nutrient Content and Composition IV. Plant Breeding Strategies for Increasing Intake of Shortfall Nutrients Literature Cited

327 328 350 374 377

Subject Index

393

Cumulative Subject Index

395

Cumulative Contributor Index

415

Contributors

S. R. Bhat National Research Centre on Plant Biotechnology, Indian Agricultural Research Institute, New Delhi 110012 India V. L. Chopra National Research Centre on Plant Biotechnology, Indian Agricultural Research Institute, New Delhi 110012 India Beverly A. Clevidence Food Components and Health Laboratory, United States Department of Agriculture—Agricultural Research Service, Beltsville Agricultural Research Center, Beltsville, Maryland 20705 USA Jeff J. Doyle L. H. Bailey Hortorium, Department of Plant Biology, Cornell University, Ithaca, New York 14853 USA David B. Haytowitz Nutrient Data Laboratory, United States Department of Agriculture—Agricultural Research Service, Beltsville Agricultural Research Center, Beltsville, Maryland 20705 USA Stan C. Hokanson University of Minnesota, Department of Horticultural Science, 1970 Folwell Avenue, St. Paul, MN 55108 USA Joannne M. Holden Nutrient Data Laboratory, United States Department of Agriculture—Agricultural Research Service, Beltsville Agricultural Research Center, Beltsville, Maryland 20705 USA P. B. Kirti Plant Science Department, University of Hyderabad, Hyderabad, 500046 India A. Ashok Kumar International Crop Research Institute for the Semi-Arid Tropics (ICRISAT), Patancheru 502 324, Andhra Pradesh, India K. R. Lamkey 2101 Agronomy Hall, Iowa State University, Ames, Iowa, USA Nagib M. A. Nassar Departamento de Genetica e Morfologia, Universidade de Brasilia, 70919 Brasilia, Brazil Rodomiro Ortiz Centro Internacional de Mejoramiento de Maiz y Trigo (CIMMYT), El Batan, Texcoco, Apdo. Postal 6-641, 06600 Mexico, D.F. Mexico Reid G. Palmer United States Department of Agriculture, Agricultural Research Service, Corn Insects and Crop Genetics Research Unit, Department of Agronomy, Iowa State University, Ames, Iowa 50011 USA Linda M. Pollak Corn Insects and Crop Genetics Research Unit, United States Department of Agriculture, Agricultural Research Service, Department of Agronomy, Iowa State University, Ames, Iowa 50011 USA Z. A. Pretorius Department of Plant Sciences, University of the Free State, Bloemfontein, 9300 South Africa ix

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CONTRIBUTERS

Shyam Prakash National Research Centre on Plant Biotechnology, Indian Agricultural Research Institute, New Delhi 110012 India C. F. Quiros Department of Vegetable Crops, University of California, Davis, California 95616 USA S. Ramesh International Crop Research Institute for the Semi-Arid Tropics (ICRISAT), Patancheru 502 324, Andhra Pradesh, India Belum V. S. Reddy International Crop Research Institute for the Semi-Arid Tropics (ICRISAT), Patancheru 502 324, Andhra Pradesh, India P. Sanjana Reddy International Crop Research Institute for the Semi-Arid Tropics (ICRISAT), Patancheru 502 324, Andhra Pradesh, India Johannes D. Rossouw Monsanto Singapore Co (PTE) Ltd., 151 Lorong Chuan 06-08 New Tech Park, Singapore H. D. Silva Monsanto Brazil, Rodovia Uberlaˆndia-Araxa´, Uberlandia, MG, Brazil Philipp W. Simon Vegetable Crops Research Unit, United States Department of Agriculture, Agricultural Research Service, Department of Horticulture, University of Wisconsin, Madison, Wisconsin 53706 USA Vance M. Whitaker University of Minnesota, Department of Horticultural Science, 1970 Folwell Avenue, St. Paul, MN 55108 USA

Anthony H. D. Brown

1 Dedication: Anthony H. D. Brown Conservation Geneticist Reid G. Palmer United States Department of Agriculture Agricultural Research Service Corn Insects and Crop Genetics Research Unit Department of Agronomy Iowa State University Ames, Iowa 50011 USA Jeff J. Doyle L. H. Bailey Hortorium Department of Plant Biology Cornell University Ithaca, New York 14853 USA

I. BIOGRAPHICAL SKETCH II. RESEARCH ACCOMPLISHMENTS A. Conservation Genetics B. Plant Mating Systems and Population Structure III. THE MAN IV. HONORS AND AWARDS ACKNOWLEDGMENT SELECTED PUBLICATIONS OF ANTHONY H. D. BROWN

This volume of Plant Breeding Reviews is dedicated to Anthony (Tony) H. D. Brown, known internationally for his research in conservation and population genetics and plant breeding. Dr. Brown’s primary contributions in the area of conservation genetics followed two major themes: optimum sampling strategies and core collections. His life’s

Plant Breeding Reviews, Volume 31 Edited by Jules Janick Copyright & 2009 John Wiley & Sons, Inc. 1

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activities in this area were inspired primarily by his long friendship and close working relationship with Sir Otto Frankel. His research in population genetics focused on the estimation of mating systems and their impact on plant population structure while his research in breeding was on the use of wild relatives in crop improvement. Dr. Brown started with the Commonwealth Scientific and Industrial Research Organization (CSIRO) in 1972 as a research scientist and retired as a chief research scientist in 2006. He is now an Honorary Research Fellow in the Centre for Plant Biodiversity Research, CSIRO Plant Industry, Canberra, Australia.

I. BIOGRAPHICAL SKETCH Tony Brown was born on November 25, 1941, in Waverley, Sydney NSW, Australia. It was wartime and his father, Arthur Brown, was in Darwin, Australia, serving as Squadron Leader in the Royal Australian Air Force. Arthur was from three or more generations of Australian stock. Tony’s maternal grandfather, Hugh Milligan, son of Scottish immigrants, was an eminent primary school headmaster. Hugh’s task was to register Tony’s birth, the agreed name being Anthony Hugh Dean Brown. Hugh urged that the last two names be hyphenated because plain ‘‘Brown’’ was insufficiently distinguished for a future Macquarie Street specialist doctor. However, Tony’s mother Joyce intervened and said, ‘‘Plain Brown is good enough for me, it should be for my son.’’ This ensured that name hyphenation could await future needs. Yet Hugh had other major influences on Tony, inspiring a love of plants, of arithmetic shortcuts, and of parsing sentences. That three initials were an encumbrance emerged later in the United States, where names and official forms were triplet coded, allowing only one middle initial. And the inevitable inversion happened after publications on alcohol dehydrogenase, when the AHD became ADH, which spawned a growing list of mutant miscitations in the Science Citation Index. In high school Tony was inspired to seek a research career as an agricultural chemist after watching a film made by CSIRO on how the discovery of remedying trace element deficiencies in some depleted soils of South Australia converted them to cropping. Consequently, Tony enrolled in the Faculty of Agriculture, University of Sydney, as a salaried employee of the Colonial Sugar Refining Company. This was an era when companies were competing to recruit future graduates. At Sydney, inspirational lectures on Mendelian genetics by Professor ‘‘Spinny’’ Smith-White led to a final fourth year in genetics. Jim Peacock, later to become Chief CSIRO Plant Industry, was in his final year as Spinny’s PhD

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student. He shaped the honors course, and infused enthusiasm as the DNA era was unfolding. Jim had spent some time in Adelaide University, and brought back insights from meeting Sir R. A. Fisher and the team of population genetics students there. Jim devised an unforgettable experiment for the honors genetics class with 16 blue and 16 yellow plastic beads in a jar to simulate genetic drift theory. The jar was shaken and inverted 16 times and the color of the first two beads noted. After 16 repeats, the jar was opened and its contents adjusted to the new observed gene frequency. Over 50 population replications were run over tens of generations, or until fixation blissfully occurred. Why was fixation happening faster than predicted? Sampling with or without replacement? Late into the night the rattle of balls in the jar resounded down the college corridors, until crash. . .extinction: The neck of the jar wore through and broke. However, the experiment had sown the seed of a lifelong interest in sampling issues. On graduating (in 1963), Tony was assigned by the Colonial Sugar Refining Company to its sugarcane experiment station in Lautoka, Fiji. This was a major transition, from collegiate to colonial life, and he was fully briefed at the head office in Sydney on how to behave toward the local population. The company itself was in transition, hiring local staff as officers, and the country was preparing for independence. Tony’s immediate boss was Joe Daniels, a sugarcane breeder respected around the world and a scholarly and imaginative leader. All communications were directed through the mill manager, including scientific reports. Tony had an early lesson in communication when management enrolled him in an in-house training course on report writing. The fact that management chose Tony’s report on fiber content to be one anonymous example of bad writing firmly made a point. It was an object lesson in the ‘‘Gunning fog index,’’ which is a function of the average length of sentences and the number of words with more than two syllables. The index is intended to equate to the number of years of education that a reader requires to understand the writing. Clearly no one in the head office had sufficient schooling to read Tony’s report. Joe Daniels introduced Tony to the magnet of wild crop relatives. It may seem hard to imagine in today’s bottom-line corporate world, but Joe was able to get the company’s approval for a visionary, ambitious project entitled ‘‘Re-creating the Pathway of Evolution of Sugarcane’’ by deliberately selecting Saccharum spontaneum for sucrose content. To achieve the screening of large populations for sucrose, Joe and Tony installed an autoanalyzer to run around the clock. For two weeks, Tony was on an unforgettable night shift feeding thousands of ethanol extracts into the analyzer and reading the output. That and the project to tag leafhoppers, vectors of Fiji disease, with radiolabel P-32

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to track their movement in cane fields in retrospect seem incongruous in a small island colony. After nearly four years in Fiji, Tony returned in 1966 to academia and graduate school. He chose to work with Dr. Robert W. Allard at the University of California, Davis, primarily because of his classic plant breeding book and his research on quantitative genetics. On arrival, however, Tony found that Professor Allard was convinced that the new isozyme technique would open the door to empirical population genetics in plants. Professor Allard recommended that the PhD project should not be on quantitative genetics but on isozyme variation in Zea mays. This would fit better with his assigning Tony as half-time research assistant to implement an isozyme lab. In so many ways, this was a opportune moment to arrive in Davis and share the excitement and friendship of the Allard lab (particularly Drs. M. T. Clegg, S. K. Jain, D. R. Marshall, and B. S. Weir). The scientific collaborations begun at Davis continued in projects for several decades and led later to sabbatical visits at Stanford University and the University of California, Riverside. UC Davis was thus a watershed in Tony’s science and life including marriage. With the completion of his PhD in 1969, Tony was appointed as a lecturer in Biology at the University of York, England. A seminar by Professor Warren Ewens in Leeds on the sampling theory of neutral genes had a lasting influence on Tony’s research. After three years, Tony returned to Australia to CSIRO Plant Industry as a research scientist in Canberra in 1972. There, two sons, Laurence and Christopher, were born. At CSIRO, Tony collaborated with Dr. Don Marshall, who had preceded him from UC Davis, and with Dr. Bruce Weir then at Massey, New Zealand. One early project was on the charge-state model of electrophoretic variation, from which a number of experiences flowed. One experience was to have their joint work scooped by Drs. T. Ohta and M. Kimura. On another occasion, a manuscript by Tony and Don was being subjected to the internal CSIRO editorial process and was sent by the panel for review to Professor P. A. P. Moran at Australian National University. Professor Moran was intrigued by the problem and concerned about some aspects of the existence and convergence properties of the distribution. He not only submitted his review, but more important also phoned Tony after hours to discuss this paper. This led Professor Moran to write a series of theoretical papers, and this thinking was referred to by Dr. J. F. C. Kingman in the history of coalescent that he wrote for Genetics in 2000. More influential than the existence of wandering distributions and the electrophoretic profile was Tony’s work on sampling strategies for plant genetic resources. Sir Otto Frankel had challenged Tony and Don

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to think critically on this subject. Later he promoted their strategy in international meetings and used it to challenge conventional collecting practice, particularly when he thought sampling was excessive. In the field, such theoretical strategies are but a guide, requiring adjustment to reality. This is particularly so for sampling the diversity of wild relatives, where one is deliberately seeking populations in diverse habitats and of greatly varying size. Along with the excitement of discovering variation new to science in its native setting came the experience of diverse human situations. Tony’s first real germplasm collecting mission was an object lesson in adjusting theory to field reality. This was a frenetic mission to Iran with Israeli professors Dani Zohary and Eibi Nevo. The trip went from Mehran near the border with Iraq, across the Zagros Mountains and the southern Caspian shores to Gonbad-e-Qabus, just two years before the 1979 revolution. With portraits of the shah’s family in every hotel room, the future course of events was not evident. At the hotel in Andimesk, the grim faces of the hotel staff were unforgettable as they examined the scientists’ passports. Although the target of the trip was wild cereals, particularly wild barley, the diversity being grown by farmers in the many barley fields was inspiring. This led to a sampling deliberately aimed at testing the allozyme diversity and genetic structure of these landraces, particularly to see whether the richness of diversity so apparent to the eye was just a mixture of a few genotypes. That research ultimately led to Tony’s principal commitment as Honorary Research Fellow with the International Plant Genetic Resources Institute in their project on the significance of crop genetic diversity still present on farms in traditional agroecosystems (with Drs. Toby Hodgkin and Devra Jarvis). The research focus was to develop a scientific basis of the use and conservation in situ of this diversity. When are germplasm collections large enough? This question came to Sir Otto with force after a visit to the excellent large world rice collection developed by Dr. T. T. Chang at the International Rice Research Institute. Few collections would have the resources to attain the size and organization of that for rice. Furthermore the human impulse to collect beyond need would lessen the utility of many collections. Sir Otto therefore proposed that core collections should be set up and called on Tony to read the draft. It was stunning to consider that Otto Frankel, renowned internationally as a champion of crisis germplasm collecting, should now advocate the slashing of collections by an order of magnitude. After much debate, a compromise concept of the ‘‘reserve’’ collection emerged. But for years thereafter Sir Otto often said, ‘‘Tony, I have you to blame for the reserve collection,’’ feeling

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that the reserve concept had dulled his original challenge to gene-bank managers to prune their holdings.

II. RESEARCH ACCOMPLISHMENTS A. Conservation Genetics Sir Otto Frankel was a member of the FAO Expert Panel on Plant Exploration and Introduction, and was preparing for the 1973 FAO/IBP Technical Conference on Crop Plant Genetic Resources in Rome. He felt that previous papers on plant collecting were strongly biased toward the practical details of collecting expeditions and that little emphasis had been given to the science of plant exploration. The original paper presented at the technical conference by Drs. Don Marshall and Tony Brown entitled ‘‘Optimum Sampling Strategies in Genetic Conservation’’ (subsequently published in the book Crop Genetic Resources for Today and Tomorrow edited by O. H. Frankel and J. G. Hawkes) was controversial but has since been widely accepted and expanded to cover other issues, such as sampling in biological control programs. Tony followed his early work on sampling strategies by extensive work on developing the concept of core collections. This concept, first introduced in 1984, was to facilitate the use of genetic resources in the major crops. By the mid-1980s it was felt that many collections, especially in the major crops, had grown so large that their mere size was likely to deter their extensive use by individual scientists, breeders, or students, except for a few characters that could be readily and rapidly discerned on single plants. It was proposed that giving priority in evaluation to a smaller number of accessions would facilitate greater use of germplasm collections, particularly for a range of characters. In a series of papers over the last 20 years, Tony has provided much of the underlying scientific rationale for the establishment and use of core collections. When first introduced, the core collection concept, because it challenged accepted dogma, was controversial, but it now has become widely applied in practice.

B. Plant Mating Systems and Population Structure A key theme spanning Tony’s research career has been the measurement of plant mating systems and their impact on population variation and structure, and the implications of these differences in terms of evolution and the collection and conservation of genetic diversity.

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One of his earliest papers with Professor Allard, which was based on his PhD research, reported the use of isozyme polymorphisms to estimate mating system parameters in open-pollinated maize populations. Over his career Tony developed procedures not only for the estimation of mating system parameters in both predominantly inbreeding and outbreeding populations but for also for apomictic species. Tony also worked with a wide range of colleagues in applying these techniques in species as diverse as Eucalyptus, Lupinus, wild Hordeum, and a number of colonizing weed species (with Drs. Jeremy Burdon and Spencer Barrett). Tony’s work on population structure was focused on genetic polymorphism, heterozygosity, multilocus associations, and population differentiation. A theoretical project with Dr. Marc Feldman, which is enjoying renewed attention with the burgeoning DNA sequence data, dealt with the measuring and testing of multilocus associations. Another example is the analysis of published isozyme data undertaken with Dr. Dan Schoen, which showed that not only do inbreeding and outbreeding species differ in overall levels of genetic diversity, but they also differ in the amount of among population variance of gene diversity. Inbreeding species exhibited much greater variation in how their populations are structured than do the populations of individual outbreeding species. We have to be clear that the comparison is the variability between the populations of one species; that is, populations 1 and 2 of species A, not population 1 of species A with population 1 of species B. Hordeum Research. Another major research theme of Tony’s was the genetic structure of natural populations of the wild progenitor (Hordeum spontaneum) of cultivated barley and the utilization of the variation in the wild species in barley improvement. His interest in wild barley was sparked in part by his experience at Davis and because of an invitation by Professor Nevo in 1976 to spend a six-month sabbatical in the Institute of Evolution, University of Haifa, in Israel. This opened an opportunity for Tony to collect wild barleys in the field and use enzyme markers to understand their genetic structure. The finding of extensive isozyme variation in wild barley raised the issue as to whether this indicated equally high levels of potentially useful variation for barley improvement. Screening of accessions collected in Israel and Iran revealed extensive resistance to barley scald and a range of other diseases and quality traits. Tony developed advanced backcross lines carrying scald resistance from the wild species, which proved to be a useful tool in investigating the genetics of seedling resistance and the tagging and pyramiding of resistance genes.

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This work led to the release of a commercial cultivar (Tantangara) carrying a known scald resistance gene from wild barley. An allied project, also conducted with Dr. Dave Garvin, was the use of molecular markers in breeding adapted proanthocyanidin-free barley. Glycine Research. The legume genus Glycine includes G. max (soybean) and its wild progenitor, G. soja. These annual species are native to northern Asia and so would seem to have little or nothing to do with Australia. Yet, surprisingly, their closest relatives, and the only other members of the genus Glycine, are native to Australia. This group of wild perennial species, Glycine subgenus Glycine, represents the tertiary gene pool for the soybean and is thus of potential economic importance. Collecting and characterizing these perennials has been a major focus of Tony’s work. The potential of this uniquely Australian resource was recognized at CSIRO by Dr. Don Marshall, working initially with Paul Broue` and Jim Grace. Subsequent staff changes led to Tony taking over the program in 1982. At that time there were fewer than 10 species recognized in subgenus Glycine, but that has changed dramatically. In 1982, the International Board of Plant Genetic Resources (IBPGR) held a workshop on soybean genetic resources at Urbana, Illinois, where Tony met those who were already, or would become, among the key figures in soybean diversity research, including Drs. Theodore Hymowitz, Reid Palmer, Randy Nelson, Christine Newell, and Duncan Vaughn. At the time of this workshop, papers on crosses between soybean and perennial Glycine species by the Hymowitz group and by the CSIRO group (Broue` and Marshall) were in draft, and there was tremendous excitement about the potential of the perennials for plant breeding, particularly as sources of drought- and disease-resistance genes. Tony came to the workshop with rough maps of the Australian Glycine distributions, based on existing herbarium records. This led to proposals for germplasm collecting trips to various poorly explored regions of Australia. As a result, IBPGR funded field trips conducted by Tony to north Queensland (1983), the Kimberley district, Western Australia, and Northern Territory (1984), south Queensland (1985), and eastern Victoria (1985). The first two of these trips were co-led by another leading figure in perennial Glycine research, Dr. Theodore Hymowitz (University of Illinois), with whom Tony led two later trips funded by the U.S. Department of Agriculture, to Western Australia (1993) and to the central arid zone of the continent (1996). As a result of these trips, the number of accessions in the CSIRO perennial soybean germplasm collection grew from a few hundred in 1982 to over 2,000 at present.

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It was through Tony’s role as curator of the perennial Glycine seed collection that he began a longtime collaboration with Dr. Jeff Doyle (Cornell University) and his wife, Jane Doyle, when Dr. Doyle contacted CSIRO requesting seed for systematic studies in 1982. Tony’s detailed knowledge of Glycine has guided their collaboration, which has produced numerous papers on the molecular phylogenetics of the subgenus. The chloroplast phylogeny of Glycine corroborated the existence of the genome groups that were based on cytological data amassed by the CSIRO and the University of Illinois groups, and offered the first hypothesis of relationships among these groups of species. The availability of a phylogeny based on defined molecular markers shared among all species also allowed the affinities of newly described species to be determined without recourse to the painstaking studies of chromosome pairing in difficult-to-produce artificial F1 hybrids conducted by Tony and colleagues in the 1980s and by the group at Illinois. Phylogenies based on nuclear markers subsequently showed some incongruence with chloroplast sequences, and some relationships in Glycine remain unresolved. Despite these limitations, molecular systematic approaches have replaced artificial hybridization as the standard method for categorizing new species in the subgenus. The field collections made these studies possible and also enabled traditional taxonomic studies of the genus by other workers, notably Drs. Mary Tindale, Lyn Craven, and Bernard Pfeil. Thanks to all of their efforts and the ongoing work of Dr. Hymowitz and colleagues, the number of species in the subgenus has grown to 25, including 3 new species formally named in 2006. Of the 25, 11 were first found in the field trips led by Tony. Additional species remain to be described in this diverse group. Diploid (2n ¼ 38, 40) members of subgenus Glycine are confined almost exclusively to the Australian continent; one species makes it across the Torresian Strait to the tip of Papua New Guinea. But the subgenus also contains tetraploids (2n ¼ 78–80), some of which are distributed throughout the Pacific, in Timor, the Philippines, New Caledonia, Vanuatu, Taiwan, and the Ryukya Islands. Tony’s work, along with parallel studies by Dr. Hymowitz and his group, has been instrumental in unraveling the origins of these species, pinpointing their diploid progenitors, and elucidating processes such as multiple origins and lineage recombination, by which these allopolyploids have become genetically diverse and geographically widespread. With this wealth of data, and the resources developed by years of collecting, subgenus Glycine has become a significant model for studying the process of allopolyploid evolution in plants. Tony’s dynamic collaborations on Glycine are a stellar example of cooperation

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to solve biological problems and are a tribute to his congenial and collegial personality. In the two and a half decades since Tony assumed responsibility for CSIRO’s perennial Glycine research, he published papers on a number of other topics, including disease resistance, seed size, floral biology, several on population genetics, and the distribution of calcium oxalate crystals in Glycine and allies. And through all of this, Tony drew on his Glycine work as a complement to his studies on Hordeum and other plants, to refine and illustrate his views on germplasm contributions, the area to which he has dedicated himself for many years.

III. THE MAN Dr. Brown has a passion for conservation genetics, from his formative years with sugarcane to the present with Glycine species. His admiration for Otto Frankel, his diligent research at CSIRO, and his affiliation with the International Plant Genetic Resources Institute (now Bioversity International), Rome, Italy (1982–present), are evident in his many contributions. CSIRO Plant Industry as his home base has been an excellent and supportive research environment, where Tony worked jointly with many outstanding colleagues, including Drs. Jeremy Burdon (who is the current chief of Plant Industry), Curt Brubaker, Andrew Young, Jake Jacobsen, and several others. Indeed, these characteristics of Plant Industry owe much to Sir Otto who, as a former chief of division, instilled a vision of excellence in plant research. The discovery of taxa new to science is the unique reward for the collector of wild species related to important crops. Each of his many trips had memorable incidents for Tony, and three are mentioned here. If you happened to be one of the few vehicles driving the remote dirt Peninsula ‘‘highway’’ in Cape York, north Queensland, in July of 1983, you may have seen three collectors (Ted Hymowitz from Illinois and Jim Grace and Tony from CSIRO) sprawled on the lawn outside the Lakeland pub below the billboard saying ‘‘Ice Cold Beer.’’ This was no early knock off; they actually were sampling rare, tiny Glycine tomentella plants. The billboard had nothing to do with site selection; a collector must check all habitats. The roadside pub, a lone building in the rural landscape, was a haven for the thirsty traveler, and it surrounds a haven for wild plants that grazing animals would otherwise decimate. Thus, sampling strategies for germplasm collection adapt to reality. Day’s end of another trip found Tony with U.S. soybean breeder Dr. Bill Kenworthy, graduate student Michael Doyle, and Jim Grace in

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the Carnarvon Gorge of central Queensland, which proved a special site, rich in diverse new taxa. Excitement died, however, when halfway along the 150-km return trip, it became clear there was insufficient fuel to reach home at Injune. There followed a long silent drive, meeting no other vehicles. Despite eking out the last drop of fuel, the vehicle slowed to a stop 15 km north of their destination. Tony and Jim remained with the vehicle; Bill and Michael chose to jog and walk to town for help, guided by moonlight and the smell of road-killed cattle and kangaroos. During the long wait at the vehicle, the silent darkness was broken by another vehicle, the first sighted since Carnarvon and, luckily, approaching the road to town, from the property right where the vehicle had stopped. When they apprised the driver of the pickup of their situation, he pointed to the rifle hung above his rear window and replied in a rural Texas accent, ‘‘Just as well you told me. If I’d been forced to stop on the road in the dark by two desperados on foot, looking for a ride to town, I’d answer with this.’’ Meeting the wildlife is a feature of any field trip in Australia. A trip to collect wild Australian Gossypium species, and to evaluate the risks to them of GM cotton, with botanist Professor Herbert Hurka from Osnabrueck, Germany, brought them to a remote Corona farm 70 km north of Broken Hill, western NSW. Herbert was intrigued by the caged talking sulphur-crested cockatoo. The farmer’s wife had warned them that the bird had lived in a hotel in the ‘‘silver city’’ but was banished because of bad language. Clearly the garrulous bird enjoyed the attention of the team of rare visitors, and Herbert lingered to converse with it while the CSIRO team sampled. When he turned to leave, the bird had a fail-safe method to retrieve attention. To the visiting professor, it screeched ‘‘A***hole’’—a fully effective way to grab Herbert’s notice. Sir Otto Frankel was one of the major influences of Tony’s science and life. His unyielding insistence on high standards and exactness led to many legendary stories. Memorable for Tony was a Christmas Eve lunch at which Tony hosted Sir Otto and Lady Margaret, along with Professor Herbert and Ute Hurka and family members. At one point, Tony introduced a wine he was particularly enjoying, and asked who would like some of this excellent Orlando chardonnay, Otto’s response was immediate and emphatic: ‘‘That wine is good, but is certainly NOT excellent!!’’ Silence fell; then he asked, ‘‘Which vintage?’’ Stunned, Tony checked the label and replied, ‘‘1988.’’ Back came the riposte: ‘‘1987 is better!!’’ Those quips have often proved useful, not only when recalling Otto’s outspokenness. Elements of Tony’s migratory career and strategic sampling interests are reflected in the lives of his two sons. Currently, Laurence is a lecturer

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in migration history in the Department of History, University of Manchester, UK; and Chris is an investment banker with the Mergers and Acquisitions Section of UBS, New York, USA. Tony has a new younger family of three stepchildren who are themselves embarking on diverse careers.

IV. HONORS AND AWARDS Dr. Brown has been extensively recognized for his contributions and achievements to conservation genetics. To further broaden his expertise, Tony has been a visiting professor at Stanford University and the University of California, Riverside, visiting research fellow at Haifa University, Israel, and a visiting scientist at the Universitaet Osnabrueck, Germany. Tony has excelled in his editorial duties for the journals Genetics, Molecular Biology and Evolution, and Conservation Genetics as well as serving as editor or coeditor of 10 books, and conference and symposia proceedings. Of the eight plant collecting missions, Tony has been leader or coleader of six in Australia, one in Israel, and one in Iran. Tony has been the International Plant Genetic Resources Institute (IPGRI) technical advisor and on the organizing committee of 18 international workshops in 10 different countries. Perhaps the most rewarding honor was the award as Honorary Research Fellow by the IPGRI, Rome, Italy. The initial award was in 1997 and Tony has been reappointed three times, most recently with Bioversity International, IPGRI’s new name.

ACKNOWLEDGMENT The authors thank Dr. Don Marshall of Plant Breeding Solutions Pty. Ltd., Hamilton, NSW, Australia, for his contributions to the text and for his critical review of this chapter.

SELECTED PUBLICATIONS OF ANTHONY H. D. BROWN Brown, A.H.D., J. Daniels, and B.D.H. Latter. 1968. Quantitative genetics of sugarcane. I. Analysis of variation in a commercial hybrid sugarcane population. Theor. Appl. Genet. 38:361–369. Brown, A.H.D. and R.W. Allard. 1969. Inheritance of isozyme differences among the inbred parents of a reciprocal recurrent selection population of maize. Crop Sci. 9:72–75.

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Brown, A.H.D. 1970. The estimation of Wright’s fixation index from genotypic frequencies. Genetica 41:399–406. Brown, A.H.D. and R.W. Allard. 1970. Estimation of the mating system in open–pollinated maize populations using isozyme polymorphisms. Genetics 66:133–145. Brown, A.H.D. 1971. Isozyme variation under selection in Zea mays. Nature 232:570. Brown, A.H.D. and R.W. Allard 1971. Effect of reciprocal recurrent selection for yield on isozyme polymorphisms in maize (Zea mays L.). Crop Sci. 11:888–893. Marshall, D.R. and A.H.D. Brown. 1973. Stability of performance of mixtures and multilines. Euphytica 22:405–412. Brown, A.H.D., D.R. Marshall, and L. Albrecht. 1974. The maintenance of alcohol dehydrogenase polymorphism in Bromus mollis L. Aust. J. Biol. Sci. 27:545–559. Marshall, D.R. and A.H.D. Brown. 1974. Estimation of the level of apomixis in plant populations. Hered. 32:321–333. Brown, A.H.D., A.C. Matheson, and K.G. Eldridge. 1975. Estimation of the mating system of Eucalyptus obliqua L. Herit using allozyme polymorphisms. Aust. J. Bot. 23:931–949. Brown, A.H.D. 1975. Efficient experimental designs for the estimation of genetic parameters in plant populations. Biometrics 31:145–160. Brown, A.H.D. 1975. Sample sizes required to detect linkage disequilibrium between two or three loci. Theor. Pop. Biol. 8:184–210. Brown, A.H.D., D.R. Marshall, and L. Albrecht. 1975. Profiles of electrophoretic alleles in natural populations. Genet. Res. Camb. 25:137–143. Brown, A.H.D., D.R. Marshall, and B.S. Weir. 1975. Population differentiation under the charge state model. Genetics 81:739–748. Marshall, D.R. and A.H.D. Brown. 1975. The charge state model of protein polymorphism in natural populations. J. Molec. Evol. 6:149–163. Marshall, D.R. and A.H.D. Brown. 1975. Optimum sampling strategies in genetic conservation. pp. 53–80. In: O.H. Frankel and J.G. Hawkes (eds.), I.B. P.2. Crop Genetic Resources for Today and Tomorrow. Cambridge Univ. Press, Cambridge. Brown, A.H.D., D.R. Marshall, and J. Munday. 1976. The adaptedness of variants at an alcohol dehydrogenase locus in Bromus mollis L. (Soft Bromegrass). Aust. J. Biol. Sci. 29:389–396. Weir, B.S., A.H.D. Brown, and D.R. Marshall. 1976. Testing for selective neutrality of electrophoretically detectable protein polymorphisms. Genetics 84:639–659. Brown, A.H.D., E. Nevo, and D. Zohary. 1977. Association of alleles at esterase loci in wild barley Hordeum spontaneum. Nature 268:430–431. Brown, A.H.D. 1978. Isozymes, plant population genetic structure and genetic conservation. Theor. Appl. Genet. 52:145–157. Brown, A.H.D., E. Nevo, D. Zohary, and O. Dagan. 1978. Genetic variation in natural populations of wild barley (Hordeum spontaneum). Genetica 49:97–108. Brown, A.H.D., D. Zohary, and E. Nevo. 1978. Outcrossing rates and heterozygosity in natural populations of Hordeum spontaneum Koch in Israel. Hered. 41:49–62. Brown, A.H.D. 1979. Enzyme polymorphism in plant population. Theor. Pop. Biol. 15:1–42. Nevo, E., D. Zohary, A.H.D. Brown, and M. Haber. 1979. Genetic diversity and environmental associations of wild barley, Hordeum spontaneum, in Israel. Evolution 33:815–833. Doll, H. and A.H.D. Brown. 1979. Hordein variation in wild (Hordeum spontaneum) and cultivated (H. vulgare) barley. Can. J. Genet. Cytol. 21:391–404. Brown, A.H.D. 1980. Genetic basis of alcohol dehydrogenase polymorphism in Hordeum spontaneum. J. Hered. 70:127–128.

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Brown, A.H.D. and L. Albrecht. 1980. Variable outcrossing and the genetic structure and predominantly self-pollinated species. J. Theor. Biol. 82:591–606. Brown, A.H.D., M.W. Feldman, and E. Nevo. 1980. Multilocus structure of natural populations of Hordeum spontaneum. Genetics 96:523–536. Corrigendum May 1981, p. 238A. Green, A.G., A.H.D. Brown, and R.N. Oram. 1980. Determination of outcrossing rate in a breeding population of Lupinus albus L. (White Lupin). Z. Pflanzenzuchtg. 84:181–191. Brown, A.H.D. and M.W. Feldman. 1981. Population structure of multilocus associations. Proc. Natl. Acad. Sci. U.S. 78:5913–5916. Brown, A.H.D. and D.R. Marshall. 1981. Evolutionary changes accompanying colonization in plants. pp. 351–363. In: G.G.E. Scudder and J.L. Reveal (eds.), Evolution Today, Proc. Second Int. Congr. Syst. and Evol. Biol. Univ. British Columbia, Vancouver. Hunt Institute for Botanical Documentation, Pittsburgh. Marshall, D.R. and A.H.D. Brown. 1981. The evolution of apomixis. Hered. 47:1–15. Marshall, D.R. and A.H.D. Brown. 1981. Wheat genetic resources. pp. 21–40. In: W. J. Peacock and L.T. Evans (eds.), Wheat Science, Today and Tomorrow. Cambridge Univ. Press, Cambridge. Brown, A.H.D. and J.V. Jacobsen. 1982. Genetic basis and natural variation of alpha– amylase isozymes in barley. Genet. Res. Camb. 40:315–324. Brown, A.H.D. and J. Munday. 1982. Population genetic structure and optimal sampling of land races of barley from Iran. Genetica 58:85–96. Erratum 60:237. Nevo, E., E. Golenberg, A. Beiles, A.H.D. Brown, and D. Zohary. 1982. Genetic diversity of environmental associations of wild wheat, Triticum dicoccoides in Israel. Theor. Appl. Genet. 62:241–254. Brown, A.H.D. 1983. Barley, pp. 57–77. In: S.D. Tanksley, and T.J. Orton (eds.), Isozymes in plant genetics and breeding, Part B. Elsevier, Amsterdam. Brown, A.H.D. and J.J. Burdon. 1983. Multilocus diversity in an outbreeding weed, Echium plantagineum L. Aust. J. Biol. Sci. 36:503–509. Brown, A.H.D. and M.T. Clegg. 1983. Analysis of variation in related DNA sequences. pp. 107– 132. In: B.S. Weir (ed.), Statistical analysis of DNA sequence data. Marcel Dekker, New York. Brown, A.H.D. and B.S. Weir. 1983. Measuring genetic variability in plant populations. pp. 219–239. In: S.D. Tanksley. and T.J. Orton (eds.), Isozymes in plant genetics and breeding, Part A. Elsevier, Amsterdam. Burdon, J.J., D.R. Marshall, and A.H.D. Brown. 1983. Demographic and genetic changes in populations of Echium plantagineum L. J. Ecology 71:667–679. Brown, A.H.D. 1984. Multilocus organization of plant populations. pp. 159–169. In: K. Wohrmann and V. Loeschcke (eds.), Population biology and evolution. Springer Verlag, Berlin. Clegg, M.T., A.H.D. Brown, and P.R. Whitfeld. 1984. Chloroplast DNA diversity in wild and cultivated barley: Implications for genetic conservation. Genet. Res. Camb. 43: 339–343. Hanson, A.D. and A.H.D. Brown. 1984. Three alcohol dehydrogenase genes in wild and cultivated barley: characterization of the products of variant alleles. Biochem. Genet. 22:495–515. Grant, J.E., A.H.D. Brown, and J.P. Grace. 1984. Cytological and isozyme diversity in Glycine tomentella Hayata (Leguminosae). Aust. J. Bot. 32:665–677. Grant, J.E., J.P. Grace, A.H.D. Brown, and E. Putievsky. 1984. Interspecific hybridization in Glycine subgenus Glycine Willd. (Leguminosae). Aust. J. Bot. 32:655–663. Schroeder, H.E. and A.H.D. Brown. 1984. Inheritance of legumin and albumin contents in a cross between round and wrinkled peas. Theor. Appl. Genet. 68:101–107. Zurawski, G., M.T. Clegg, and A.H.D. Brown. 1984. The nature of nucleotide sequence divergence between barley and maize chloroplast DNA. Genetics 106:735–749.

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Brown, A.H.D., J.E. Grant, J.J. Burdon, J.P. Grace, and R. Pullen. 1985. Collection and utilization of wild perennial Glycine. pp. 345–352. In: R. Shibles (ed.), Proc. World Soybean Research Conference III. Westview Press, Boulder, Colorado. Doyle, M.J. and A.H.D. Brown. 1985. Numerical analysis of isozyme variation in Glycine tomentella. Biochem. Syst. Ecol. 13:413–419. Brown, A.H.D., J.E. Grant, and R. Pullen. 1986. Outcrossing and paternity in Glycine argyrea by paired fruit analysis. Biol. J. Linn. Soc. 29:283–294. Burdon, J.J. and A.H.D. Brown. 1986. Population genetics of Echium plantagineum L.—a target weed for biological control. Aust. J. Biol. Sci. 39:369–378. Doyle, M.J., J.E. Grant, and A.H.D. Brown. 1986. Reproductive isolation between isozyme groups of Glycine tomentella (Leguminosae), and spontaneous doubling in their hybrids. Aust. J. Bot. 34:523–535. Grant, J.E., R. Pullen, A.H.D. Brown, J.P. Grace, and P.M. Gresshof. 1986. Cytogenetic affinity between the new species Glycine argyrea and its congeners. J. Hered. 77: 423–426. Brown, A.H.D. and J.J. Burdon. 1987. Mating systems and colonizing success in plants. pp. 115–131. In: A.J. Gray, M.J. Crawley, and P.J. Edwards (eds.), Colonization, succession and stability. 26th Symposium of British Ecol. Soc. Blackwell Scientific, Oxford. Henry, R.J. and A.H.D. Brown. 1987. Variation in the carbohydrate composition of wild barley (Hordeum spontaneum) grain. Z. Pflanzenzu¨chtung 98:97–103. Hoffman, N.E., D. Hondred, A.D. Hanson, and A.H.D. Brown. 1988. Lactate dehydrogenase isozymes in barley: Polymorphism and genetic basis. J. Hered. 79:110–114. Brown, A.H.D., J. Munday, and R.N. Oram. 1988. Use of isozyme-marked segments from wild barley (Hordeum spontaneum) in barley breeding. Plant Breed. 100:280–288. Brown, A.H.D. 1989. The case for core collections. pp. 136–156. In: A.H.D. Brown, O.H. Frankel, D.R. Marshall, and T. Williams (eds.), The use of plant genetic resources. Cambridge Univ. Press, Cambridge. Brown, A.H.D. 1989. Core collections: A practical approach to genetic resources management. Genome 31:818–824. Brown, A.H.D. 1989. Genetic characterization of plant mating systems. pp. 145–162. In: A.H.D. Brown, M.T. Clegg, A.L. Kahler, and B.S. Weir (eds.), Plant population genetics, breeding and genetic resources. Sinaeuer Associates, Sunderland, Massachusetts. Brown, A.H.D., J.J. Burdon, and A.M. Jarosz. 1989. Isozyme analysis of plant mating systems. pp. 73–86. In: D. Soltis and P. Soltis (eds.), Isozymes in plant biology. Dioscorides Press, Portland, Oregon. Brown, A.H.D., G.J. Lawrence, M. Jenkin, J. Douglass, and E. Gregory. 1989. Linkage drag in backcross breeding. J. Hered. 80:234–239. Doyle, J.J. and A.H.D. Brown. 1989. 5S nuclear ribosomal gene variation in the Glycine tomentella polyploid complex. Syst. Bot. 14:398–407. Hurka, H., S. Freunder, A.H.D. Brown, and U. Plantholt. 1989. Aspartate amino transferase isozymes in the genus Capsella (Brassicaceae): Subcellular location, gene duplication and polymorphism. Biochem. Genetics 27:77–90. Kenworthy, W.J., A.H.D. Brown, and G.A. Thibou. 1989. Variation in flowering response to photoperiod in perennial Glycine species. Crop Sci. 29:678–682. Brown, A.H.D. 1990. The role of isozyme studies in molecular systematics. Aust. Syst. Bot. 3:39–46. Brown, A.H.D., J.J. Burdon and J.P. Grace. 1990. Genetic structure of Glycine canescens, a perennial relative of soybean. Theor. Appl. Genet. 79:729–736. Doyle, J.J., J.L. Doyle, and A.H.D. Brown. 1990. Analysis of a polyploid complex in Glycine with chloroplast and nuclear DNA. Aust. Syst. Bot. 3:125–136.

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Doyle, J.J., J.L. Doyle, and A.H.D. Brown. 1990. A chloroplast DNA phylogeny of the wild perennial relatives of soybean (Glycine subgenus Glycine): Congruence with morphological and crossing groups. Evolution 44:371–389. Doyle, J.J., J.L. Doyle, and A.H.D. Brown. 1990. Chloroplast DNA phylogenetic affinities of newly described species in Glycine (Leguminosae: Phaseoleae). Syst. Bot. 15:466–471. Doyle, J.J., J.L. Doyle, and A.H.D. Brown. 1990. Chloroplast DNA polymorphism and phylogeny in the B genome of Glycine subgenus Glycine (Leguminosae). Amer. J. Botany 77:772–782. Doyle, J.J., J.L. Doyle, A.H.D. Brown, and J.P. Grace. 1990. Multiple origins of polyploids in the Glycine tabacina complex inferred from chloroplast DNA polymorphism. Proc. Natl. Acad. Sci. USA 87:714–717. Doyle, J.J., J.L. Doyle, J.P. Grace, and A.H.D. Brown. 1990. Reproductively isolated polyploid races of Glycine tabacina (Leguminosae) had different chloroplast genome donors. Syst. Bot. 15:173–181. Feuerstein, U., A.H.D. Brown, and J.J. Burdon. 1990. Linkage of rust resistance genes from wild barley (Hordeum spontaneum) with isozyme markers. Plant Breed. 104:318–324. Schoen, D.J. and A.H.D. Brown. 1991. Intraspecific variation in population gene diversity and effective population size correlates with the mating system in plants. Proc. Natl. Acad. Sci. USA 88:4494–4497. Abbott, D.C., J.J. Burdon, A.M. Jarosz, A.H.D. Brown, W.J. Muller, and B.J. Read. 1991. The relationship between seedling infection types and field reactions to leaf scald in Clipper barley backcross lines. Aust. J. Agric. Res. 42:801–809. Brown, A.H.D. and J.D. Briggs. 1991. Sampling strategies for genetic variation in ex situ collections of endangered plant species. pp. 99–119. In: D.A. Falk and K.E. Holsinger (eds.), Genetics and Conservation of Rare Plants. Oxford Univ. Press, Oxford. Lagudah, E.S., R. Appels, A.H.D. Brown, and D. McNeil. 1991. The molecular-genetic analysis of Triticum tauschii, the D-genome donor to hexaploid wheat. Genome 34: 375–386. MacLeod, L.C., R.C.M. Lance, and A.H.D. Brown. 1991. Chromosomal mapping of the Glb 1 locus encoding (1!3), (1!4)–ß–D–glucan 4–glucanohydrolase EI in barley. J. Cereal Sci. 13:291–298. Schoen, D.J. and A.H.D. Brown. 1991. Whole and part-flower self-pollination in Glycine clandestina and G. argyrea and the evolution of autogamy. Evolution 45:1651–1664. Abbott, D.C., A.H.D. Brown, and J.J. Burdon. 1992. Genes for scald resistance from wild barley (Hordeum vulgare ssp spontaneum) and their linkage to isozyme markers. Euphytica 61:225–231. Brown, A.H.D. 1992. Genetic variation and resources in cultivated barley and wild Hordeum. Barley Genetics 6:669–682. Brown, A.H.D. 1992. Human impact on plant gene pools and sampling for their conservation. Oikos. 63:109–118. Schoen, D.J., J.J. Burdon, and A.H.D. Brown. 1992. Resistance of Glycine tomentella to soybean leaf rust Phakopsora pachyrhizi in relation to ploidy level and geographic distribution. Theor. Appl. Genet. 83:827–832. Schoen, D.J. and A.H.D. Brown. 1993. Conservation of allelic richness in wild crop relatives is aided by assessment of genetic markers. Proc. Natl. Acad. Sci. USA 90:10623–10627. Brown, A.H.D. and D.J. Schoen. 1994. A revised measure of association of gene diversity values. Hereditas 120:77–79. Burdon, J.J., D.C. Abbott, A.H.D. Brown, and J.S. Brown. 1994. Genetic structure of the scald pathogen (Rhynchosporium secalis) in South East Australia: Implications for control strategies. Aust. J. Agric. Res. 45:1445–1454.

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Guerin, J.R., R.C.M. Lance, A.H.D. Brown, and D.C. Abbott. 1994. Mapping of malt endopeptidase, diaphorase and esterase loci on barley chromosome 3L. Plant Breed. 112:279–284. Prober, S.M. and A.H.D. Brown. 1994. Conservation of the grassy white box woodlands. I. Population genetics and fragmentation of Eucalyptus albens Benth. Conservation Biol. 8:1003–1013. Abbott, D.C., E.S. Lagudah, and A.H.D. Brown. 1995. Identification of RFLPs flanking a scald resistance gene on barley chromosome 6. J. Hered. 86:152–154. Brown, A.H.D. 1995. The core collection at the crossroads. pp. 3–19. In: T. Hodgkin, A.H.D. Brown, T.J.L. van Hintum, and E.A.V. Morales (eds.), Core collections of plant genetic resources. John Wiley, Chichester. Brown, A.H.D. and D.R. Marshall. 1995. A basic sampling strategy: Theory and practice. pp. 75–91. In: L. Guarino, V. Ramanatha Rao, and R. Reid (eds.), Collecting plant genetic diversity technical guidelines. CAB International, Wallingford. Frankel, O.H., A.H.D. Brown, and J.J. Burdon. 1995. The conservation of plant biodiversity. Cambridge Univ. Press, Cambridge. Brown, A.H.D., D.F. Garvin, J.J. Burdon, D.C. Abbott, and B.J. Read. 1996. The effect of combining scald resistance genes on disease levels, yield and quality traits in barley. Theor. Appl. Genet. 93:361–366. Young, A., G.T. Boyle, and A.H.D. Brown. 1996. The population genetic consequences of habitat fragmentation for plants. Trends in Ecology and Evolution 11:413–418. Young, A.G. and A.H.D. Brown. 1996. Comparative population genetic structure on the rare woodland shrub Daviesia suaveolens and its common congener D. mimosoides. Conservation Biol. 10:1220–1228. Brown, A.H.D., C.L. Brubaker, and J.P. Grace. 1997. The regeneration of germplasm samples: Wild versus cultivated species. Crop Sci. 37:7–13. Brown, A.H.D., C.L. Brubaker, and M.J. Kilby. 1997. Assessing the risk of cotton transgene escape into wild Australian Gossypium species. pp. 83–94. In: G.D. McLean, P.M. Waterhouse, G. Evans, and M.I. Gibbs (eds.), The commercialisation of transgenic crops: Risk, benefit and trade considerations. Bureau of Resource Sciences, Kingston, ACT, Australia. Garvin, D.F., A.H.D. Brown, and J.J. Burdon. 1997. Inheritance and chromosome locations of novel scald resistance genes derived from Iranian and Turkish wild barleys. Theor. Appl. Genet. 94:1087–1091. Roulin, S., P. Xu, A.H.D. Brown, and G.B. Fincher. 1997. Expression of specific (1!3)-bGlucanase genes in leaves of near-isogenic resistant and susceptible barley lines infected with the leaf scald fungus (Rhynchosporium secalis). Phys. Mol. Plant Path. 50:245–261. Garvin, D.F., J.E. Miller-Garvin, E.A. Viccars, J.V. Jacobsen, and A.H.D. Brown. 1998. Identification of molecular markers linked to ant28, a mutation that eliminates proanthocyanidin in barley seeds. Crop Sci. 38:1250–1255. Prober, S.M., L.H. Spindler, and A.H.D. Brown. 1998. Conservation of the grassy white box woodlands: Effects of remnant population size on genetic diversity of the outcrossing, allotetraploid herb, Microseris lanceolata. Conservation Biol. 12:1279–1290. Young, A.G. and A.H.D. Brown. 1998. Comparative analysis of mating systems in the rare woodland shrub Daviesia suaveolens and its congener D. mimosoides. Hered. 80: 374–381. Brown, A.H.D. 1999. The genetic structure of crop landraces and the challenge to conserve them in situ on farms. pp. 29–48. In: S.B. Brush (ed.), Genes in the field: Conserving plant diversity on farms. Lewis Publishers, Boca Raton, FL. Brubaker, C.L., A.H.D. Brown, J. McD. McStewart, M.J. Kilby, and J.P. Grace. 1999. Production of fertile hybrid germplasm with diploid Australian Gossypium species for cotton improvement. Euphytica 108:199–213.

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Burdon, J.J.P.H. Thrall, and A.H.D. Brown. 1999. Resistance and virulence structure in two Linum marginale—Melampsora lini host-pathogen metapopulations with different mating systems. Evolution 53:704–716. Doyle, J.J., J.L. Doyle, and A.H.D. Brown. 1999. Incongruence in the diploid B-genome species complex of Glycine (Leguminosae) revisited: Histone H3-D alleles vs. chloroplast haplotypes. Molec. Biol. Evol. 16:354–362. Doyle, J.J., J.L. Doyle, and A.H.D. Brown. 1999. Origins, colonization, and lineage recombination in a widespread perennial soybean polyploid complex. Proc. Nat. Acad. Sci. USA 96:10741–10745. Marshall, D.R. and A.H.D. Brown. 1999. Sampling wild legume populations. pp. 78–89. In: S.J. Bennett and P.S. Cocks (eds), Genetic resources of Mediterranean pasture and forage legumes. Kluwer Acad. Press, Dordrecht. Young, A.G., A.H.D. Brown, and F.C. Zich. 1999. Genetic structure of fragmented populations of the endangered grassland daisy Rutidosis leptorrhynchoides. Conservation Biol. 13:256–265. Young, A.G. and A.H.D. Brown. 1999. Paternal bottlenecks in fragmented populations of the grassland daisy Rutidosis leptorrhynchoides. Genet. Res. 73:111–117. Abbott, D.C., J.J. Burdon, A.H.D. Brown, B.J. Read, and D. Bittisnich. 2000. The incidence of barley scald in cultivar mixtures. Aust. J. Agric. Res. 51:355–360. Brown, A.H.D. and C.L. Brubaker. 2000. Genetics and the conservation and use of Australian wild relatives of crops. Aust. J. Bot. 48:297–303. Brown, A.H.D. and C.M. Hardner. 2000. Sampling the gene pools of forest trees for ex situ conservation. pp. 185–196. In: A. Young, T. Boyle, and D. Boshier (eds.), Forest conservation genetics: Principles and practice. CSIRO, Melbourne, Australia. Brown, A.H.D. and A.G. Young. 2000. Genetic diversity in tetraploid populations of the endangered daisy Rutidosis leptorrhynchoides and implications for its conservation. Hered. 85:122–129. Doyle, J.J., J.L. Doyle, A.H.D. Brown, and B.L. Pfeil. 2000. Confirmation of shared and divergent genomes in the Glycine tabacina polyploid complex (Leguminosae) using histone H3-D sequences. Syst. Bot. 25:437–448. Garvin, D.F., A.H.D. Brown, H. Raman, and B.J. Read. 2000. Genetic mapping of the barley Rrs14 scald resistance gene with RFLP, isozyme and seed storage protein markers. Plant Breed. 119:193–196. Brown, A.H.D. and C.L. Brubaker. 2001. Indicators for sustainable management of plant genetic resources—how well are we doing? pp. 249–262. In: J.M.M. Engels, V. Ramanatha Rao, A.H.D. Brown, and M T. Jackson (eds.), Managing plant genetic diversity. CAB International, Wallingford, Oxon, UK. Lin, J.-Z., A.H.D. Brown, and M.T. Clegg. 2001. Heterogeneous geographic patterns of nucleotide sequence diversity between two alcohol dehydrogenase genes in wild barley (Hordeum vulgare ssp. spontaneum). Proc. Nat. Acad. Sci. USA 98:531–536. Teshome, A., A.H.D. Brown, and T. Hodgkin. 2001. Diversity in landraces of cereal and legume crops. Plant Breed. Rev. 21:221–261. Brown, A.H.D., J.L. Doyle, J.P. Grace, and J.J. Doyle. 2002. Molecular phylogenetic relationships within and among diploid races of Glycine tomentella (Leguminosae). Aust. Syst. Bot. 15:37–47. Bleeker, W., A. Franzke, K. Pollman, A.H.D. Brown, and H. Hurka. 2002. Phylogeny and biogeography of southern hemisphere high mountain Cardamine species (Brassicaceae). Aust. Syst. Bot. 15:575–581.

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Doyle, J.J., J.L. Doyle, A.H.D. Brown, and R.G. Palmer. 2002. Genomes, multiple origins, and lineage recombination in the Glycine tomentella (Leguminosae) polyploid complex: histone H3-D gene sequences. Evolution 56:1388–1402. Rauscher, J.T., J.J. Doyle, and A.H.D. Brown. 2002. Internal transcribed spacer repeat– specific primers and the analysis of hybridization in the Glycine tomentella (Leguminosae) polyploid complex. Molec. Ecol. 11:2691–2702. Brubaker, C.L. and A.H.D. Brown. 2003. The use of multiple alien chromosome addition aneuploids facilitates genetic linkage mapping of the Gossypium G genome. Genome 46:774–791. Genger, R.K., A.H.D. Brown, W. Knogge, K. Nesbitt, and J.J. Burdon. 2003. Development of SCAR markers linked to a scald resistance gene derived from wild barley. Euphytica 134:149–159. Genger, R.K., K.J. Williams, H. Raman, B.J. Read, H. Wallwork, J.J. Burdon, and A.H.D. Brown. 2003. Leaf scald resistance genes in Hordeum vulgare and Hordeum vulgare ssp. spontaneum: parallels between cultivated and wild barley. Aust. J. Agric. Res. 54:1335–1342. Doyle, J.J., J.L. Doyle, J.T. Rauscher, and A.H.D. Brown. 2003. Diploid and polyploidy reticulate evolution throughout the history of the perennial soybeans (Glycine subg. Glycine). New Phytologist 161:121–132. Murray, B.R., A.H.D. Brown, and J.P. Grace. 2003. Geographic gradients in seed size among and within perennial Australian Glycine species. Aust. J. Bot. 51:47–56. Rau, D., A.H.D. Brown, C.L. Brubaker, G. Attene, V. Balmas, E. Saba, and R. Papa. 2003. Population genetic structure of Pyrenophora teres Drechs., the causal agent of net blotch in Sardinian landraces of barley complex (Hordeum vulgare L.). Theor. Appl. Genet. 106:947–959. Doyle, J.J., J.L. Doyle, J.T. Rauscher, and A.H.D. Brown. 2004. Evolution of the perennial soybean polyploid (Glycine subgenus Glycine): A study of contrasts. Biol. J. Linnean Soc. 82:583–597. Joly, S., J.T. Rauscher, S.L. Sherman-Broyles, A.H.D. Brown, and J.J. Doyle. 2004. Evolution of the 18S-5.8S-26S nuclear ribosomal gene family and its expression in natural and artificial Glycine allopolyploids. Molec. Biol. Evol. 21:1409–1421. Murray, B.R., A.H.D. Brown, C.R. Dickman, and M.S. Crowther. 2004. Geographical gradients in seed mass in relation to climate.J. Biogeography 31:379–388. Rauscher, J.T., J.J. Doyle, and A.H.D. Brown. 2004. Multiple origins and nrDNA internal transcribed spacer homoeologue evolution in the Glycine tomentella (Leguminosae) allopolyploid complex. Genetics 166:987–998. Cervantes-Martinez, T., H.T. Horner, R.G. Palmer, T. Hymowitz, and A.H.D. Brown. 2005. Calcium oxalate crystal macropatterns in leaves of species from groups Glycine and Shuteria (Glycininae; Phaseoleae; Papilionoideae; Fabaceae). Can. J. Bot. 83:1410–1421 Genger, R.K., K. Nesbitt, A.H.D. Brown, D.C. Abbott, and J.J. Burdon. 2005. A novel barley scald resistance locus: Genetic mapping of the Rrs15 scald resistance gene derived from wild barley, Hordeum vulgare ssp. spontaneum. Plant Breed. 124: 137–141. Rau, D., F.J. Maier, R. Papa, A.H.D. Brown, V. Balmas, E. Saba, W. Shafer, and G. Attene, 2005. Isolation and characterization of the mating-type locus of the barley pathogen Pyrenophora teres frequencies of mating-type idiomorphs within and among fungal populations collected from barley landraces. Genome 48:855–869. Pfeil, B.E., L.A. Craven, A.H.D. Brown, B.G. Murray, and J.J. Doyle. 2006. Three new species of northern Australian Glycine (Fabaceae, Phaseolae) G-gracei, G. montis-dougle and G. syndetika. Aust. Syst. Bot. 19:245–258.

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Brown, A.H.D. and T. Hodgkin. 2007. Measuring, managing and maintaining crop genetic diversity on-farm. pp. 13–33. In: D. Jarvis, C. Paddoch, and D. Williams (eds.), Managing biodiversity in agricultural ecosystems. Columbia University Press, New York. Jarvis, D.I., A.H.D. Brown, V.I. Imbruce, J. Ochoa, M. Sadiki, E. Karamura, P. Trutmann, and M.R. Finckh. 2007. Managing crop disease in traditional agroecosystems: The benefits and hazards of genetic diversity. pp. 292–319. In: D. Jarvis, C. Paddoch, and D. Williams (eds.), Managing biodiversity in agricultural ecosystems. Columbia University Press, New York. Triono, T., M.D. Crisp, A.H.D, Brown, and J.G. West. 2007. A phylogency of Pouteria (Sapotaceae) from Malesia and Australasia. Aust. Syst. Bot. 20:107–118. Rau, D., G. Attene, A.H.D. Brown, L. Nanni, F.J. Maier, V. Balmas, E. Saba, W. Schaefer, and R. Papa. 2007. Phylogeny and evolution of mating-type genes from Pyrenophora teres, the causal agent of barley ‘‘net blotch’’ disease. Current Genetics 51:377–392. Jarvis, D.I., A.H.D. Brown, et al. 2008. A global perspective of the richness and evenness of traditional crop-variety diversity maintained by farming communities. Proc. Nat. Acad. Sci. USA 105:5326–5331.

2 Brassica and Its Close Allies: Cytogenetics and Evolution Shyam Prakash National Research Centre on Plant Biotechnology Indian Agricultural Research Institute New Delhi 110012 India S. R. Bhat National Research Centre on Plant Biotechnology Indian Agricultural Research Institute New Delhi 110012 India C. F. Quiros Department of Vegetable Crops University of California Davis, California 95616 USA P. B. Kirti Plant Science Department University of Hyderabad Hyderabad, 500046 India V. L. Chopra National Research Centre on Plant Biotechnology Indian Agricultural Research Institute New Delhi 110012 India

I. INTRODUCTION II. CYTOGENETICS A. Cytogenetic Architecture of Brassica Coenospecies B. Crop Species

Plant Breeding Reviews, Volume 31 Edited by Jules Janick Copyright & 2009 John Wiley & Sons, Inc. 21

22

III.

IV.

V. VI. VII.

VIII. IX.

X.

S. PRAKASH, S. R. BHAT, C. F. QUIROS, P. B. KIRTI, AND V. L. CHOPRA 1. Nature of Diploid Species 2. Nature of Alloploid Species 3. Nuclear DNA 4. Karyotypes 5. Pachytene Chromosomes 6. Satellite Chromosomes and rDNA Loci 7. Archetype and Evolution of Genomes GENOME MANIPULATION A. Resyntheses of Natural Allopolyploid Brassica spp. B. Agronomic Potential of Synthetics C. Diploidization of Allopolyploid Species D. Raphanobrassica E. Higher Allopolyploids in U Triangle Species through Protoplast Fusion WIDE HYBRIDIZATION A. Sexual Hybrids B. Somatic Hybrids C. Introgression of Genes CYTOPLASMIC SUBSTITUTION AND MALE STERILITY GENOME DISSECTION AND DEVELOPMENT OF CHROMOSOME ADDITION LINES MITOCHONDRIAL GENOME A. Organization B. Gene Content C. Mitochondrial Plasmids PLASTID GENOME POTENTIAL ROLE OF ARABIDOPSIS THALIANA IN BRASSICA IMPROVEMENT A. A. thaliana as a Model Crucifer B. Cytology and Possible Origin of the A. thaliana Genome C. Synteny Conservation D. Synteny-Based Gene Discovery and Cloning E. Arabidopsis Knowledge–Based Gene Discovery and Brassica Improvement 1. Understanding Domestication 2. Understanding Metabolism 3. Testing for Gene Function by Complementary Transformation CHLOROPLAST GENOMES AND THEIR PHYLOGENETIC IMPLICATIONS A. Subtribe Brassicinae 1. Brassica 2. Diplotaxis 3. Erucastrum 4. Sinapis 5. Trachystoma 6. Hirschfeldia incana 7. Sinapidendron 8. Coincya 9. Eruca B. Subtribe Raphaninae C. Subtribe Moricandiinae D. General Considerations

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XI. EVOLUTION OF MORPHOLOGICAL CHARACTERS A. Cotyledons B. Adult Leaves C. Fruits D. Isthmus Concept XII. CONCLUDING REMARKS ACKNOWLEDGMENTS LITERATURE CITED

ABBREVIATIONS ACO ADH AFLP Ag-NOR BACs BTL CAGs cp CP CMS Cytodeme DAPI ESTs FISH GISH GOT GSL IDH ISSR ITC ITS LAP MDH mt NOR PrBn 6 PGD 6 PGDH PGI PGM QTL

Aconitase Alcohol deshydrogenase Amplified fragment length polymorphism Silver-stained nucleolus organizer region Bacterial artificial chromosomes Binary trait loci Conserved Arabidopsis genome sequences Chloroplast Condensation pattern Cytoplasmic male sterility Crossing group 40 ,6-diamidino-2-phenylindole Expressed sequence tags Fluorescence in situ hybridization Genomic in situ hybridization Glutamate oxaloacetate transaminase Glucosinolate Isocitric dehydrogenase Inter-simple sequence repeats Isothiocynanates Internal transcribed spacers of nuclear ribosomal DNA genes Leucine amino-peptidase Malate dehydrogenase Mitochondria Nucleolus organizer region Pairing regulator Brassica napus 6-phosphogluconase dehydrogenase 6-phosphogluconate deshydrogenase Phosphoglucoisomerase Phosphoglucomutase Quantitative trait loci

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RAPD RFLP rDNA rRNA SDH SSR TE TPI

S. PRAKASH, S. R. BHAT, C. F. QUIROS, P. B. KIRTI, AND V. L. CHOPRA

Randomly amplified polymorphic DNA Restriction fragment length polymorphism Ribosomal DNA genes Ribosomal RNA Shikimic acid dehydrogenase Simple sequence repeats Transposable element Triose phosphate isomerase

I. INTRODUCTION Brassica species, Brassicaceae (Cruciferae), provide an important component of human diet as major sources of edible oil and vegetables (Table 2.1). The antiquity of crops belonging to the genus Brassica is manifest from references in ancient literature of the Indian, Chinese, Greek, and Roman civilizations (Prakash and Hinata 1980; Go´mezCampo and Prakash, 1999). A number of taxonomic treatments of this family are available since 1700. Prominent among these are by Tournefort (1700), Linnaeus (1753), deCandolle (1821), Hooker (1862), Baillon (1871), and Prantl (1891). However, the most comprehensive one is by Schulz (1919, 1936), a German schoolteacher (Hedge 1976; Prakash and Hinata 1980; Gomez-Campo 1999b). A recent molecular account of the family has been provided by Beilstein et al. (2006). Brassiceae is one among the 19 tribes recognized by Schulz in the family and is divided into 7 to 9 subtribes (Go´mez-Campo 1980, 1999b). Brassica is the core genus in the subtribe Brassicinae. Several members of related subtribes, such as Raphaninae and Moricandiinae, exhibit close genetic affinities with Brassica. However, morphological distintiveness of these three subtribes is not well substantiated and molecular data provide scanty support for their independent status. A majority of the species related to Brassica are wild and weedy. They possess, however, useful genes that may confer agronomic advantages and/or enhance the quality and utility of crop species. In fact, genetic enrichment of crop species with genes from wild allies is a major approach for many crop improvement programs. Such gene transfer can be achieved both by conventional plant breeding methods and through biotechnology. Cytogenetic investigations on Brassica, initiated in the second decade of the 20th century, were confined to determining the chromosome numbers and studying chromosome pairing in interspecific hybrids, which finally led to unraveling the genetic architecture of crop species. Small size of chromosomes, scarcity of distinctive cytogenetic landmarks,

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Table 2.1. Taxonomic components of Brassica and related genera and their usage. Botanical name B. nigra B. oleracea var. acephala var. capitata var. sabauda var. gemmifera var. gongylodes var. botrytis var. italica var. fruticosa var. alboglabra B. rapa spp. oleifera var. brown sarson var. yellow sarson var. toria ssp. rapifera ssp. chinensis ssp. pekinensis ssp. nipposinica ssp. parachinensis B. carinata B. juncea B. napus spp. oleifera spp. rapifera Eruca sativa Raphanus sativus Sinapis alba

Common name

Usage

black mustard

condiment (seed)

kale cabbage savoy cabbage brussels sprouts kohlrabi cauliflower broccoli branching bush kale Chinese kale

vegetable, fodder (leaves) vegetable (head) vegetable (terminal buds) vegetable (head) vegetable, fodder (stem) vegetable (inflorescence) vegetable (inflorescence) fodder (leaves) vegetable (stem, leaves)

turnip rape brown sarson yellow sarson toria turnip bok choi Chinese cabbage — — Ethiopian mustard mustard

oilseed oilseed oilseed oilseed fodder, vegetable (root) vegetable (leaves) vegetable, fodder (head) vegetable (leaves) vegetable (leaves) vegetable, oilseed oilseed, vegetable

rapeseed rutabaga, swede rocket, taramira radish white mustard

oilseed fodder vegetable, nonedible oilseed vegetable, fodder oilseed

and not being amenable to pachytene investigations were major deterrants to cytogenetical analyses. Advances in tissue culture techniques, including ovary and embryo rescue and protoplast fusion, since the 1950s made varied cytogenetic material available to investigate genome homologies and facilitated introgression of useful nuclear genes even across conventional generic boundaries. Such investigations require reliable markers for chromosome identification. A significant step toward this development has been the extensive use of molecular markers. Molecular biology in Brassica started with the determination of female parents of allopolyploid species using chloroplast DNA RFLPs by Palmer et al. (1983a). Use of genomic and fluorescence in situ hybridization (GISH and FISH respectively) methodology, in combination with ribosomal DNA markers have given new directions in genome analysis

26

S. PRAKASH, S. R. BHAT, C. F. QUIROS, P. B. KIRTI, AND V. L. CHOPRA

and characterization of parental genome components as well as precise identification of the individual chromosomes and location of gene sequences directly on the chromosomes. These investigations led to the generation of first FISH-based molecular karyotypes (Fukui et al. 1998). At the same time, chloroplast and mitochondrial DNA RFLPs have been used extensively to elucidate phylogeny of Brassica and related genera. Molecular markers also have been identified to tag important agronomic traits. This research has not only substantiated some of the already existing concepts but also proposed several new ones. Potential sources of germplasm have been identified outside of the conventional boundaries, thus increasing the range of available germplasm relevant to Brassica improvement. Genomic studies on Arabidopsis, a crucifer and closely related to brassicas, has given a new direction in evolutionary studies of the family Brassicaceae and in particular the members of the genus Brassica. Inferences from comparative genomics between Arabidopsis and Brassica species have elucidated evolutionary processes. Arabidopsis has become a model plant in the field of experimental biology because of its several unique features: short life span, autogamy, and ease of tissue culture. Its entire genome has recently been sequenced (Arabidopsis Genome Initiative 2000). Biologically, Brassica and allied taxa have been grouped collectively and referred to as Brassica coenospecies (Harberd 1972). This review is an attempt to synthesize available literature and developments in Brassica coenospecies from classical to molecular cytology and application of genomic information to throw light on genome organization, genome manipulation, and phylogeny in Brassica and related genera. Informative reviews dealing with some of these aspects are given in Biology of Brassica Coenospecies (Go´mez-Campo 1999a) and Biotechnology in Agriculture and Forestry: Brassica (Pua and Douglas 2004). II. CYTOGENETICS A. Cytogenetic Architecture of Brassica Coenospecies Brassica coenospecies is not a taxonomic but a biological unit. More appropriately it is a cytogenetical concept. It is defined as that part of tribe Brassiceae of Schulz whose members are closely related to crop Brassica species and are potentially capable of exchanging genetic material with them. Harberd (1972) was the first to propose this concept on the basis of his extensive cytotaxonomical investigations on Brassica crop and related wild species. Harberd’s focus was on subtribe

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Brassicinae of Schulz comprising of 11 genera and also on a part of related subtribe Raphaninae. His investigations involved studies on chromosome pairing and extent of fertility of hybrids. Harberd proposed that nine genera from subtribe Brassicinae—Brassica, Coincya, Diplotaxis, Eruca, Erucastrum, Hirschfeldia, Sinapis, Sinapidendron, and Trachystoma—and two genera from subtribe Raphaninae—Enarthrocarpus and Raphanus—constitute Brassica coenospecies. Harberd (1972) involved a wide spectrum of species in his hybridization program and studied this germplasm biologically rather than taxonomically to classify it into cytodemes or crossing groups to resolve the confusion about their species and generic status. A cytodeme is defined as a group consisting of any number of species or genera that have the same chromosome number, and crosses between them always yield fertile hybrids. Harberd (1972) established 38 cytodemes in the coenospecies. This number was further extended by Takahata and Hinata (1983). In fact, Harberd’s results revealed, for the first time, extensive genome homoeology across species and generic boundaries, implying that Brassica coenospecies constitutes a large gene pool and thus opening the possibilities of transferring agronomically desirable traits to crop species. The boundaries of coenospecies have further expanded with developments in molecular biology that have resulted in massive incongruities with established taxonomy. Chloroplast DNA RFLP studies on members of other related subtribes also suggest that delemitation of genera and species by Schulz does not fully reflect the natural boundaries (Warwick and Black 1991; Pradhan et al. 1992). These investigations strongly support the inclusion of not only Raphanus and Enarthrocarpus in the coenospecies as suggested by Harberd (1972, 1976), but also of three more genera—Moricandia, Pseuderucaria, and Rytidocarpus—from the related subtribe Moricandiinae (Warwick and Black 1997). At present, 63 cytodemes are recognized in coenospecies that spread over 14 taxonomically defined genera, as shown in Fig. 2.1 and Table 2.2 (Prakash et al. 1999). Determination of chromosome number of B. campestris syn. B. rapa (Takamine 1916) was the beginning of cytogenetic research on Brassica and in the Brassicaceae. Chromosome numbers for many species were determined between 1928 and 1932 (Jaretzky 1932; Manton 1932). Manton (1932) did pioneering work on cytology of Brassicaceae, publishing new chromosome reports and reviewing the work on the whole family. Credit for initiating cytogenetical researches on crop species goes to Karpechenko of Russia, Morinaga and U of Japan. Mizushima of Japan deserves the credit for work on wild germplasm. These researchers investigated different aspects and elucidated genomic homoeologies in a

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Table 2.2. Cytodemes in Brassica coenospecies. Chromosome no. (n) 7

8

9

10

11

12

13 14

Principal species Brassica deflexa Boiss. Diplotaxis erucoides (L.) DC. Erucastrum virgatum C. Presl. Erucastrum varium Durieu Sinapis aucheri (Boiss.) O.E. Schulz Hirschfeldia incana (L.) Lagreze-Fossat Pseuderucaria spp. O.E. Schulz Brassica nigra (L.) Koch Brassica fruiticulosa Cyr. (þ maurorum þ spinescens) Diplotaxis siettiana Maire Erucastrum abyssinicum (A. Rich.) O.E. Schulz Erucastrum nasturtiifolium (Poiret) O.E. Schulz (þ leucanthum) Erucastrum strigosum (Thunb.) O.E. Schulz Trachystoma spp. Brassica oleracea L. and 8 wild Mediterranean allied species Brassica oxyrrhina Coss. Diplotaxis assurgens (Del.) Gren. Diplotaxis catholica (L.) DC. Diplotaxis tenuisiliqua Del. Diplotaxis virgata (Cav.) DC. Diplotaxis berthautii Braun-Blanq. and Maire Erucastrum cardaminoides Webb and Berth. (þ canariense þ ifniense) Raphanus L. all species and subspecies Sinapis arvensis L. (þ allioni) Sinapis pubescens L. Brassica tournefortii Gouan. Brassica barrelieri (L.) Janka Brassica gravinae Ten. Brassica repanda (Willd.) DC. (þ desnottesii) Brassica rapa L. (þ many cultivated subspecies) Diplotaxis siifolia G. Kunze Diplotaxis viminea (L.) DC Enarthrocarpus spp. Sinapidendron spp. Brassica souliei Batt. Diplotaxis acris (Forsk.) Boiss. Brassica elongata Ehrh. Diplotaxis tenuifolia (L.) DC. (þ pitardiana) Eruca spp. Mill. Coincya spp. (syn. Hutera and Rhynchosinapis) Sinapis alba L. Sinapis flexuosa Poir. Diplotaxis harra (Forsk.) Boiss. (þ several subsps.) Erucastrum virgatum C. Presl. (subsp. pseudosinapis) Moricandia arvensis (L.) DC. Moricandia moricandioides (Boiss.) Heywood Rytidocarpus moricandioides Coss.

2. BRASSICA AND ITS CLOSE ALLIES: CYTOGENETICS AND EVOLUTION Table 2.2. (Continued) Chromosome no. (n) 15

Principal species Erucastrum gallicum (Willd.) O.E. Schulz Erucastrum elatum (Ball.) O.E. Schulz Brassica cossoniana (Boiss. & Reut.) (4x) North African subspecies Brassica balearica Pers. Erucastrum nasturtiifolium (Poiret) O.E. Schulz (4x) Erucastrum abyssinicum (A. Rich.) O.E. Schulz (4x) Brassica carinata A. Braun Brassica juncea (L.) Czern & Coss. Brassica napus L. Brassica gravinae Ten. (4x) Diplotaxis muralis (L.) DC. Brassica dimorpha Coss. & Dur. Coincya spp. (4x) Moricandia suffruticosa (Desf.) Coss. & Dur. Moricandia spinosa Pomel Brassica repanda (Willd.) DC. (High Atlas)

16

17 18 19 20 21 22 24 28 42 80?

Source: From Prakash et al. 1999; C. Go´mez-Campo, personal communication.

Family

Brassicaceae

Tribe

Brassicaceae

Subtribe

Brassicinae

Genera

Brassica (20) Coincya (1) Diplotaxis (13) Eruca (1)

Raphaninae

Enarthrocarpus (1) Raphanus (1)

Moricandiinae

Moricandia (1) Pseuderucaria (1) Rytidocarpus (1)

Erucastrum (11) Hirschfeldia (1) Sinapis (5) Sinapidendron (1) Trachystoma (1) Fig. 2.1. Archetecture of Brassica coenospecies (number of cytodemes in brackets. (Source: From Prakash et al. 1999.)

29

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S. PRAKASH, S. R. BHAT, C. F. QUIROS, P. B. KIRTI, AND V. L. CHOPRA

series of publications. Extensive taxonomical investigations on wild germplasm have been carried out by Go´mez-Campo (1999b). The lowest chromosome number in coenospecies, n ¼ 7, is characteristic of seven cytodemes. Harberd (1972) was of the view that cytodeme with n ¼ 14 or higher chromosome numbers should be attributed to polyploidy. According to this view, 43 cytodemes are diploids where every chromosome number from n ¼ 7 to n ¼ 13 is represented. However, variations in isozyme numbers of a vast range of taxa in the tribe Brassiceae suggest that genera with n ¼ 14 to 18 are not necessarily polyploids of n ¼ 7 to 13 genomes (Anderson and Warwick 1998). Around 50% of the cytodemes have gametic chromosome number n ¼ 9 and n ¼ 10. Polyploidy also played a role as both auto- and allopolyploids are represented by 20 cytodemes (Table 2.3). The majority are tetraploids. This polyploidy level is exceeded only in some accessions of Moricandia spinosa (2n ¼ 84, x ¼ 6) and Brassica repanda (2n ¼ 160, x ¼ 8) (Prakash et al. 1999). The genus Moricandia seems to be exclusively polyploid (Al-Shebaz 1984). B. Crop Species Genome analysis in crop species, pioneered by Morinaga (1928; 1929a, b,c; 1931; 1933; 1934a,b) was based on hybridizing high-chromosome species with low-chromosome species and interpreting the chromosome pairing behavior of the hybrids. This research led Morinaga (1934) to propose that crop brassicas comprise six species. Of these, three are low-chromosome monogenomic diploids—B. nigra (n ¼ 8), B. oleracea (n ¼ 9), and B. rapa (syn. B. campestris, n ¼ 10)—and three are high-chromosome digenomics—B. carinata (n ¼ 17), B. juncea (n ¼ 18), and B. napus (n ¼ 19), which evolved in nature through convergent alloploid evolution between any two of the diploid species. Morinaga also assigned genome symbols to these species. U (1935) represented this cytogenetical relationship diagramatically, in what is now commonly referred to as U triangle (Fig. 2.2). These relationships have, in recent years, been substantiated by cytogenetics, molecular analysis of nuclear and chloroplast DNA, and by genomic and fluorescence in situ hybridization (Snowdon et al. 2003; Snowdon 2007). This complex of diploids and allopolyploids is now considered a model system for investigations on polyploidy in crop plants (Lukens et al. 2006; Pires et al. 2006). 1. Nature of Diploid Species. The three basic diploid species, B. nigra, B. oleracea, and B. rapa, represent an ascending aneuploid series

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Table 2.3. Polyploid cytodemes in Brassica coenospecies. Allopolyploids Brassica Brassica Brassica Brassica

carinata, n ¼ 17 juncea, n ¼ 18 napus, n ¼ 19 balearca, n ¼ 16

Diplotaxis muralis, n ¼ 21 Erucastrum gallicum, n ¼ 15 Erucastrum elatum, n ¼ 15 Tentative autopolyploids Moricandia arvensis, n ¼ 14 Moricandia moricandiodes, n ¼ 14 Rytidocarpus moricandiodes, n ¼ 14 Erucastrum virgatum (subsp. pseudosinapis), n ¼ 14 Brassica cossoneana, n ¼ 16 Erucastrum abyssinicum, n ¼ 16 Erucastrum nasturtiifolium, n ¼ 16 Brassica gravinae, n ¼ 20

Diploid progenitors B. B. B. B.

nigra, B. oleracea rapa, B. nigra oleracea, B. rapa oleracea group  another species D. viminea, D. tenuifolia

E. leucanthum  sp.? Hirschfeldia incana  Erucastrum littoreum Diploid homolog

Reference U 1935 U 1935 U 1935 Snogerup and Persoon 1983 Harberd 1976; Mummenhoff et al. 1993: Ueno et al. 2006 Harberd 1976 Go´mez-Campo 1983; Sa´nchez-Ye´lamo 1992 Reference

unknown

Harberd 1976

unknown

Harberd 1976

unknown

Harberd 1976

E.virgatum B. maurorum, n ¼ 8 E. abyssinicum, n ¼ 8 E. nasturtiifolium, n ¼ 8

Harberd 1976 Pradhan et al. 1992 Harberd 1976 Harberd 1976

B. gravinae, n ¼ 10

Takahata and Hinata 1983 Go´mez-Campo 1980 Harberd 1976 Sobrino-Vesperinas 1980 Sobrino–Vesperinas 1980 Galland 1988

Brassica dimorpha, n ¼ 22 Coincya spp., n ¼ 24 Moricandia suffruticosa, n ¼ 28 Moricandia spinosa, n ¼ 42

B. soullei, n ¼ 11 Coincya sp., n ¼ 12 Moricandia sp., n ¼ 14 Moricandia sp., n ¼ 14

Brassica repanda, n ¼ 80

B. repanda, n ¼ 10

Source: From Prakash et al. 1999.

(Manton 1932) and are regarded as secondary polyploids. Evidence for this conclusion were adduced from chromosome associations at meiosis in their respective haploids (Thompson 1956; Prakash 1974b; Armstrong and Keller 1981). Investigations on pachytene chromosome analysis (Ro¨bbelen 1960; Venkateswarlu and Kamla 1971), isozyme markers, and rDNA genes (Quiros et al. 1987) suggested that these species originated from a now-extinct archetype with a probable basic chromosome number of x ¼ 6. It was believed that selective doubling of some chromosomes in this archetype led to the evolution of the three diploid genomes. However, results of recent investigations on nuclear,

32

S. PRAKASH, S. R. BHAT, C. F. QUIROS, P. B. KIRTI, AND V. L. CHOPRA

B. carinata n = 17, bc

B. oleracea

B. nigra

n = 9, c

n = 8, b

B. napus

B. rapa

n = 19, ac

n = 10, a

B. juncea n = 18, ab

Fig. 2.2. Cytogenetic relationships of crop brassicas (U, 1935). Solid and broken lines in the allopolyploids represent female and male parents. respectively) (Source: According to Palmer et al. 1983a).

mitochondrial, and chloroplast DNA (Palmer 1988; Song et al. 1988a; Warwick and Black 1991; Pradhan et al. 1992) have discounted this theory of monophyletic origin and have instead suggested their origin from two linages: B. oleracea and B. rapa originating from one archetype and B. nigra evolving from the other. Nevertheless, these genomes share close homologies, as revealed by cytogenetical (Mizushima 1950a; Prakash and Hinata 1980; Attia and Ro¨bbelen 1986) and molecular studies (Hosaka et al. 1990; Teutonico and Osborn 1994; Truco et al. 1996; Parkin et al. 2003). Cytogenetical investigations in digenomic and trigenomic interspecific hybrids involving the three basic species showed high frequency of bivalents and multivalents. A good number of these were suggested to arise due to allosyndesis. GISH analysis confirmed three allosyndetic bivalents between B and A/C (Ge and Li 2007) and five bivalents between A and C genomes (Liu et al. 2006). All three genomes contain similar genetic information with many duplications (Slocum et al., 1990; Chyi et al. 1992; Jackson et al. 2000; Parkin et al. 2003); just the organization and distribution on chromosomes is different (Truco et al. 1996). Chromosome differentiation and repatterning has occurred mainly through duplications and translocations (Quiros et al. 1988; Hosaka et al. 1990; McGrath et al. 1990; Truco and Quiros 1994) and also deletions (Hu and Quiros 1991). These changes were tolerated and adjusted because of the secondary

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33

balanced nature of these genomes (Kianian and Quiros 1992a). Also, a large number of rearrangements separated the B genome from the A or C genome. In comparison, A and C genomes are less differentiated (Lagercrantz 1998). Genomes A and C are also cytogenetically very close (Mizushima 1950a; Olsson 1960b), a fact substantiated by: (1) FISH mapping of two families of repetitive DNA that are common to pericentromeric regions of most chromosomes of A and C genomes but are absent in the B genome (Harrison and Heslop-Harrison 1995); (2) structural analysis of rDNA intergenic spacers (Bhatia et al. 1996); (3) colinearity between them as revealed by comparative analysis (Scheffler et al. 1997); and (4) extent of homoeologous pairing detected by GISH (Snowdon et al. 1997a; Ge and Li 2007) and FISH and molecular markers (Nicolas et al. 2007). Interestingly, RFLP analysis of rDNA reveals, on the contrary, closer affinities between B and C genomes (Hasterok and Maluszynska 2000a). Also, as detected by microsatellites (Bornet and Blanchard 2004), the C genome is more conserved than A or B. Among the three basic species, two types of cytoplasm exist: the B type found in B. nigra and the A/C type occurring in B. rapa and B. oleracea. The A and B types are quite distinct although they retain homology to a large extent (Palmer et al. 1983a; Yanagino et al. 1987; Warwick and Black 1991; Pradhan et al. 1992).

2. Nature of Alloploid Species. The three high-chromosome species, B. carinata, B. juncea, and B. napus, originated through convergent alloploid evolution involving different combinations of the diploid species. This is amply demonstrated by several sets of evidence including taxonomy, artificial syntheses, molecular analysis, and chromosome mapping. The allopolyploids have attained a balance forming regular bivalents and are devoid of any homoeologous pairing. Information from Fraction-1 protein (Uchimiya and Wildman 1978) and chloroplast and mitochondrial DNA restriction patterns (Palmer et al. 1983a; Ichikawa and Hirai 1983; Erickson et al. 1983; Yanagino et al. 1987; Warwick and Black 1991; Pradhan et al. 1992; Cunha et al. 2004) have revealed directionality of natural hybridizations. It has conclusively been established that B. nigra and B. rapa contributed the cytoplasm to B. carinata and B. juncea, respectively. However, patterns of chloroplast DNA restriction of B. napus were different from both B. rapa and B. oleracea but approached that of B. oleracea (Erickson et al. 1983; Palmer et al. 1983a). Studies on comparative chloroplast and mitochondrial RFLP patterns strongly implicated B. montana, belonging to B. oleracea group, as the cytoplasm donor of B. napus (Song and

34

S. PRAKASH, S. R. BHAT, C. F. QUIROS, P. B. KIRTI, AND V. L. CHOPRA

Osborn 1992). However, plastid SSRs analysis does not support this view; on the contrary, it suggests that B. rapa is the more likely plastid genome donor of B. napus (Flannery et al. 2006). Various investigations also suggest that while chloroplast genomes have been conserved in B. carinata and B. juncea since their origin, the chloroplast genome of B. napus has gone through evolutionary alteration (Palmer et al. 1983a). Both mitochondrial and chloroplast genomes in all the three species have been coinherited over generations (Palmer 1988). It has also been observed that cytoplasm has exerted considerable influence on the evolution of nuclear genomes of alloploid species. When the parental diploid species of allopolyploid has highly differentiated cytoplasm, as in B. juncea and B. carinata, the nuclear genomes contributed by the male parents were altered considerably more compared to the nuclear genomes of female parents (Song et al. 1988a, 1995). Thus in B. juncea, nuclear genome A has remained mostly intact while nuclear genome B has changed considerably; B. carinata has B nuclear genome unchanged but considerably altered C genome. In B. napus, both A and C genomes have undergone a similar extent of changes. However, ISSR marker analysis suggests that B-genome-carrying allopolyploid species B. carinata and B. juncea underwent more modifications than B. napus. Also, in B. napus, A genome has changed more than C genome; A genome in B. juncea and C genome in B. carinata are drastically modified; and B. juncea and B. carinata have relatively conserved B genome (Liu and Wang 2006). Strong evidence suggests rapid and extensive structural rearrangements of chromosomes in these alloploid species since their evolution (Slocum et al. 1990; Song et al. 1991, 1995; Kianian and Quiros 1992a,b; Poulsen et al. 1993; Harrison and HeslopHarrison 1995). In fact, chromosomal rearrangements arising from homoeologous recombination between A and C genomes are widespread. Referred to as interstitial homoeologous reciprocal translocations they are a cause for origin of morphological variants in B. napus (Osborn et al. 2003; Udall et al. 2005) and are considered a progressive step in evolution (Nicolas et al. 2007). Morphological variants of nonhomologous origin have been obtained in B. juncea (Prakash 1973b). Occurrence of such events was earlier inferred from analysis of segregating populations employing molecular markers (Parkin et al. 1995; Sharpe et al. 1995). However, comparative analysis of RFLP marker-based chromosome maps of natural and synthesized B. juncea does not support Song et al. (1995) and indicate that B. juncea has remained largely unchanged since its origin (Axelsson et al. 2000). In spite of incorporation of these changes, the average chromosome size has remained unaltered (Parkin and Lydiate 1997).

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35

3. Nuclear DNA. Nuclear DNA content and nuclear volume were first estimated by Yamaguchi and Tsunoda (1969) in B. rapa, B. oleracea, and naturally occurring and synthetic strains of B. napus. They observed that the values for synthetic B. napus were the sum of the constituent parents. However, there was an appreciable reduction in total DNA content in natural forms. These authors proposed that nuclear DNA had been lost subsequent to evolution of allotetraploids. Verma and Rees (1974) further investigated this problem by estimating the amount of DNA in diploids and their allotetraploid derivatives in root meristem nuclei at the GI phase. No significant intraspecific variation in nuclear DNA amount was observed. However, differences exist at the interspecific level. In spite of the fact that values for allotetraploids were very close to the sum values of their constituent parents, reduction from the expected values for every species was observed. They postulated that the lower values in tetraploids result from underestimation of DNA due to higher nuclear density. It was also suggested that values observed by Yamaguchi and Tsunoda (1969) were based on dense nuclei and were underestimations; thus, when corrected, the values showed no significant deviations from those anticipated. Therefore, decrease in the amount of DNA was not associated with allopolyploidy. One significant observation by Verma and Rees (1974) was a remarkable reduction in nuclear size in natural allotetraploids, which suggested condensation of chromosomal material that probably reflected an adaptive switching off of redundant gene copies. In several recent investigations, DNA values have been estimated afresh (Arumugunathan and Earle 1991; Narayan 1998; Bennet and Leitch 2005; Johnston et al. 2005). A general observation is the evolution of DNA content from low to high in the genus. These studies also support the earlier observations of Yamaguchi and Tsunoda (1969) that there has been a decrease in DNA content in the present-day alloploid species. A decrease of 6% was observed by Narayan (1998), and the values for B. napus, B. juncea, and B. carinata are 0.095, 0.094, and 0.049 pg less respectively than the sum of their parental species (Johnston et al. 2005, Table 2.4). 4. Karyotypes. Brassica chromosomes are very small in size, as indicated by haploid 1C genome size (Bennet and Leitch 2005). They are also poorly differentiated in structure without distinctive cytological landmarks. Consequently, to date no precise nomenclature is available. Karyotype studies were initiated as early as 1934 by Catcheside and were followed by Alam (1936), Richharia (1937), and Sikka (1940). These investigations were confined to observations on the frequency of various chromosome types, number of satellites, and

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S. PRAKASH, S. R. BHAT, C. F. QUIROS, P. B. KIRTI, AND V. L. CHOPRA

Table 2.4. 1c nuclear DNA content and genome size in Brassica species. Species B. B. B. B. B. B.

nigra oleracea rapa carinata juncea napus

1c nuclear DNA content (pg  se)

Genome size (1 x) (Mbp)

0:647  0009 0:710  0:002 0:539  0:018 1:308  0:018 1:092  0:001 1:154  0:006

632 696 529 642 534 566

Source: From Johnston et al. 2005.

nucleoli in different genomes. They distinguished chromosomes into long, medium, small, and very small with median, subterminal and terminal constrictions. In recent years, karyotypes, particularly of diploid species, based on mitotic (Olin-Fatih and Heneen 1992; Cheng et al. 1995a; Fukui et al. 1998; Hasterok and Maluszynska 2000a; Hasterok et al. 2005a) and meiotic chromosome (Cheng et al. 1994b; Mackowiak and Heneen 1999; Koo et al. 2004) phenotypes, have been constructed using different staining. Mitotic prometaphase and meiotic diakinesis offer better possibilities for characterizing individual chromosomes and constructing karyotypes. Since early 1990s, use of FISH with ribosomal DNA probes has further helped in generating chromosome markers. Molecular karyotypes based on FISH have been generated, enabling more reliable identification of individual chromosomes. Maluszynska and Heslop-Harrison (1993), Snowdon et al. (1997a) and Fukui et al. (1998) employed FISH with a 45S rDNA probe for individual chromosome identification. However, simultaneous probing with 45S rDNA and 5S rDNA in B. napus, Sinapis alba and Raphanus sativus (Schrader et al. 2000) and in all the six species of U triangle (Hasterok et al 2001) proved to be more informative as they provided numerous signals on somatic chromosomes revealing new landmarks. Fukui (1998) developed a system for computer imaging of plant chromosomes that led to the definition of a new parameter, the ‘‘condensation pattern’’ (CP) for chromosome analysis. It is an effective and reproducible parameter and very useful in identification of small chromosomes. It is a general observation that somatic chromosomes of A and C genomes are morphologically very similar and difficult to distinguish (Olin-Fatih and Heneen 1992) although their condensation patterns differ in prometaphase chromosomes (Cheng et al. 1995a). However, making use of FISH and GISH, it has been possible to identify individual chromosomes of A, B, and C genomes and also to match chromosomes with corresponding counterparts in alloploid species

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37

with considerable reliability (Snowdon et al. 1997b, 2002; Kamisugi et al. 1998; Maluszynska and Hasterok 2005). Also, these investigations are very helpful in integrating genetical maps, emerging from analysis of molecular markers, with physical maps based on morphometric analysis. Although many papers describing the karyotypes of these species have appeared since 1937, overall conclusions can be summarized in this way: Chromosomes of A genome are morphologically most diverse, B genome chromosomes are much more uniform and difficult to identify individually, and C genome chromosomes are poorly differentiated in morphology and size, and undergo variable degree of condensation of hetero- and euchromatin in the chromosome arms (Olin-Fatih 1994). Discrepencies in nomenclature and numbering of chromosomes have occurred due to polymorphisms in the rDNA sites and contraction rates of the chromosomes. Brassica nigra. Hasterok and Maluszynska (2000a) observed that B. nigra chromosomes are more or less similar in size, ranging from 2.47 to 3.57 mm, and are morphologically undistinguishable. Only two types of chromosomes are present: median (no. 1–4) and submedian (no. 5–8). Chromosomes 7 and 8 contain secondary constriction and satellite on the short arm. Earlier, Mackowiak and Heneen (1999) presented a karyotype based on diakinesis bivalents wherein each chromosome exhibited a specific pattern of chromatin condensation or darkly stained regions. The eight bivalents were classified into three groups. Group 1, comprising pairs 1 and 2, has darkly stained median position signifying pericentric chromatin. Pair 2 was the smallest. Group 2 comprised pairs 3 to 5 with a submedian-subterminal, darkly stained region that represent pericentric chromatin. Group 3 comprised pairs 6 to 8 with relatively large-size subterminal-terminal darkly stained region. Pair 6 was larger than pairs 7 and 8. Pairs 6 to 8 were the satellited nucleolar chromosomes. Brassica oleracea. Cheng et al. (1995a) described a karyotype for B. oleracea where the absolute length ranged from 2.8 to 4.5 mm. The genome is comprised of three median group (1–3), four submedian group (4–7), and two subterminal group chromosomes with a nonsatellite pair (8) and a satellite pair (9). Armstrong et al. (1998) reported partial karyotype of B. oleracea var. alboglabra based on cytological landmarks (centromere locations) provided by FISH localization of three repetitive DNA sequences. Five of the nine chromosomes could be identified with precision.

38

S. PRAKASH, S. R. BHAT, C. F. QUIROS, P. B. KIRTI, AND V. L. CHOPRA

Chromosomes 2 and 4 carry 45S rDNA clusters. Chromosome 2 is also submetracentric. Chromosomes 4, 6, 8, and 9 are very much similar. In all, there are seven metacentric/submetacentric and two pairs of satellited acrocentric chromosomes. Hasterok and Maluszynska (2000a) studied a B. oleracea karyotype where chromosomes are poorly differentiated and are in the range of 2.17 to 2.97 mm. The karyotype consists of 1–4 median and 5–8 submedian chromosomes. Chromosome 9 has a secondary constriction and satellite on the short arm. Brassica rapa. Karyotypes of B. rapa, in the earlier studies, have been constructed on Giemsa-stained mitotic metaphase or pro-metaphase chromosomes (Nishibayashi 1992; Olin-Fatih and Heneen 1992; OlinFatih 1994; Cheng et al. 1995a). Making use of condensation pattern profiles and FISH patterns employing a 45S rDNA probe, Fukui et al. (1998) generated a karyotype of B. rapa. Subsequently, Snowdon et al. (2002) and Lim et al. (2005) also used FISH for chromosomal localization of rDNA loci and generated a karyotype. Cheng et al. (1995a) observed four large-sized chromosome pairs (1–4) in the median group, four relatively small-size pairs (5–8) in the submedian group, and a satellited (9) and a nonsatellited pair (10) in the subterminal group. The absolute length ranged from 2.1 to 5.2 mm. In general, chromosome 1 is the largest; chromosomes 2 and 4 are of similar size; chromosome 5 is the large one in submediann group; chromosome 6 has a less submedian centromere than chromosome 7. Chromosome 7 has a submedian centromere and is larger than chromosome 8. Chromosome 9 has a satellite on the short arm. However, Kim et al. (1998) reported six median, two submedian, and two subterminal chromosomes. Hasterok and Maluszynska (2000a) characterized individual chromosomes in the A genome morphologically. The absolute length ranged from 1.77 to 3.73 mm. Based on centromere position, there are five median (1–5), three submedian (6–8), and two subterminal pairs (9–10). Chromosome 10 has a secondary constriction and a prominent satellite on the short arm. Recently, Koo et al. (2004) developed a molecular cytogenetic map of B. rapa ssp. pekinensis. The average length of mitotic metaphase chromosomes is in the range of 1.46 mm to 3.30 mm. Earlier, Kim et al. (1998) observed the range 1.01 to 2.06 mm for B. rapa. A map of B. rapa var. pekinensis was developed of mitotic metaphase chromosomes using 45S and 5S rDNA and pericentromeric tandem repeats as probes (Lim et al. 2005). Centromere index varied from 50.7% to 32.9%, indicating that the chromosomes are metacentric or submetacentric. The average total length of the haploid complement was 32.47 mm. Chromosomes 1 and 2 were the longest (4.5 and 5.83 mm respectively),

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the latter one being a nucleolus chromosome including the satellite and NOR. The short arm of this chromosome possessed 45S and 5S rDNA sites. Chromosomes 1, 3, 4, and 5 had 45S rDNA loci in their long arm. Chromosome 10 was the shortest (2.85 mm), and its short arm occupied a 5S rDNA site. The number of rDNA sites in the interphase nuclei varied from 4 to 10. The three high-chromosome allotetraploid species have numerous chromosomes, making karyotype formation very difficult when based just on morphometric features. In recent investigations, Hasterok and Maluszynska (2000b) and Kulak et al. (2002) have presented karyotypes of B. carinata, B. juncea, and B. napus on features combining morphometric information and multicolor FISH. Brassica carinata. Its karyotype consists of fairly uniform chromosomes, both in morphology and in length, ranging from 1.56 to 2.40 mm. Two groups of chromosomes can be distinguished: median (1–6) and submedian (7–15). There are two pairs of satellite chromosomes (16–17) with distinct secondary constrictions (Kulak et al. 2002). Brassica juncea. The chromosome length in B. juncea ranges from 1.38 to 3.25 mm, and the karyotype comprises of median (1–6) and submedian groups (7–15).Two chromosomes (17–18) are NOR-bearing with prominent secondary constrictions in the short arm. Chromosome 16, although NOR bearing, does not have a distinct secondary constriction/satellite region (Kulak et al. 2002). The extent of variations in chromosome size and morphology is due to A genome chromosomes. Brassica napus. This karyotype has been studied extensively (OlinFatih and Heneen 1992; Skarzhinskaya et al. 1998; Hasterok and Maluszynska 2000b; Snowdon et al. 2000a, 2002; Wei et al 2005) and is comprised of relatively small and poorly differentiated chromosomes. According to Hasterok and Maluszynska (2000b) and Kulak et al. (2002), B. napus chromosomes range from 1.52 to 3.3 mm in length and are small and similar in shape. They occur in three groups: median/ metacentric (1–4), submedian/submetacentric (5–17), and subterminal/subtelocentric (18–19), which carry satellites. Although chromosome condensation patterns in B. oleracea and B. rapa differ in prometaphase chromosomes (Cheng et al. 1995a), Kamisugi et al. (1988) were successful in identifying B. rapa–type and B. oleracea– type chromosomes in B. napus. A direct association with their homologs, however, could not be made. Distribution of rRNA genes on

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S. PRAKASH, S. R. BHAT, C. F. QUIROS, P. B. KIRTI, AND V. L. CHOPRA

chromosomes can be helpful in this regard. For example, A genome chromosomes are characterized by pericentromeric localization, and C genome chromosomes have terminally distributed rRNA genes (Hasterok and Maluszynska 2000b). These were clearly identified using rDNA hybridization and DAPI staining by Snowdon et al. (2002). Another B. napus karyotype has been constructed based on Cot-1 DNA FISH banding patterns by Wei et al. (2005). Their results agreed with the earlier reports. It was demonstrated that this technique can be used with precision to identify individual chromosomes and would be very helpful in recognizing homologous and nonhomologous chromosome pairing. 5. Pachytene Chromosomes. The cytologically difficult nature of material has restricted investigation on pachytene chromosome morphology to only a few attempts. Ro¨bbelen (1960), for the first time, analyzed pachytene chromosomes in the three basic species: B. nigra, B. oleracea, and B. rapa. The chromosomes revealed differentiation into proximal heterochromatic and distal euchromatic segments. Individual chromosomes within the genomes were identified by the number, size, and distribution pattern of the heterochromatic segments near the centromeres. The chromosomes were classified on the basis of their absolute length and were distinguished into five different types: very short (up to 20 mm); short (20–25 mm); medium (25–30 mm); long (30–40 mm); and very long (more than 40 mm). The B genome of B. nigra consists of one very long, four long, and three medium chromosomes of which five are metacentric; chromosome 3 (type C) has a satellite. B. oleracea, representing the C genome, has one very long, five long, two medium, and one short chromosomes. One chromosome is subtelocentric (nucleolus organizing chromosome No. 1, type A), two are submetacentric, and the rest are metacentric. The A genome B. rapa is characterized by one very long, four long, three medium, one short, and one very short chromosomes. Four chromosomes have subterminal and two have median centrometres. Nucleoli are attached to chromosomes 1 and 2 (both of type A) in B. rapa, to chromosomes 1 and 4 (types A and C) in B. oleracea, and to 1 and 3 (types A and C) in B. nigra. B. nigra genome has higher content of heterochromatin as compared to B. oleracea and B. rapa. Ziolkowski and Sadowski (2002) presented a high-resolution pachytene karyotype of B. oleracea using FISH. Each bivalent had a pericentromeric, heterochromatic region devided by a centromere. Up to 11 chromatin blocks corresponding to 9 pericentromeric heterochromatic

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regions and 2 nucleolus organizer regions were observed. Two NORassociated chromosomes were acrocentric, containing heterochromatin blocks at the ends of their short arm, and were designated C4 and C7. A prominent chromomere was present on the long arm of a submetacentric chromosome. Submetacentric chromosome C2 displayed three 5S rDNA loci on the same arm with medium (M), strong (S) and weak (W) FISH intensities. While locus M was very close to centromere, the two adjacent loci, S and W, were more distal. These three loci offer prominent landmarks for C2 chromosome of B. oleracea. Pachytene chromosome karyotype of B. rapa generated by Koo et al. (2004) was based on multicolor FISH and comprised of two metacentric (nos. 1, 6), five submetacentric (nos. 3, 4, 5, 9, and 10), two subtelocentric (nos. 7 and 8), and one acrocentric (no. 2) chromosomes. Their corresponding centromeric index ranges were 38.8% to 41.0%, 29.5% to 36.7%, 17.4% to 20.2%, and 9.38% respectively. The mean lengths varied from 23.7 to 51.3 mm with a total of 385 mm. As compared to mitotic metaphase chromosome length (1.46–3.30 mm), it is 17.5-fold higher. DAPI staining revealed variable length of heterochromatic blocks in the pericentromeric regions of all the chromosomes. Also, small heterochromatic regions, with a total length of 38.2 mm and approximately 10% of the total length of pachytene chromosomes, were observed on the long arm of chromosomes 3, 4, 5, and 7. FISH indicated 5S rDNA loci on pericentromeric regions of the short arms of chromosomes 2 and 10 and the long arm of chromosome 7. Similarly, 45S rDNA loci were observed on pericentromeric regions of short arms of chromosomes 1, 2, 4, and 5 and the long arm of chromosome 7. A 5S rDNA locus, observed on the long arm of bivalent no. 7, had not been detected on mitotic metaphase chromosomes in any earlier investigations. Ro¨bbelen (1960) recognized six basic types of chromosomes in each genome based on absolute length, symmetry of arms, and shape of heterochromatic centromeric region. These six types are: ‘‘A’’ with a distal heterochromatic satellite involved in nucleolus organization ‘‘B’’ with two heterochromatic segments of equal size near the centromere ‘‘C’’ with a small chromomere and two heterochromatic segments near the centromere ‘‘D’’ with three heterochromatic segments distributed in a 1:2 pattern near the centromere

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S. PRAKASH, S. R. BHAT, C. F. QUIROS, P. B. KIRTI, AND V. L. CHOPRA

‘‘E’’ with four or more heterochromatic segments ‘‘F’’ with two unequal heterochromatic segments. Based on these observations, Ro¨bbelen (1960) proposed the genetic constitution of the three basic species. B. rapa has two chromosome types, A and D, in tetrasomic and type F in hexasomic condition and the constitution AABCDDEFFF. B. nigra is tetrasomic for chromosomes D and F and has the constitution ABCDDEFF. B. oleracea is a triple tetrasomic for three chromosome types B, C, and E, with the constitution ABBCCDEEF. Venkateswarlu and Kamala (1971) arrived at a conclusion very similar to that of Ro¨bbelen (1960). They also identified six basic types of chromosomes. However, their observations regarding the type of chromosomes present in disomic or tetrasomic condition differed. According to them, the A genome has the genetic constitution AABCDDEFFF; B genome has the constitution ABCDEEFF; and C genome has the constitution ABCCDDEEF. These authors opined that basic genomes originated from loss of different sets of chromosomes from an allotetraploid (2n ¼ 20) rather than from duplication of different chromosomes, as proposed by Ro¨bbelen (1960). Generating karyotypes based on meiotic chromosome preparations rather than mitotic ones has a number of advantages although the clumping of pericentromeric heterochromatin makes the resolution of individual chromosomes difficult. Thus, combining pachytene and metaphase chromosome analysis for efficient physical mapping by FISH would be advantageous (Ziolkowski and Sadowski 2002). 6. Satellite Chromosomes and rDNA Loci. The small size of Brassica somatic chromosomes makes it difficult to identify all satellite chromosomes. In recent years, these have been identified with precision with silver staining and FISH, by detecting active nucleolar organizer regions (NORs) and nucleolar activity of ribosomal RNA (Table 2.5). Cheng and Heneen (1995), for the first time, used silver staining to detect the number and sites of nucleolar organizer regions and the number of nucleoli in Brassica species. Giemsa staining was used to characterize the satellite chromosomes. Their studies at prometaphase/early metaphase revealed that while one pair of chromosomes has secondary constriction and satellites in B. rapa, three such pairs exist in B. nigra, the last being substantiated by Hasterok et al. (2005a). The silver-stained nucleolar organizing regions were localized at the secondary constrictions of the satellite chromosomes. B. rapa revealed two NORs and two nucleoli; six NORs and six nucleoli were

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Table 2.5. Satellite chromosomes in Brassica and allied genera. Species B. nigra

No. satellite chromosomes 4

6 B. oleracea

2

B. rapa

2

B. carinata B. juncea

4 6

B. napus

4 2 4

Raphanus sativus Sinapis alba Sinapis arvensis

4 6 6

Reference Sikka 1940; Ro¨bbelen 1960; This 1990; Lan et al. 1991; Hasterok and Maluszynska 2000a Cheng and Heneen 1995; Mackowiak and Heneen 1999; Hasterok et al. 2005a Sikka 1940, Wang and Luo 1987; Olin-Fatih and Heneen 1992; Cheng et al. 1995a; Armstrong et al. 1998; Hasterok and Maluszynska 2000a; Ziolkowski and Sadowski 2002; Hasterok et al. 2005a,b Sikka 1940; Nishibayashi 1992; Olin-Fatih and Heneen 1992; Cheng and Heneen 1995; Cheng et al. 1995a; Hasterok and Maluszynska 2000a; Hasterok et al. 2005a,b; Lim et al. 2005 Kulak et al. 2002 Sikka 1940; Maluszynska and Hasterok 2005 Kulak et al. 2002 Olin-Fatih and Heneen 1992; Olin-Fatih 1994, 1996; Skarzhinskaya et al. 1998 Snowdon et al. 1997a; Hasterok and Maluszynska 2000b; Kulak et al. 2002 Mukharjee 1979 Geber and Schweizer 1988 Cheng and Heneen 1995

observed in B. nigra in early and late telophase. The two Ag-NORs in B. rapa represent one pair of active rDNA loci. Maluszynska and Heslop-Harrison (1993), Snowdon et al. (1997a), and Fukui et al. (1998), on the contrary, reported five pairs of rDNA loci following in situ hybridization. These represent both active and inactive rDNA sites while silver staining reveals only the sites with active rDNA. It appears that four pairs of these sites are inactive in nucleolus formation in B. rapa (Cheng and Heneen 1995). The observations on B. nigra having six pairs do not correspond with the earlier investigations of Sikka (1940) and Lan et al. (1991), where only four satellite chromosomes were observed. Ro¨bbelen (1960) also observed that only four chromosomes were associated with nucleoli at pachytene of meiosis in B. nigra. This et al. (1990) also assigned rDNA markers to two pairs of B. nigra

44

S. PRAKASH, S. R. BHAT, C. F. QUIROS, P. B. KIRTI, AND V. L. CHOPRA

chromosomes using B. nigra–specific rDNA RFLP markers. Closely related species Sinapis arvensis has five distinct satellited chromosome while one chromosome has a faint satellite. It is one of the two homologs that are heteromorphic to the size of satellite (Cheng and Heneen 1995). Six Ag-NORs were observed. The Ag-bands varied in size between and within pairs: two large, three intermediate, and one less distinct and small. These authors also suggested that higher number of NORs in B. nigra and Sinapis arvensis as compared to B. rapa and B. oleracea implies that the former two species have been more conservative during evolution. Brassica oleracea possesses only one pair of chromosomes exhibiting secondary constrictions and satellites as revealed by mitotic analysis (Sikka 1940; Wang and Luo 1987; Olin-Fatih and Heneen 1992; Cheng et al. 1995a). Giemsa staining also revealed one pair associated with secondary constriction and satellites (Cheng et al. 1995a). However, at pachytene of meiosis, two pairs of chromosomes are associated with nucleoli (Ro¨bbelen 1960; Kamala 1976). This fact gets support from the occurrence of four nucleoli—two major and two minor—observed by Cheng et al. (1995a). These authors also suggested that one satellited chromosome pair (no. 9) contains major active rRNA loci leading to the formation of major nucleoli and the nonsatellite pair (no. 8) carries minor active rRNA gene loci responsible for micronuclei formation. The size difference between major and minor pairs of rRNA gene loci probably suggests a reduction of most copies of rRNA genes on chromosome pair 8 during and/or after the speciation of B. oleracea. In all the three diploid species, silver staining confirmed that active rRNA genes are located at the secondary constriction of NOR chromosomes. Allopolyploid species have either the sum of or fewer satellite chromosomes than the diploids. B. carinata and B. juncea both have two pairs of satellite chromosomes (nos. 16–17 and 17–18, respectively) with distinct secondary constrictions (Kulak et al. 2002). In B. juncea, chromosome no. 16 is also a NOR-bearing type, but it lacks a distinct secondary constriction/satellite region. B. napus karyotype revealed one pair of satellited chromosomes based on C-banding (Olin-Fatih and Heneen 1992; Olin-Fatih 1994, 1996) and giemsa staining (Skarzhinskaya et al. 1998), However, after in situ hybridization with rDNA, two pairs (nos. 18–19) with distinct secondary constrictions and satellites were observed (Hasterok and Maluszynska 2000b; Snowdon et al. 2000a; Kulak et al. 2002). Both pairs of NORs are similar in size and morphology. The related species Raphanus sativus has two pairs (Mukharjee 1979) while Sinapis alba and S. arvensis carry two pairs each of NORs (Geber and Schweiger 1988; Cheng and Heneen 1995).

2. BRASSICA AND ITS CLOSE ALLIES: CYTOGENETICS AND EVOLUTION

45

The knowledge of relative physical location and the number of rDNA gene loci is relevant for construction of physical maps of chromosomes and for phylogenetic studies. Two types of ribosomal RNA genes—5S rDNA and 45S rDNA—encoding for 18S-5.8S-25S ribosomal RNAs have been studied extensively in Brassica through FISH to find their locations on chromosomes. These genes also provide excellent cytological markers for karyotype analysis, particularly where the chromosomes are very small. FISH was first used in Brassica by Maluszynska and Heslop-Harrison (1993), who investigated all the six species of U triangle to determine the number of rDNA loci. They observed that the diploid species have more than one pair of rDNA loci (B. nigra 2, B. oleracea 3, and B. rapa 5), and the derived alloploids have fewer rDNA loci than the sum of parental species (Table 2.6). However, chromosome location of hybridization sites was generally unclear. The first precise localization of 45S rDNA sequence on mitotic chromosomes was determined in B. rapa, B. oleracea, and B. napus by Snowdon et al. (1997a). Subsequently, Fukui et al. (1998) and Hasterok and Maluszynska (2000a) analyzed rDNA sites followed by 5S rDNA sites (Schrader et al. 2000; Snowdon et al. 2000a; Hasterok et al. 2001; Kulak et al. 2002; Koo et al. 2004; Hasterok et al. 2005a). Recently two comprehensive investigations on number and physical localization of rDNA loci in a large number of genera and species in Brassica coenospecies have been carried out using both rDNA probes. Ali et al. (2005) studied karyotypes of 37 species in five tribes of Brassicaceae, many of which belong to Brassica coenospecies, and also compiled corresponding data of others (Table 2.6). In the investigation of Hasterok et al. (2006), a major focus was polymorphism number and also localization of rDNA loci in many accessions of Brassica species and related genera. These investigations clearly demonstrated the efficacy of rDNA markers in identifying the chromosomes and construction of karyotypes. Wide variations in the number and localization of rDNA sites have been observed among species in these studies (Table 2.6). The karyotypes of diploid species—B. nigra, B. oleracea, and Eruca sativa—carry the minimum number of two 5S rDNA sites. The highest number of sites was reported in B. rapa, ranging from 6 (Hasterok et al. 2001) to 10 (Hasterok et al. 2005a,b, 2006; Lim et al. 2005). The lowest number of two 45S rDNA loci among the diploids was observed in B. tournefortii while the highest was in B. rapa (10). Among the allopolyploids, the number of 5S rDNA sites varied from 4 in B. carinata to a high of 14 in B. napus (Ali et al. 2005). 45S rDNA sites are in the range of 8 to 10 in B. carinata, 12 to 16 in B. juncea, and 10 to 14 in B. napus (Hasterok et al. 2006). Regarding the

46

4 6–10

4 4 (i)y

B. oxyrrhina (18) B. rapa (20)

B. tournefortii (20) B. carinata (34)

2 8–10

4 10 (i)y

4 4 2 þ 2 minor 4 þ 2 minor (i)y (two 5S rDNA sites on the same chromosome)

B. fruticulosa (16) B. oleracea (18)

4–6

with 45S rDNA

2 (t)

z

with 5S rDNA

B. nigra (16)

Taxon (2 n)

2 —

2 4–8

2 —

—

with 5S and 45S rDNA

Number of Chromosomes

Reference Maluszynska and Heslop-Harrison 1993 (only 45S rDNA); Fukui et al. 1998 (only 45S rDNA); Hasterok and Maluszynska 2000a (only 45S rDNA); Hasterok et al. 2001; Ali et al. 2005; Hasterok et al. 2006 Hasterok et al. 2006 Maluszynska and Heslop-Harrison 1993 (only 45S rDNA); Snowdon et al. 1997a (only 45S rDNA); Armstrong et al. 1998; Fukui et al. 1998 (only 45S rDNA); Hasterok and Maluszynska 2000a (only 45S rDNA); Hasterok et al. 2001; Ziolkowski and Sadowski 2002; Snowdon et al. 2002; Ali et al. 2005; Hasterok et al. 2005a, 2006 Hasterok et al. 2006 Maluszynska and Heslop-Harrison 1993 (only 45S rDNA); Snowdon et al. 1997a (only 45S rDNA); Fukui et al. 1998 (only 45S rDNA), Hasterok and Maluszynska 2000a (only 45S rDNA); Hasterok et al. 2001; Snowdon et al. 2002; Koo et al. 2004; Ali et al. 2005; Hasterok et al. 2005a, 2006; Lim et al. 2005 Hasterok et al. 2006 Maluszynska and Heslop-Harrison 1993 (only 45S rDNA); Hasterok and Maluszynska 2000c (only 45S rDNA); Hasterok et al. 2001; Kulak et al. 2002; Ali et al. 2005; Hasterok et al. 2006

Table 2.6. Number and position of 5S and 45S rDNA sites in Brassica and allied genera.

47

8 6 6–7 8 þ 4 minor 4 6 5–6

4 (i)y 6 2 (i)y 4 (i)y 4 2 (i)y 4–5 (i)y

8 4 (t)

8 4 (t)z 6 22

Diplotaxis siifolia (20) Diplotaxis muralis (42) Eruca sativa (22) Erucastrum gallicum (30 ) Hirschfeldia incana (16) Moricandia arvensis (28) Raphanus sativus (18)

R. sativus (tetraploid) (36) Sinapis alba (24)

S. alba (tetraploid) (48) Sinapis arvensis (18) Sinapidendron frutescens (20) Orychophragmus violaceus (24)

4 2 2 8

4 2

2 — 3

2 2 2

5–8

6

Maluszynska and Heslop-Harrison 1993 (only 45S rDNA); Hasterok and Maluszynska 2000c (only 45S rDNA); Hasterok et al. 2001; Kulak et al. 2002; Ali et al. 2005; Hasterok et al. 2006 Maluszynska and Heslop-Harrison 1993 (only 45S rDNA); Snowdon et al. 1997a (only 45S rDNA); Kamisugi et al. 1998; Hasterok and Maluszynska 2000c (only 45S rDNA); Schrader et al. 2000; Snowdon et al. 2000a; Hasterok et al. 2001; Kulak et al. 2002; Snowdon et al. 2002; Ali et al. 2005; Hasterok et al. 2006 Ali et al. 2005 Hasterok et al. 2006 Ali et al. 2005; Hasterok et al. 2006 Ali et al. 2005 Ali et al. 2005 Ali et al. 2005 Schrader et al. 2000; Ali et al. 2005; Hasterok et al. 2006 Hasterok et al. 2006 Schrader et al. 2000; Ali et al. 2005; Hasterok et al. 2006 Hasterok et al. 2006 Ali et al. 2005; Hasterok et al. 2006 Ali et al. 2005 Li et al. 2005

y

z

t ¼ terminal/subterminal position of 5S rDNA. i ¼ interstitial position of 5S rDNA exclusively; interstitial and terminal position of 45S rDNA. Source: Compiled from Ali et al. 2005; Hasterok et al. 2006.

16 6 6 8

8 6–8

10–14 (i)y

9–14 (two 5S rDNA sites on the same chromosome)

B. napus (38)

12–16 (i)y

8–10

B. juncea (36)

48

S. PRAKASH, S. R. BHAT, C. F. QUIROS, P. B. KIRTI, AND V. L. CHOPRA

location of these rDNA sites, B. rapa has 4 25S rDNA loci near the centromere of metacentric chromosomes 1, 4, 5, and 7. Chromosome 2 bears NOR and contains the fifth largest 25S rDNA locus extending over NOR and satellite. Chromosome 7 has a large 25S locus located interstitially and colocalized with a large 5S rDNA locus. Short arms of chromosome 2 and 10, the largest and smallest acrocentric chromosomes, respectively, in B. rapa genome have two more 5S loci (Snowdon et al. 2002). Koo et al. (2004) and Hasterok et al (2005a) also observed the same number of 45S and 5S rDNA loci at the same locations. Of these 10 rDNA loci, only 2 are active, distributed on the secondary constriction of chromosome 10 (Hasterok and Maluszynska 2000a). Koo et al. (2004) studied pachytene bivalents and observed 5S rDNA loci on pericentromeric region of short arm of chromosomes 2 and 10 and the long arm of chromosome 7. The long arm of chromosome 7 exhibited another 5S rDNA site, which was not detected in mitotic metaphase. These authors believe that two closely linked 5S rDNA loci could not be detected in earlier investigations because of lower resolution of FISH on mitotic chromosomes. Localization of 45S rDNA loci was revealed on pericentromeric regions of the short arm of chromosomes 1, 2, 4, and 5 and the long arm of chromosome 7. Brassica oleracea genome has two 18S-5.8S-25S rDNA sites subtelomerically on the short arms of two satellited acrocentric chromosomes (nos. 4 and 7). The third one occurs adjacent to the centromere on the short arm of chromosome 2, which is submetacentric. On the long arm of this chromosome, 5S rDNA sequences are located with closely adjacent major and minor loci (Armstrong et al. 1998). These results match those of Hasterok et al. (2001, 2005a) and Snowdon et al. (2002), who observed 5S rDNA genes in two closely adjacent loci on the long arm of a single large submetacentric chromosome 4. Two acrocentric satellite-possessing chromosomes (nos. 2 and 7) have 25S loci at the terminal ends of their short arm, which extends over the satellite. A novel 5S rDNA locus was also detected by Ziolkowski and Sadowski (2002). B. nigra has three pairs of 25S loci; one pair is located on the short arm of chromosome 6 and two pairs are located at the secondary constriction and satellite of chromosome pairs 7 and 8. Only these two pairs of loci are transcriptionally active (Hasterok and Maluszynska 2000c). The derived alloploids have fewer rDNA loci than the sum of their parents. B. carinata has four 5S and eight 25S rDNA sites, and no chromosome carries both the genes closely linked or colocalized. Ten sites of 5S and 16 sites of 25S rDNA are observed in B. juncea.

2. BRASSICA AND ITS CLOSE ALLIES: CYTOGENETICS AND EVOLUTION

49

Chromosomes 4, 10, and 16 have colocalized both the gene sites (Kulak et al. 2002). Employing imaging methods in combination with FISH, Kamisugi et al. (1998) detected 25S/18S rDNA loci in the centromeric and distal regions of seven chromosomes in B. napus. While eight 5S rDNA loci were observed on five chromosomes mainly in the centromeric regions, two chromosomes carried both 25S/18S and 5S rDNA loci in close proximity. Regarding their localization, according to Snowdon et al. (1997a, 2000a), the largest site covers the satellite and short arm of the largest NOR-carrying chromosome. The second largest is located on a telomeric NOR-like structure on the short arm of a large subtelocentric pair, and the smallest locus is at the telomere on the short arm of a smaller submetacentric chromosome. The three other loci are localized at or near the centromeres of metacentric chromosomes. About the origin of these rDNA sites carrying chromosomes, Snowdon et al. (1997a) inferred that the two largest noncentromeric signal blocks and the NOR-carrying chromosomes closely resemble those of B. rapa and B. oleracea. Similarly, the three largest centromerically located loci in B. napus match to those of B. rapa. Schrader et al. (2000) indicated that in B. napus, one of the pair with 5S rDNA gene sites belongs to B. oleracea. Additionally, two submetacentric chromosomes having two closely adjacent 5S DNA clusters belong to B. oleracea. The other four pairs probably derived from the B. rapa progenitor. A comparison of chromosome sets of B. napus, B. oleracea and B. rapa revealed that B. napus chromosomes carrying rDNA loci could be matched with those of constituent parents (Snowdon et al. 2002). Chromosomes possessing rDNA loci could be identified based on size and centromere position. The chromosomes belonging to A and C genomes could clearly be distinguished with minor discrepancies. In general, these observations closely correspond to those of Kamisugi et al. (1998). Maluszynska and Hasterok (2005), using two-color GISH, successfully discriminated partaking genomes in B. juncea and assigned chromosomes to A and B genomes. Molecular analysis also indicated that in allopolyploids, B. nigra rRNA genes are dominant over those of B. rapa, which are in turn dominant over B. oleracea (Chen and Pickard 1997; Ge and Li 2007). However, according to Hasterok and Maluszynska (2000c), the number of Ag NORs in the alloploid species is equal to the sum of active NORs in diploid parental species, clearly indicating an absence of nuclear dominance in root meristematic cells. Views have been proposed regarding the evolutionary loss of rDNA in B. napus. It might be the result of genome organization due to translocation events between A and C genome chromosomes (Sharpe

50

S. PRAKASH, S. R. BHAT, C. F. QUIROS, P. B. KIRTI, AND V. L. CHOPRA

et al. 1995). However, B. napus is of very recent origin, and we also have the conflicting views that the partaking A and C genomes are unaltered to a large extent (Parkin et al. 1995). In fact, Delseny et al. (1990) have reported that rDNA-carrying chromosomes of B. oleracea have not undergone any major structural changes since the evolution of B. napus. Earlier, Bennet and Smith (1991) suggested that there has been a large reduction in copy number of rDNA in present-day B. napus as compared to ancestral forms, which is due to a reduction in B. oleracea–type rDNA in existing B. napus forms from a total copy number of 1,500 to one of 800, while B. rapa–type rDNA is unaltered. Maluszynska and HeslopHarrison (1993) are of the view that a C-genome locus has been lost in B. napus due to reduction in B. oleracea–type rDNA. However, Snowdon et al. (1997a, 2000a) believe that both ancestral B. oleracea loci are still present, with reduced rDNA copy numbers, as suggested earlier by Bennet and Smith (1991). They also proposed that the smallest and relatively insignificant rDNA locus from B. rapa is absent in B. napus. On the whole, since substantial C-genome rDNA has been lost, it appears that A-genome rDNA is of greater genetic importance than C-genome rDNA in B. napus. In the species belonging to related genera Sinapis alba and Raphanus sativus, Schrader et al. (2000) detected 5S and 25S rRNA genes using double FISH. Sinapis alba possesses four 5S rDNA loci on the short arm of two submetacentric chromosome pairs and six 25S rDNA loci on the short arm of six chromosomes. In one of the chromosome pairs, both rDNA genes were closely linked on the short arm. These results were confirmed by Ali et al. (2005). Schrader et al. (2000) suggested that of six sites of 25S rDNA genes in S. alba, three represent active and three inactive loci. Raphanus sativus displays four sites each of 5S and 25S rDNA gene loci. Both rDNA genes were localized distally on the short arm (25S) and near the centromere on the long arm (25S) on one of the longest chromosome pair. The other 5S or 25S rDNA genes were separately located on the long arm of two chromosome pairs (Scharder et al. 2000; Ali et al. 2005). Investigations on 15 species in Brassica and allied genera (Table 2.6) reveal that 5S rDNA loci occupied interestitial positions in four species—Eruca sativa, Erucastrum gallicum, B. carinata, and Raphanus sativus—and terminal/subterminal and also interestitial positions in other species (Ali et al 2005). Hirschfeldia incana, Sinapis arvensis, S. alba, Diplotaxis siifolia, Sinapidendron frutescens, Eruca sativa, and Raphanus sativus have one chromosome pair carrying both loci. Sinapis arvensis and Raphanus sativus have these on opposite arms of chromosomes; in others these are located adjacently on the same arm.

2. BRASSICA AND ITS CLOSE ALLIES: CYTOGENETICS AND EVOLUTION

51

A comparative analysis clearly revealed the occurrence of polymorphism in number and chromosomal distribution of rDNA loci among different ecotypes of a species and their population (Hasterok et al. 2006). Inter- and/or intravarietal polymorphism was evident in B. oleracea, B. rapa, B. carinata, B. juncea, B. napus, and Raphanus sativus. It was also observed that Brassica species carrying A genome—B. rapa, B. juncea, and B. napus—are highly polymorphic and contain high numbers of rDNA sites (Hasterok et al. 2001).

7. Archetype and Evolution of Genomes. Several investigators (Thompson 1956; Ro¨bbelen 1960; Prakash and Hinata 1980) have proposed that basic diploid species are polyploids that evolved from a common ancestor in the ancient past. While evidence are presented that this archetype was a smaller genome, views on the genetic constitution of the ancestral genome are variable, proposing the ancestral basic numbers x ¼ 37: Catcheside (1934, 1937), basing his conclusion on secondary chromosome associations in B. napus and B. oleracea, suggested x ¼ 6 as the basic number. He also believed that the primitive haploid number in Brassicaceae is x ¼ 7 from which x ¼ 6 arose by fusion of two chromosomes. Alam (1936) and Haga (1938) supported Catcheside’s views after observing secondary pairing in the three basic diploid species. Sikka (1940) proposed a basic number x ¼ 5, from the study of secondary bivalent associations in three species—B. monensis (syn. Coincya monensis, 2n ¼ 24), B. sinapistrium (syn. Sinapis arvensis, 2n ¼ 18), and B. nigra—and suggested the evolution in Brassica toward tetraploidy, citing the series 2n ¼ 30, 60, 90, and 120 in the genus Crambe, all multiples of x ¼ 5. However, the pachytene chromosome analysis in diploid species revealing six basic types of chromosomes led Ro¨bbelen (1960) to suggest that the archetype had x ¼ 6 as the basic constitution. This is supported by meiotic chromosome pairing in haploids of B. nigra and B. tournefortii (Prakash 1974a,b) and substantiated by GISH in trigenomic ABC hybrids (Ge and Li 2007). An entirely different number of x ¼ 3 was proposed by Hussein and Abobakr (1976) based on secondary bivalent associations in B. oleracea. This view found support from: (1) Chen and Heneen (1991) and Cheng and Heneen (1995), who observed three pairs of satellited chromosomes with active NORs in B. nigra and S. arvensis; (2) three pairs of chromosomes carrying 25S rDNA gene loci in B. nigra (Fukui et al. 1998), B. oleracea (Maluszynska and Heslop-Harrison 1993; Snowdon et al. 1997a; Armstrong et al. 1998; Ali et al. 2005; Hasterok et al. 2006), S. alba (Schrader et al. 2000), and S. arvensis (Ali et al. 2005);

52

S. PRAKASH, S. R. BHAT, C. F. QUIROS, P. B. KIRTI, AND V. L. CHOPRA

and (3) three copies of the gene encoding acyl-CoA-binding protein in B. rapa and B. oleracea (Hills et al. 1994). It is inferred that the number of three chromosome pairs carrying the 25S rDNA gene is basic for the family Brassicaceae. The earlier view was that the three basic diploid species evolved from a common archetype following duplication of whole chromosomes (i.e., aneuploidy) accompanied by differentiation following structural changes. The mapping data, in contrast, clearly discounts the role of polysomy or duplication of whole chromosomes (Quiros 1999). Recent information emerging from use of molecular markers firmly disproves the theory of monophyletic origin and instead suggests a biphyletic origin of the diploid species. It concludes that B. oleracea/ B. rapa originated from one archetype while B. nigra originated from the other (Song et al. 1990; Warwick and Black 1991; Pradhan et al. 1992). Cytogenetical investigations preceeded in predicting this genetic divergence between B. nigra and B. oleracea/B. rapa based on chromosome pairing in hybrids (Mizushima 1950a; Prakash and Hinata 1980). Subsequently, molecular analysis has been very revealing. The first evidence came from nuclear RFLPs by Song et al. (1988a), which was substantiated by other investigations. Information from nuclear, chloroplast, and mitochondrial DNA RFLPs has established that the primitive genome diversified into two lineages and all the taxa in subtribe Brassicinae fall in these two lineages (Warwick and Black 1991; Pradhan et al. 1992). This view also gets support from a comparative study of molecular markers (Lagercrantz 1998) and rDNA intergenic spacer (Bhatia et al. 1996). The evolutionary divergence is also reflected in their cytoplasm (Palmer 1988; Warwick and Black 1991; Pradhan et al. 1992). B. oleracea and B. rapa cytoplasms are closer to each other than either is to B. nigra (Palmer 1988). Evidence also indicates that the A genome was derived in the distant past from an already existing C genome, as these two genomes have extensive genomic regions of conserved homology (Slocum 1989). Many investigations during the past 15 years have dealt with comparative mapping of various genera and species, which throws light on the constitution of archetype and history of polyploidy. Truco et al. (1996) suggested a range of x ¼ 5 to 7 of this ancestral archetype, which was further narrowed down to x ¼ 4 by Lagercrantz (1998). Evidence cited in support was that several species of the related subtribe Lepidieae also have n ¼ 4 chromosome number, the lowest in Brassicaceae family (Mulligan 1964). Henry et al. (2006), based on recent genome sequencing data, also support n ¼ 4 as the basal

2. BRASSICA AND ITS CLOSE ALLIES: CYTOGENETICS AND EVOLUTION

53

genomic number of the family, followed by tetraploidization before the separation of the Brassica and Arabidopsis lineages. An interesting hypothesis proposed by Lagercrantz and Lydiate (1996), Lagercrantz (1998), and O’Neil and Bancroft (2000) envisages that (i) Arabidopsis shares common ancestry with Brassica crop species, and (ii) three ancestral species with x ¼ 5 and whose genomes were similar to Arabidopsis genome gave rise to a hexaploid following hybridization between them. This was the ancestral archetype of A, B, and C genomes from which the basic genomes evolved through reduction in chromosome number by extensive chromosome fusion. This view, known as the triplication theory, gets support from the fact that some loci are triplicated as detected by molecular markers (Cavell et al. 1998; O’Neil and Bancroft 2000; Parkin et al. 2002, 2003, 2005; Rana et al. 2004; Lysak et al. 2005, 2007; Park et al. 2005; Yang et al. 2005; Lim et al. 2006; Matthew and Lydiate 2006; Nelson and Lydiate 2006; Ziolkowski et al. 2006; Yang et al. 2006) and also that diploid Brassica genomes contain approximately three times the DNA of Arabidopsis genome (Arumugnathan and Earle 1991). The event of hexaploidy occurred around 7.9 to 14.6 million years ago (Lysak et al. 2005). However, the occurrence of a large proportion of heterochromatin, repetitive DNA (Gupta et al. 1990, 1992; Iwabuchi et al. 1991), transposable elements (TE) (Zhang and Wessler 2004; Gao et al. 2005; Lim et al. 2007), and the ancestral role of Arabidopsis would argue against this hypothesis. Lukens et al. (2004) also found no strong evidence of the role of the ancestral hexaploid genome. Furthermore, considering that the ancestral species to the Brassicaceae was a tetraploid of 2n ¼ 4x ¼ 16 (Henry et al. 2006), it already explains the origin of species such as B. nigra, also with 2n ¼ 16, without the need to invoke another round of polyploidization or hexaploidy. Another hypothesis referred to as cyclic amphiploidy was proposed by Truco et al. (1996) and Quiros (1999), which suggests that hybridizations occurred among different species with x ¼ 4 and/or x ¼ 5 chromosome constitution. These species were derived from an ancestral Brassica archetype genome with x ¼ 4 or 5. Each pair of species crosses gave rise to A, B, and C genomes and are partial allopolyploids (Fig. 2.3). Also sequidiploidy resulting from hybridization of allotetraploids to ancestral diploids could have been originated genomes of various chromosome numbers. Structural chromosomal modifications, mostly reciprocal translocations, played a pivotal role initially in the origin of these genomes (Quiros 1999). Recent evidence also supports further and constant genome modification by insertion and movement of TE (Zhang and Wessler 2004; Gao et al. 2005;

54

S. PRAKASH, S. R. BHAT, C. F. QUIROS, P. B. KIRTI, AND V. L. CHOPRA

Ancestral genome x =4, 5 ?

Chromosomal changes Geographic isolation Derived genomes x=4, 5

Chromosomal changes Z1

TE Z2

1st cycle of allopolyploidy

Diploid species genomes

Z4

Z3

Zn

Aneuploidy B

A

C

Structural changes 2nd cycle of allopolyploidy

Cultivated allopolyploids

BC

AB

AC

Fig. 2.3. Cyclic amphidiploidy and the origin of Brassica diploid and allopolyloid species solid and broken lines represent female and male parents respectively in allopolyploids. (Source: After Quiros 1999.)

Lim et al. 2007). Surprisingly, the genome size of these basic species has remained practically unaltered in spite of changes in chromosome numbers and structure. Based on marker arrangement conservation, Truco et al. (1996) proposed a model of genome evolution and phylogenetic relationships among the chromosomes of the three basic species considering two assumptions: that (1) A and C genomes are closely related, and possibly C genome is the predecessor of A genome; and (2) the genus Brassica is of biphletic origin. It envisages that the ancient genome possessed at least five and no more than seven chromosomes. B and/or C genome chromosomes evolved from six ancestral chromosomes (W1 to W6) (Fig. 2.4). C genome chromosomes also gave rise to A genome chromosomes. Two intermediate chromosomes Bx and Cx originated from W1. Bx produced B1, B2, B4, and B8 chromosomes and the Cx

2. BRASSICA AND ITS CLOSE ALLIES: CYTOGENETICS AND EVOLUTION

A10

A5

C5

55

A4

B6

C1

B3

A1

B5 W3

C4 A6

C7

W5

W4

C×

W2

A3

C3

W1

W6

C6

A7

C8

B×

B1

A8 C9

B8 B7

B4

A9

B2 C2

A2

Fig. 2.4. Hypothetical ancestral genome of six chromosomes (W1 to W6) originating specific A, B and C genome chromosomes deduced by homoeologous relationships. Bx and Cx are intermediate chromosomes. Broken lines indicate tentative homologies. (Source: From Truco et al. 1996.)

chromosome gave rise to A7. Chromosomes Bx and C1 were similar in their genetic content. Chromosomes B7 and C9 might have originated from W6 or independently, one from W6 and other from a seventh ancestral chromosome, W7. These two chromosomes, B7 and C9, do not share homology with any other group. In spite of their biphyletic origin, the three basic genomes still share regions of homology, as determined by Truco et al. (1996), expressed in cM by adding the distance of chromosome segments sharing homology between the two genomes following the comparison of linkage maps of these species. The lowest homology is between A and B genomes, which share 92.7 and 219.5 cM of their genomes respectively and results in up to six bivalents in hybrids between them (Prakash 1973a,b). Homology between B and C genomes is intermediate with 223 and 365.7 cM respectively and form up to four bivalents between them (Mizushima

56

S. PRAKASH, S. R. BHAT, C. F. QUIROS, P. B. KIRTI, AND V. L. CHOPRA

1950a; Song et al.1993). The highest homology is observed between A and C genomes where they share 337.2 and 487.2 cM respectively and form up to nine bivalents (Olsson 1960b). ISSR data also reflected these relationships, as observed by Liu and Wang (2006). The average genetic distance between B. rapa and B. oleracea is 0.499, indicating close homology; between B. rapa and B. nigra, 0.528; and between B. oleracea and B. nigra, 0.615 showing clearly the divergence between A/C and B genomes. In fact, the genomic contents of A and C genomes are equivalent, and rearrangements are the cause of difference in their chromosome number (Parkin et al. 2003). To summarize, basic Brassica genomes evolved and differentiated from an originally smaller genome. Chromosome arrangements due to homoeologous recombination and hybridizations were the major factors in their stabilization. These three species are, in fact, secondary polyploids with regions of shared ancestry. As expected, duplications are widespread in these genomes.

III. GENOME MANIPULATION A. Resyntheses of Natural Allopolyploid Brassica spp. Soon after the proposals of Morinaga (1928–1934) on genome analysis, Nagaharu U (1935) presented the cytogenetical relationships among different Brassica crop species. He is also credited with substantiating it by experimental synthesis of B. napus. A little later Ramanujam and Srinivasachar (1943) successfully produced B. juncea. Simultaneously, Frandsen (1943, 1947) synthesized all the three allopolyploid species. These early academic reports were primarily aimed at verifying Morinaga’s proposals. Subsequently, the activities shifted to enhancement of genetic variability of breeding value utilizing extensive genetic and phenotypic range of variation available in constituent parents. It is now believed that intensive breeding, particularly in B. napus, which is a relatively younger species, and also in B. juncea, has led to a narrowing of the genetic base. In contrast, experimentally resynthesized strains have considerably broadened the gene pool for use by the breeders. These synthetics are also valuable in heterosis breeding programs; several investigations report high heterosis in crosses between synthetics and crop cultivars (Pradhan et al. 1993; Jain et al 1994; Girke et al. 1999; Seyis et al. 2003). Literature on sexually and parasexually produced versions of naturally occurring Brassica species is summarized in Tables 2.7 and 2.8.

2. BRASSICA AND ITS CLOSE ALLIES: CYTOGENETICS AND EVOLUTION

57

Table 2.7. Major investigations on artificial synthesis of natural allopolyploid species B. carinata, B. juncea, and B. napus through sexual hybridization. Species

Reference

B. carinata (B. nigra  B. oleracea and reciprocals) B. juncea (B. rapa  B. nigra and reciprocals)

B. napus (B. rapa  B. oleracea and reciprocals) Oil rape

Forage rape Rutabaga

Heading form

Frandsen 1943; Mizushima 1950b; Pearson 1972; Prakash et al. 1984; Song et al. 1993 Frandsen 1943; Ramanujam and Srinivasachar 1943; Olsson 1960a; Prakash 1973a,b; Campbell et al. 1990, 1991; Song et al. 1993; Srivastava et al. 2001, 2004; Se´guin-Swartz et al. 2004

U 1935; Karpechenko and Bogdanova 1937; Frandsen 1947; Rudorf 1950; Hoffmann and Peters 1958; Olsson 1960b; Gland 1982; Prakash and Raut 1983; Chen et al. 1988a,b; Chen and Hennen 1989; Akbar 1989; Hossain et al. 1990; Mithen and Magrath 1992; Song et al. 1993; Ozminkowski and Jourdan 1994a,b; Beschorner et al. 1995; Heath and Earle 1996; Girke et al. 1999; Lu et al. 2001; Rahman et al. 2001; Zhang et al. 2002; Happastadius et al. 2003; Luhs et al. 2003; Seyis et al. 2003; Niu et al. 2004; Zhang et al. 2004; Abel et al. 2005; Rahman 2005; Zhou et al. 2007; Wen et al. 2008 Hosoda 1950, 1953, 1961; Feng 1955; Sarashima 1967, 1973; Hosoda et al. 1969; Nishi et al. 1970 Olsson et al. 1955; Olsson 1960b; Hosoda et al. 1963, 1969; Namai and Hosoda 1967, 1968; Kato et al. 1968 Shinohara and Kanno 1961

Table 2.8. Major papers on synthesis of natural allopolyploid species through protoplast fusion. Brassica species carinata juncea napus

Reference Narasimhulu et al. 1992; Jourdan and Salazar 1993 Campbell et al. 1990, 1991; Se´guin-Swartz et al. 2004; Bhat et al. unpubl. Schenck and Ro¨bbelen 1982; Sundberg and Glimelius 1986; Taguchi and Kameya 1986; Robertson et al. 1987; Sundberg et al. 1987; Terada et al. 1987; Rosen et al. 1988; Jourdan et al. 1889b; Hossain and Asahira 1992; Yamagishi et al. 1992; Ozminkowski and Jourdan 1993, 1994a,b; Hansen and Earle 1994; Heath and Earle 1995, 1996, 1997; Ren et al. 2000; Lian and Lim 2001

58

S. PRAKASH, S. R. BHAT, C. F. QUIROS, P. B. KIRTI, AND V. L. CHOPRA

The objectives in these allopolyploid syntheses vary, from purely academic (e.g., Song et al. 1993; Srivastava et al. 2001, 2004), to developing agricultural forms that include early and productive B. carinata forms for Indian conditions (Prakash et al. 1984), highseed-yielding B. juncea (Olsson 1960a; Prakash 1973a), early-maturing B. napus suitable for the Indian subcontinent (Prakash and Raut 1983; Akbar 1989), productive oil seed B. napus (Olsson 1960b; Seyis et al. 2003), fodder forms of B. napus (Namai and Hosoda 1967, 1968; Ellerstro¨m and Sjo¨din 1973), root-forming sweeds or rutabagas (Olsson et al. 1955; Kato et al. 1968; Namai and Hosoda 1968), and a new headforming vegetable form (Shinohara and Kanno 1961; Takeda 1986). A major objective in B. napus syntheses in recent years has been the modification of oil and meal quality (Lu et al. 2001; Lu¨hs et al. 2003; Seyis et al. 2005) and incorporation of yellow seed coat color (Shirzadegan and Ro¨bbelen 1985; Liu and Gao 1987; Chen et al. 1988; Tang et al. 1997; Meng et al. 1998; Baetzel et al. 1999; Rahman 2001; Wen et al. 2008). Initially these studies were carried out chiefly in Japan, Sweden, Germany, and India, and later in other countries. In fact, resynthesis has been widely attempted for improvement of B. napus (Olsson and Ellerstro¨m 1980; Chen and Hennen 1989b; Lu¨hs et al. 2002; Friedt et al. 2003). Synthetic alloploids of all three species have been obtained through sexual hybridizations from reciprocal crosses. In some combinations, hybrids are difficult to obtain because of postfertilization barriers, which are generally overcome by sequential ovary culture or embryo rescue. Failure of normal endosperm development following fertilization is the major cause for hybrid embryo abortion (Inomata 1976; Wojciechowski 1985). Highly significant influence of maternal genotype on crossability, particularly in the synthesis of B. napus, has been observed (Diederichen and Sacrista´n 1994; Lu et al. 2001; Abel et al. 2005). Similarily, the role of cytoplasm has also been implicated in the success of hybridization (Song et al. 1993). It is observed that hybrids with the same cytoplasm as natural alloploids are easier to synthesize than others (Song et al. 1993). Synthetic B. juncea with B. nigra cytoplasm and B. napus with B. oleracea cytoplasm are difficult to obtain. In general, synthesis of B. juncea and B. carinata is easier than synthesis of B. napus (Prakash and Chopra 1991; Song et al. 1993; Stewart 2004). However, in vitro techniques, such as embryo and ovary culture, have made possible the generation of a large number of synthetic alloploids of choice. Protoplast fusion technology for Brassica became available in 1980s and enabled synthesis of somatic hybrids of desired combinations. Besides

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59

enlarging nuclear variability, this methodology generates novel combinations of cytoplasmic organelles and has received considerable attention in recent years. Synthesis of somatic hybrids of B. napus by Schenck and Ro¨bbelen (1982) was the earliest success. Most reports on somatic hybridization relate to synthesis of B. napus. Ozminkowski and Jourdan (1994a,b) and Heath and Earle (1996) reconstructed B. napus both sexually and following somatic cell fusion. B. napus allohaploids have also been synthesized by fusing pollen protoplasts of B. oleracea var. italica and haploid mesophyll protoplasts of B. rapa; Fan et al. (2007) present the first report about a hybrid formation between two haploid protoplasts. Hybrids could be obtained faster through somatic fusion because of avoidance of chromosome doubling in F1 sexual hybrids for restoring fertility. Somatic hybrids have also been obtained involving B. carinata and B. juncea (Table 2.8). These investigations on somatic hybridizations represent significant breakthroughs in interspecific hybridizations. Synthetics obtained following sexual hybridization and chromosome doubling have the sum of the parental chromosome number. The somatic hybrids also have, in general, these summations. However, in some somatically produced B. napus, there were deviations, where the plants possessed variable chromosome number ranging from 33 to 57 (Table 2.9). These resulted from triple fusions, such as one B. oleracea and two B. rapa protoplasts (2n ¼ 58) or vice versa, resulting in digenomic hexaploid AAAAACC (2n ¼ 58) and AACCCC (2n ¼ 56) plants (Terada et al. 1987; Heath and Earle 1996). Aneuploids with somatic chromosome number 33, 49, 54, 57 were also recorded. These probably originated by chromosome elimination during regeneration and subsequent development of plants. Somatic hybrid plants of B. carinata had the normal chromosome number of 34 (Narasimhulu et al. 1992; Jourdan and Salazar 1993). Campbell (1993), Se´guin-Swartz et al. (2004), and Bhat et al. (unpublished) also observed the normal somatic chromosome number in B. juncea somatic hybrid plants (Table 2.9). Meiosis in the synthetic allopolyploids obtained from either sexual or somatic hybridizations was disturbed in early generations, primarily due to the preponderance of univalents and multivalents. However, it was more normal in somatic hybrids. Up to four quadrivalents in B. carinata (Mizushima 1950b; Narasimhulu et al. 1992) and up to three quadrivalents and two trivalents occurred in B. napus (Sarashima 1973; Prakash and Raut 1983; Wen et al. 2008). Higher associations were rare or absent in B. juncea (Olsson 1960a; Prakash 1973a). The number of multivalents and univalents decreased with the

60

B. napus 12 4 10 23

5

B. juncea 3

38, 36, 54 38 38, 33, 36, 49, 56, 57 38, 50–68

36, 18 II–16 II þ 4 I

36, 18 II

34, 56

34, 17 II, 11 II þ 3 IV

B. carinata 74

64

Chromosome no. pairing at M1

No. somatic hybrids obtained

Rapa (2) Nig (1) Nig (3) Rec (2)

Ole (5) Nig (18) Novel (1)

Ole (1) Nig (3)

CP

Rapa (5) Ole (6) ND (12)

Rapa (3) Nig (2)

Rapa (3)

Ole (6) Nig (18)

Ole (1) Nig (2) Rec (1)

MT

Organelle genome composition

6–70

4–98

36–87

Pollen

very poor

very poor

19

Seed

Fertility (%)

Table 2.9. Summary of results of somatic hybrids of B. carinata, B. juncea, and B. napus.

Schenck and Ro¨bbelen 1982 Taguchi and Kameya 1986 Terada et al. 1987 Sundberg et al. 1987

Bhat et al. unpubl.

Campbell 1993

Jourdan and Salazar 1993

Narasimhulu et al. 1992

Reference

61

38 38, 56, 58 Rapa (30) Mix (9) Rapa (38) Rapa (8) Ole (16)

Rapa (3) Ole (2) 38, 58, aneuploid also Rec (12) Rapa (7) Rapa (1) Rec (10) 38,58

36–38 38, 19 II–18 II þ 2 I

z

ND ¼ not determined. Rec ¼ recombined. Number in parentheses denotes the number of plants.

38 24

109 39

72

11

34

1 5

Rapa (12) Ole (10) Rec (2)

Ole (34) Rapa (5) Ole (6)

Ole (3)

NDz

0–97

fertile –—

ND

good

low – high

sterile very poor

Cardi and Earle 1997

Heath and Earle 1997

Ozminkowski and Jourdan 1993, 1994a,b Heath and Earle 1995 Heath and Earle 1996

Jourdan et al. 1989 Sundberg and Glimelius 1991a

Robertson et al. 1987 Rosen et al. 1988

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S. PRAKASH, S. R. BHAT, C. F. QUIROS, P. B. KIRTI, AND V. L. CHOPRA

advancing generations, and meiotic stabilization with regular bivalent formation was achieved by the amphidiploid 3 (A3) generation. Sexually obtained synthetics had reduced pollen and seed fertility in early generations, sometimes as low as 6% in B. juncea (Olsson 1960a). With stabilization of meiosis and selection, fertility improved considerably. By the A5 generation, the attained fertility was much higher. A7 generation plants had fertility comparable to naturally occurring forms (Table 2.10). Somatic hybrids also had very low pollen fertility and seed set. Pollen was ineffective in producing seeds on selfing or on pollinations to natural forms of B. carinata (Jourdan and Salazar 1993). Similar observations were recorded for B. napus by Rosen et al. (1988), Sundberg et al. (1987), and Heath and Earle (1996, 1997). Since the plants had more or less regular meiosis, the reasons for a high degree of sterility are unknown. The organellar constitution of somatic hybrids does not follow any pattern; all possible combinations of mitochondria and chloroplast genomes are observed in addition to frequent intergenomic mitochondrial recombination. A majority of B. napus somatic hybrids contain B. rapa chloroplast; some have both chloroplast types (heteroplastidic); and only a few have B. oleracea chloroplasts. They contain mostly B. rapa and recombinant mitochondria. Some plants also have a mix of mitachondria and chloroplast genomes of B. rapa and B. oleracea. Most B. carinata plants contain both chloroplast and mitochondrial genomes from B. nigra, but some combine these from both the parents. A similar phenomenon is observed with in B. juncea, where combinations of chloroplast and mitchondria parental genomes have been obtained as shown in Table 2.9 (Bhat et al. unpublished). B. Agronomic Potential of Synthetics Attempts to synthesize allopolyploids and use them either directly or as useful germplasm have been made in several countries. The general observation has been that synthetics do not compete for productivity with cultivated forms but are a rich reservoir of diversity, as revealed by allozyme and molecular marker studies (Chen et al. 1989; Song et al. 1993; Becker et al. 1995; Seyis et al. 2003; Srivastava et al. 2004; Liu and Yang 2004). Their value is particularly relevant for such traits as productivity, disease resistance, and quality parameters and are utilized in backcross programs to introgress these novel characters. Hybridization between synthetics and elite breeding lines have resulted in many high-yielding cultivars because of high genetic distance between them.

63

— 17 II þ 4 IV þ 9 II

17 II–1 4 II þ 6 I

—

18 II mostly 16 II þ 4 I–18 II

—

19 II 2 IV þ 15 II–19 II 19 II mostly

19 II mostly

17 II16 II þ 2 I 17 II þ 4 IV þ 9 II

17 II þ 4 IV þ 9 II

12 II þ 10 I

18 II16 II þ 4 I 18 II14 II þ 8 I

12 II þ 12 I

19 II18 II þ 2 I 2 IV þ 2 III þ 7 II 3 IV þ 9 II þ 8I

19 II mostly

B. carinata

B. napus

B. juncea

A2

A1

Alloploid

Chromosome pairing

—

— — —

18 II 18 II mostly —

— 17 II mostly 17 II mostly —

A3

poor

7–43 8.8 7–29

poor

6.2 26.1

poor

22–31

very poor poor

A1

poor

89 24 1952

poor

26.6 45.8

poor

26–69

very low 21.3

A2

—

96 36 5892

3164 6469

5482

— 27–53

A3

—

96 59 81–94

43–92 96

87

— 47–62

A4

Seed fetility (%)

—

96 79 96

96 96

94

— 87

A5

—

96

96

96 96

04

— 92

A6

Olsson 1960b Sarashima 1973 Prakash and Raut 1983 Song et al. 1993

Song et al. 1993

Olsson 1960a Prakash 1973a

Song et al. 1993

Frandsen 1947 Mizushima and Katsuo 1953 S. Prakash unpubl.

Reference

Table 2.10. Chromosome pairing and seed fertility in sexually synthesized Brassica allopolyploids: B. carinata, B. juncea, and B. napus.

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S. PRAKASH, S. R. BHAT, C. F. QUIROS, P. B. KIRTI, AND V. L. CHOPRA

Desirable characteristics in synthetic B. carinata include dwarf stature, early flowering, and high pod density (Prakash and Chopra 1996). A derivative of a synthetic ‘Pusa Gaurav’ has been released for cultivation in India. B. juncea synthesized from leafy vegetable forms has vigorous vegetative growth with more biomass and has been identified as preferred fodder. Synthetics originating from oleiferous forms of B. rapa are high yielders and contain more oil. Some of these are also early maturing (Prakash 1973a; Prakash and Chopra 1996). High-yielding cultivars released in India, such as ‘Pusa Jagannath’ and ‘Pusa Agrani’, have originated from synthetics. Several oil-yielding forms of B. napus that are derivatives of synthetics have been marketed in Europe. These include ‘Svalo¨f Panter’, a high-oil-yielding form suitable for low-temperature conditions; ‘Svalo¨f Norde’, a high-seed and oil-yielding form with considerable resistance to Peronospora and Verticillium; and ‘Brink’ and ‘Jupiter’, which combine high seed yields with very low erucic acid content (Olsson 1986). Several synthetic zero-erucic acid lines possess a high degree of resistance to Verticillium longisporum (Rygulla et al. 2007). A number of hybrids between synthetics and cultivars in B. napus have yield potential as high as 40 q/ha (Seyis et al. 2003,2006). Some of the synthetic strains that are early maturing and productive have been found to be promising for cultivation in India (Prakash and Raut 1983) and Bangladesh (Zaman 1989), where the photo- and thermo-sensitive strains from Europe and Canada remain vegetative and do not flower. Early-flowering (RC) and vigorous strains were also obtained by Hansen and Earle (1994). Among synthetics are strains that exhibit a high degree of resistance to diseases such as black leg (Mithen and Magrath 1992), clubroot (Diederichsen and Sacrista´n 1996), bacterial soft rot (Ren et al. 2000, 2001), Verticillium wilt (Happastadius et al. 2003), turnip yellows luteovirus (TuVV) (Paetsch et al. 2003), and pod shatter (Child et al. 2003). Winter oilseed rape varieties ‘Mendal’ and ‘Tosca’ combining clubroot resistance and cv. ‘Surpass’ carrying resistance to black leg were marketed in Europe in the last two decades. Synthesis of yellow-seeded B. napus has been the objective of many investigations because of the association of yellow seededness with increased oil and lower fiber content and also the improved feed quality of meal (Shirzadegan and Ro¨bbelen 1985; Chen et al. 1988; Chen and Hennen 1992; Tang et al. 1997; Meng et al. 1998; Baetzel et al. 1999; Rahman 2001; Wen et al. 2008), and such yellow-seeded synthetic forms have been obtained. A super-long silique form of resynthesized B. napus has been developed in China (Niu et al. 2004). Genetic variants for low glucosinolate content (Gland 1982) and

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modified fatty acid composition, particularly low erucic acid, have been created through resynthesis in B. napus (Chen and Hennen 1989b; Lu et al. 2001; Lu¨hs et al. 2002, 2003; Seyis et al. 2005). Synthetic rapeseed with high erucic acid content for industrial use has also been produced (Chen et al. 1989a; Lu¨hs and Friedt 1994, 1995a,b; Weir et al. 1997; Lu¨hs et al. 1999a,b; Han et al. 2001). Heath and Earle (1996, 1997) introduced a nonshattering trait and large-size seeds in somatic hybrids of B. napus. These authors also obtained B. napus somatic hybrids that were low in linolenic acid (Heath and Earle 1997) and high in erucic acid content (Heath and Earle 1995). Synthetic selfincompatible B. napus lines have recently been obtained through sexual hybridizations for developing commercial F1 hybrids (Rahman 2005). Fodder forms of B. napus have been synthesized in Japan and Sweden using leafy and root-forming forms of B. rapa—ssp. chinensis, pekinensis, narinosa, nipposinica, and rapa. Hosoda (1950) bred a fodder rape ‘CO’, which was very popular in Japan because of its vigorous growth and winter hardiness. A novel synthetic head-forming vegetable type has been developed in Japan from the cross B. oleracea var. capitata  B. rapa ssp. pekinensis. It is a popular vegetable form released in 1968 under the name ‘Hakuran’. It has soft leaves, fewer fibers, tastes like heady lettuce, and possesses high degree of resistance to soft rot (Shinohara and Kanno 1961; Takeda 1986). Such a type has also been produced using the same parents through protoplast fusion (Taguchi and Kameya 1986). As has been pointed out by many investigators, the germplasm pool of cultivated alloploid species is narrow. The genetic base can be enlarged by synthesizing polyploids with specifically chosen types of constituent parents. Two of the diploid constituent species—B. rapa and B. oleracea—are highly polymorphic, encompassing enormous physiological and morphological variations. These are of relatively recent origin and have not participated in the evolution of natural alloploid forms. Utilization of this variation will generate novel variability not available in the conventional germplasm. At present, resynthesis is routinely carried out using in vitro techniques following reciprocal hybridizations to gain the advantage of reciprocal cytoplasms, and by protoplast fusion to obtain various combinations of cytoplasmic organelles and mitochondrial recombination. Further variability in these synthetics also arises from intergenomic recombination between the chromosomes of constituent parents during early generations of allopolyploidy (Prakash 1973b). The desirable attributes of synthetics, such as disease resistance and quality characters, must

66

S. PRAKASH, S. R. BHAT, C. F. QUIROS, P. B. KIRTI, AND V. L. CHOPRA

be integrated into high-yielding varieties either by developing semisynthetic forms or in backcross breeding programs (Kraling 1987; Friedt et al. 2003). Many studies have suggested that heterosis for seed yield in intervarietal hybrids is positively correlated with genetic distance (Jain et al. 1994; Ali et al. 1995; Diers et al. 1995; Seyis et al. 2003; Shen et al. 2003; Burton et al. 2004). The variability and genetic distance of synthetics from cultivars in cultivation can be usefully exploited for generating both highly productive hybrids and genetically enhanced cultivars. Seyis et al. (2006) demonstrated the potential of synthesized B. napus for developing experimental hybrids having high yields. C. Diploidization of Allopolyploid Species As discussed earlier, the three basic genomes of Brassica crops share considerable homoeology among their chromosomes as evidenced from frequent occurrence of bivalents in diploid hybrids AB, AC, and BC. As a consequence, intergenomic allosyndetic pairing is expected in allopolyploids. However, the extant Brassica allopolyploids exhibit diploidlike meiosis and a completely bivalent forming regime. This diploidization of polyploids is an important step in their evolution and successful colonization. Several mechanisms including structural alterations in chromosomes, incorporation of adaptive mutations that prevent homoeologous chromosome pairing, and the silencing of duplicate genes have contributed this diploidization. It has been proposed that diploidlike meiosis is genetically regulated in Brassica and its related genera (Prakash 1974c; Harberd 1976; Attia and Ro¨bbelen 1986; Eber et al. 1994; Sharpe et al. 1995; Jenczewski et al. 2003). Prakash (1974c) proposed a genetic mechanism in B. juncea that is similar to the one that operates in wheat, where a gene is responsible for diploidlike meiosis and, when, absent allows paring of homoeologous chromosomes disturbing normal meiosis. His observations are based on the high extent of chromosome pairing in plants originating from a translocation heterozygote. These plants represent duplicationdeficiency types in which a very small segment presumably carrying the pairing regulator gene(s) is missing and which accounts for normal bivalents regime. However, because of the absence of monosomic series in alloploid Brassica, it has not been possible to validate this proposal experimentally. Attia and Ro¨bbelen (1986) observed low meiotic pairing in hybrids when B. nigra was one of the parents—for example, B. rapa  B. nigra and B. nigra  B. oleracea—and suggested a genetic meiotic suppressor in B. nigra genome that restricts the homoeologous

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67

pairing. However, this view was later discounted (Busso et al. 1987). By observing frequent intergenomic recombination in a B. rapa–B. alboglabra monosomic addition line but not in trigenomic AAC hybrids, Chen et al. (1992) proposed that it could be due to a pairing control mechanism. Jenczewski et al. (2002) postulated a pairing regulator gene for diploidlike meiotic regime in an induced autotetraploid of B. oleracea. Later, a hypothesis that envisions the presence of a major gene PrBn (Pairing regulator Brassica napus) in alloploid B. napus was proposed by Jenczewski et al. (2003). These authors studied the chromosome pairing in low- and high-bivalent-forming haploids of B. napus and observed that chromosome pairing patterns are inherited in a Mendelian way, indicating the presence of a major gene for restricting the homoeologous pairing. It was also suggested that since regular bivalents are observed in all B. napus accessions, regardless of bivalent frequency in their haploids, PrBn could contribute to the regularity of chromosome pairing. It could be ineffective at hemizygous stage or at least less efficient as compared to at the diploid state (Jenczewski and Alix 2004). PrBn gene has been mapped on a C genome chromosome and displays complete penetrance. Additionally, three to six minor QTL/BTL have slight additive effect on pairing without any interaction with PrBn. However, a number of other loci interact epistatically with PrBn (Liu et al. 2006). The other mechanism implicated in stabilization of alloploids is genic changes. Song et al. (1988a), following studies on nuclear RFLPs of diploid and alloploid species observed that nuclear DNA composition of alloploid species is closer to their cytoplasmic donor species than to the male parent donors. They concluded that when constituent diploid species of alloploids have highly differentiated cytoplasm, the nuclear genomes of male parent undergo substantial changes as compared to nuclear genome contributed by cytoplasmic donor. As mentioned earlier, two divergent types of cytoplasms occur in basic species: the B type as present in B. nigra and the A/C type as present in B. rapa and B. oleracea and quite diverged from the B type. As revealed by cp DNA studies (Erickson et al. 1983; Palmer et al. 1983a), B. juncea and B. napus have A type cytoplasm, while B. carinata has B type cytoplasm. During their evolution, nuclear genomes of male parents in B. juncea and B. carinata have undergone extensive changes while the nuclear genomes of female parents remained intact. In B. napus, both the genomes A and C have evolved with similar rates of change (Song et al. 1988a). Such changes are brought about by interaction between nuclear and cytoplasmic genomes and play an important role in stabilization of raw allopolyploids. The cytoplasm exerts selection

68

S. PRAKASH, S. R. BHAT, C. F. QUIROS, P. B. KIRTI, AND V. L. CHOPRA

pressure on alien nuclear genome and brings about a harmoneous interaction between cytoplasm and both the nuclear genomes in the new environment. To understand the process of changes in the genomes, Song et al. (1993) developed a series of synthetic alloploids of B. carinata, B. juncea, and B. napus following reciprocal hybridizations and characterized them for RFLP patterns of nuclear and cytoplasmic genomes. It was observed in subsequent generations that frequency of genome changes are associated with genetic divergence of constituent diploid parents: the more the genetic divergence, the higher the frequency of changes (Song et al. 1995). These changes could have resulted from chromosome rearrangements, point mutations, gene conversion, and DNA methylation. Intergenomic homologous recombination could lead to chromosome rearrangements and provide opportunities for gene conversion–like events (Osborn 2004; Pires et al. 2004, 2006). It has been suggested that extensive genome changes occur during early generations of polyploidy, and this accelerates the evolutionary processes (Song et al. 1995; Lukens et al. 2006). Also, intergenomic heterozygosity and epigenetic changes give rise to new variations crucial to their ecological success (Schranz and Osborn 2004). Another factor in stabilizing the chromosome pairing may be the role of rRNA genes. It has been reported in a number of allopolyploids that rRNA genes from only one parent are transcribed while the transcription of such genes of the other parent is suppressed: a phenomenon referred to as nucleolar dominance. A hierarchy of nucleolar dominance has been demonstrated to be B. nigra > B. rapa > B. olerace in three Brassica allotetraploids (Chen and Pikaard 1997; Pikaard 2000; Ge and Li 2007). These results suggest that nucleolar dominance may contribute decisively in preferential stabilization of chromosomes from rRNAsdonor parent. D. Raphanobrassica Raphanobrassica does not occur in nature but is a man-made curiosity—a hybrid between Raphanus sativus and B. oleracea. There is a long history of hybridizing Raphanus with B. oleracea. A sterile hybrid was obtained as early as 1826 by Sageret. However, the first fertile Raphanobrassica was synthesized by the Russian botanist Karpechenko (1927) following chromosome doubling of the hybrid R. sativus and B. oleracea. The classical Raphanobrassica has 36 somatic chromosomes representing full chromosome complements of R. sativus and B. oleracea. Subsequently, a number of researchers

2. BRASSICA AND ITS CLOSE ALLIES: CYTOGENETICS AND EVOLUTION

69

attempted its synthesis including Kakizaki (1927), Richharia (1937), Howard (1938), Kondo (1942), Crescini (1942), Honma and Heeckt (1962), Moskov and Makarova (1969), and McNaughton (1973). Details are presented by Prakash and Hinata (1980). More recently, it has been synthesized from the cross Raphanus sativus  B. oleracea var. alboglabra (Chen and Wu 2007). All these investigations involved sexual hybridization where Raphanus was always the female parent. The reciprocal sexual hybridization was not successful. The idea in later investigations was to develop fodder forms of Raphanobrassica incorporating the enormous diversity of B. oleracea. Hybrids were obtained by crossing diploids or tetraploids; however, the success was more at the tetraploid level. In recent years strains of Raphanobrassica have also been synthesized following protoplast fusion (Hagimori et al. 1992; Yamanaka et al. 1992). Although Raphanobrassica plants were intermediate between parents, in vegetative characters, B. oleracea traits are predominant. Flowers, however, have close resemblance with radish—larger in size with white petals and violet venations. The siliqua has a nondehiscent Raphanus–like apical portion and a dehiscent Brassica–like basal portion. Considerable variation in chromosome pairing has been observed by different investigators. Karpechenko (1927) found regular meiosis with 18 bivalents. However, Richharia (1937) and Howard (1938) reported frequent quadrivalents and trivalents. McNaughton (1973) observed that while regular 18 bivalents were predominant, occasionally 2 univalents were also present. A majority of Karpechenko’s plants were pollen and seed fertile, with seed fertility ranging from 51% to 66%. However, a high degree of seed sterility was reported by Richharia (1937; he found only 0.2 to 1.3 seeds per siliqua), Howard (1938), and McNaughton (1973). In a recent investigation, repeated selections resulted in highly fertile 10th-generation plants with 14.9 seeds per siliqua (Chen and Wu 2008). Somatic hybrids had only low levels of fertility (Hagimori et al. 1992; Yamanaka et al 1992). The low fertility in Raphanobrassica has been ascribed to chromosome irregularities, but mainly genetic imbalance led to embryo abortion following endosperm deficiency (Ellerstro¨m and Zagorcheva 1977; Iwasa and Ellerstro¨m 1981). As compared to sexual hybrids where Raphanus contributed both mt and cp genomes, in somatic hybrids radish chloroplast genomes dominated (90%). The mitochondrial genome was contributed either by Raphanus or B. oleracea and some also possessed recombinant mitochondria (Hagimori et al. 1992; Yamanaka et al. 1992). Since the 1970s, the major objective has been to exploit Raphanobrassica as a possible new forage crop combining the disease resistance, fast growth rate, and high

70

S. PRAKASH, S. R. BHAT, C. F. QUIROS, P. B. KIRTI, AND V. L. CHOPRA

quality of fodder radish with winter hardiness and high productivity of B. oleracea. In fact, several superior lines designated as Radicole were developed at Svalo¨f, Sweden (Olsson and Ellerstro¨m 1980) and the Scottish Plant Breeding Institute (McNaughton 1982). Some of the strains exceeded forage rape in fresh weight and dry matter yield by 20%. They also have resistance to clubroot and downy mildew. Somatic hybrids were generated to introgress clubroot resistance from radish to B. oleracea, and these did in fact posses high degree of resistance to clubroot (Hagimori et al. 1992; Yamanaka et al. 1992). Another synthetic alloploid involving radish is Raparadish (Brassicoraphanus, 2n ¼ 38). It was obtained from the cross B. rapa  R. sativus primarily for determining the homoeology between the two genomes (Terasawa 1932; Mizushima 1950b). Later the synthesis aimed at developing a fodder type (Tokumasu and Kato 1976, 1988) and transferring resistance to beet cyst nematode from Raphanus to B. rapa (Dolstra 1982). Raparadish grows vigorously, combining the rapid growth with resistance to beet cyst nematode and clubroot (Lange et al. 1989). A detailed cytogenetical study on Brassicoraphanus synthesized for fodder has been carried out by Tokumasu and Kato (1976) and Matsuzawa et al. (2000), who recorded 15–19 II þ 08 I at M1 of meiosis, with occasional occurrence of a tri- or quadrivalent. However, the pollen fertility was low (0–89%) and the seed set was 0.01 to 0.1 seeds per siliqua after self- and open pollinations, respectively. Some of the A3 generation plants with yellow flowers showed considerably improved fertility. It was suggested that the genes for flower color are closely linked with those controlling embryo development. The genetic reconstitution due to intergenomic segmental exchange promotes development of embryos leading to higher fertility in yellowflowered plants (Kato and Tokumasu 1976; Tokumasu and Kato 1988). Matsuzawa et al. (2000) reported that two Raparadish lines had potential to be used as fodder. Further, they also obtained it from the reciprocal cross R. sativus  B. rapa, which showed mostly regular meiosis with 19 bivalents. E. Higher Allopolyploids in U Triangle Species through Protoplast Fusion The optimum level of genome organization in crop brassicas is tetraploidy, as represented by naturally occurring digenomic tetraploids. However, it is possible to bring together all the three basic genomes in one combination. In the past, such combinations were created to determine chromosome homoeology among A, B, and C

2. BRASSICA AND ITS CLOSE ALLIES: CYTOGENETICS AND EVOLUTION

71

genomes (see Prakash and Hinata 1980). In recent years, the three genomes have been brought together through protoplast fusion (Table 2.11) to make use of them as bridge species for transferring traits of agronomic importance, particularly resistance to fungal diseases, such as blackleg and clubroot caused by Phoma lingam and Plasmodiophora brassiceae, respectively. These are serious diseases on B. napus in Europe, Australia, and Canada. Genes conferring resistance are available in B. nigra and natural alloploid species containing B. nigra genome, specifically B. carinata and B. juncea (Sacrista´n and Gerdeman-Kno¨rck 1986; Sjo¨din and Glimelius 1989a,b; Zhu and Spanier 1991). The other objectives are incorporation of herbicide resistance and cytoplasmic male sterility (Kao et al. 1992; Hansen and Earle 1995; Arumugam et al. 1996).

IV. WIDE HYBRIDIZATION Hybridization in brassicas goes back to early 19th century when Sageret (1826) obtained intersub-tribal hybrid Raphanus sativus  B. oleracea and Herbert (1847) produced interspecific hybrid B. napus  B. rapa. Cytogenetical interest following determination of chromosome numbers gave a boost to wide hybridization. Initial attempts at hybridizations were for elucidating genomic homoeology. Later, attention shifted to utilizing wide hybridization for expanding genetic variability, introgressing nuclear genes that conferred desirable agronomic traits or cytoplasmic genes for inducing male sterility. Chromosome addition lines have also been generated to locate genes on specific chromosomes and for construction of genetic maps. During the last 30 years, in vitro techniques such as ovary and embryo culture and protoplast fusion have been employed successfully to obtain a large number of sexual and somatic hybrids. A. Sexual Hybrids Initial attempts at hybridizatization through conventional methods required very large numbers of pollinations to realize hybrids. Success was negligible for many combinations because of inability of pollen tubes to grow down the style to effect fertilization. Even after successful fertilization, hybrid embryos can abort due to genetic incompatibilities between the developing embryo and the endosperm or to partial sterilization of ovaries by alien pollen tube. In some instances, unilateral incompatibility has also been observed (Harberd 1976; Shivanna 1996).

72

52, BBCCCC 54, AABBCC

56, AACCCC

54, AABBCC 72, AABBCCCC 74, AAAABBCC

B. napus þ B. oleracea

B. juncea þ B. oleracea B. napus þ B. carinata B. napus þ B. juncea

Somatic chromosome number and genomes

B. carinata þ B. oleracea B. napus þ B. nigra

Somatic hybrids

Hygromycin resistance Club root resistance Atrazine resistance Polima CMS Not defined CMS Ogu, triazine tolerance Xanthomonas resistance, CMS Ogu, atrazine resistance Not defined Plastome transformation Tour CMS Black leg resistance Black leg resistance Male sterility

Disease resistance Black leg resistance

Trait to transfer

Table 2.11. Interspecific somatic hybrids within U triangle species.

Olin-Fatih et al. 1996 Nitovskaya et al. 2006a Arumugam et al. 1996 Sjo¨din and Glimelius 1989b Sjo¨din and Glimelius 1989b Szasz et al. 1991

Ryschka et al. 1996 Sjo¨din and Glimelius 1989a; Yamagishi et al. 1989; Sakhno et al. 1991 Sacrista´n et al. 1989 Gerdemann-Kno¨rck et al. 1994, 1995 Nitovskaya et al. 1988; Jourdan et al. 1989 Yarrow et al. 1990 Sundberg et al. 1991 Kao et al. 1992 Hansen and Earle 1995

Reference

2. BRASSICA AND ITS CLOSE ALLIES: CYTOGENETICS AND EVOLUTION

73

Limited investigations have been undertaken to determine the details of postfertilization barriers. Lack of a functional endosperm or its early degeneration appear to be the major reasons for abortion of hybrid embryos. Ways devised to overcome these hybridization barriers include grafting, mixed pollination, bud pollination, and stump pollination (Hosoda et al. 1963; Sarashima 1964; Namai 1971). Kameya and Hinata (1970) succeeded in performing in vitro fertilization and obtained interspecific hybrids. A modified technique of placental pollination was used by Zenkteler (1990). Embryo rescue technique has been an effective technique for overcoming postfertilization barriers and is used extensively to obtain wide hybrids. Japanese scientists, particularly Nishi and his group (1959), pioneered it in Brassica in the late 1950s (Nishi et al. 1959). Sequential culture, which involves successive culture of ovaries, ovules, and seeds/embryos, is more effective than simple ovary or ovule culture (Shivanna 1996; Wen et al. 2008). Although wide hybridizations in Brassica have been carried out for a long time, here we define it in terms of hybridizing species of secondary and tertiary gene pools. A pioneer in this area was Mizushima (1950a,b, 1968) who attempted such hybridizations involving wild germplasm. Subsequent extensive investigations were by Harberd and McArthur (1980), who reported nearly 50 distant hybrids in which a majority were intergeneric. At present, hybridization between wild and crop species has become a routine. The last 20 years have witnessed a large number of sexual hybrids comprising interspecific, intergeneric, intersubtribal, and intertribal combinations. These hybrids and their meiotic behavior are listed in Table 2.12. Sexual hybrids are characterized by a highly disorganized meiosis, particularly when both parents are diploid. Chromosomes, due to the absence of a homologous partner, remain mostly as univalents but occasionally undergo pairing and also form bivalents in a very low frequency. Bivalents, when they occur, are mostly rod-shape monochiasmates and rarely ring shape with multiple chiasmata. Multivalents in diploid hybrids occur only rarely. However, a variable number of bivalents and frequent trivalent/quadrivalents are formed in triploid (tetraploid  diploid) and tetraploid (tetraploid  tetraploid species) combinations. Harberd and McArthur (1980) observed a close relationship between mean chromosome number and mean bivalent frequency at three ploidy levels (Table 2.13). Among the diploid hybrids, high chromosome pairing has been observed in several combinations:. Sinapis arvensis  B. nigra (2n ¼ 17, 8 II, Mizushima 1950a), Diplotaxis erucoides  B. nigra (2n ¼ 15, 6 II,

74 15 16 17

Brassica maurorum (n ¼ 8)  Erucastrum varium (n ¼ 7)  Brassica nigra (n ¼ 8)

 Sinapis arvensis (n ¼ 9)

17 I–3 II þ 11 I

15 I–4 II þ 7 I 16 I–7 II þ 2 I

II þ 2 I II þ 11 I II þ 10 I II þ 4 I

10–24 I þ 1--7 II þ 0--1 IV

15 26 I–7 I–3 I–4 I–7

16 I–4 II þ 8 I

16

16 17 18 18

Harberd and McArthur 1980

15 I–5 II þ 5 I

15

16 17 18 18

Takahata and Hinata 1983

1 III þ 19 II þ 4 I

45

Erucastrum varium (n ¼ 7)  Brassica nigra (n ¼ 8) Erucastrum virgatum (n ¼ 7)  Sinapis pubescens (n ¼ 9) Hirschfeldia incana (n ¼ 7)  Brassica nigra (n ¼ 8)  Brassica napus (n ¼ 19) Brassica fruticulosa (n ¼ 8)  Brassica nigra (n ¼ 8)  Erucastrum cardaminoides (n ¼ 9)  Brassica barrelieri (n ¼ 10)  Brassica rapa (n ¼ 10)

Takahata and Hinata 1983 Takahata and Hinata 1983; Truco and Quiros 1991 Truco and Quiros 1991

Mizushima 1968; Truco and Quiros 1991 Takahata and Hinata 1983 Harberd and McArthur 1980 Mizushima 1968; Takahata and Hinata 1983; Nanda Kumar et al. 1988

Mattsson 1988 Kerlan et al. 1993

Quiros et al. 1988 Quiros et al. 1988 Harberd and McArthur 1980 Vyas et al. 1995 Vyas et al. 1995 Vyas et al. 1995 Quiros 1986a; Ringdahl et al. 1987; Delourme et al. 1989 Vyas et al. 1995

14 I–6 II þ 2 I 15 I–7 II þ 1 I 16 I–4 II þ 8 I 25 I–7 II þ 11 I 17 I–1 III þ 3 II þ 8 I 1 II þ 23 I–1 IV þ 6 II þ 9 I 1 IV þ 1 II þ 5 I–13 II þ 6--23 I

14 15 16 25 17 25 26

Diplotaxis erucoides (n ¼ 7)  Hirschfeldia incana (n ¼ 7)  Brassica nigra (n ¼ 8)  Sinapis pubescens (n ¼ 9)  Brassica oleracea (n ¼ 9)  Brassica rapa (n ¼ 10)  Brassica juncea (n ¼ 18)  Brassica napus (n ¼ 19)

Reference

2n

Hybrids

Chromosome pairing at M1 of meiosis

Table 2.12. Wide hybrids in Brassica coenospecies and their meiotic behavior.

75

Takahata and Hinata 1983 Nandkumar and Shivanna 1993 Mizushima 1980 Quiros et al. 1988 Sarashima et al. 1980 Mizushima 1950a; Momotaz et al. 1998 Mizushima 1950a Harberd and McArthur 1980 Harberd and McArthur 1980; Momotaz et al. 1998 Li and Heneen 1999 Apel et al. 1984 Harberd and McArthur 1980

16 I–4 II þ 8 I 16 I–6 II þ 4 I 16 I–4 II þ 8 I 16 I–6 II þ 4 I

16–18 16 16 18 16 16 18 18 18 21 21

 Orychophragmus violaceus (n ¼ 12)  Moricandia arvensis (n ¼ 14)  Diplotaxis muralis (n ¼ 21)

 Orychophragmus violaceus (n ¼ 12) Brassica spinescens (n ¼ 8)  Brassica nigra (n ¼ 8) Diplotaxis siettiana (n ¼ 8)  Brassica nigra (n ¼ 8)  Brassica rapa (n ¼ 10) Brassica oleracea (n ¼ 9)  Diplotaxis erucoides (n ¼ 7)  Hirschfeldia incana (n ¼ 7)  Raphanus sativus (n ¼ 9)  Sinapis arvensis (n ¼ 9)  Sinapis pubescens (n ¼ 9)  Coincya spp. (n ¼ 12)  Sinapis alba (n ¼ 12) 3 III þ 4 II þ 4 I–21 I 1 II þ 28 I–6 II þ 18 I

30

I I II þ 7 I II þ 18 I

21

18 I–5 II þ 8 18 I–5 II þ 8 2 II þ 17 I–7 2 II þ 19 I–6

(continued)

Truco and Quiros 1991

3 III þ 4 II þ 4 I–21 I

16 16 17 17

    1 II þ 15 I–8 II þ 1 I

Harberd and McArthur 1980 Quiros et al. 1988 Harberd and McArthur 1980; Prakash et al. 1982; Mattson 1988 Prakash et al. 1982 Prakash et al. 1982 Matsuzawa and Sarashima 1984 Mizushima 1950a; Harberd and McArthur 1980; Mattson 1988 Li and Heneen 1999

1 II þ 13 I–4 II þ 7 I 15 I–1 II þ 5 II þ 2 I 16 I–2 II þ 12 I

15 15 16

Brassica maurorum (n ¼ 8) Brassica spinescens (n ¼ 8) Raphanus sativus (n ¼ 9) Sinapis arvensis (n ¼ 9)

Takahata and Hinata 1983 Takahata and Hinata 1983

18 I–2 II þ 14 I 18 I–5 II þ 8 I

18 18

 Brassica barrelieri (n ¼ 10)  Brassica rapa (n ¼ 10) Brassica nigra (n ¼ 8)  Erucastrum virgatum (n ¼ 7)  Hirschfeldia incana (n ¼ 7)  Brassica fruticulosa (n ¼ 8)

76

Erucastrum canariense (n ¼ 9)  Brassica oleracea (n ¼ 9)  Brassica rapa (n ¼ 10) Erucastrum cardaminoides (n ¼ 9)  Brassica nigra (n ¼ 8)  Brassica oleracea (n ¼ 9)  Brassica rapa (n ¼ 10) Raphanus sativus (n ¼ 9)  Brassica fruticulosa (n ¼ 8)  Brassica nigra (n ¼ 8)  Brassica oleracea (n ¼ 9)  Brassica rapa (n ¼ 10)  Brassica juncea (n ¼ 18)  Brassica napus (n ¼ 19)

Brassica oxyrrhina (n ¼ 9)  Brassica nigra (n ¼ 8)  Brassica oleracea (n ¼ 9)  Raphanus sativus (n ¼ 9)  Sinapis pubescens (n ¼ 9)  Brassica barrelieri (n ¼ 10)  Brassica rapa (n ¼ 10)  Brassica tournefortii (n ¼ 10) Diplotaxis catholica (n ¼ 9)  Brassica rapa (n ¼ 10)  Brassica juncea (n ¼ 18) Diplotaxis virgata (n ¼ 9)  Brassica rapa (n ¼ 10)  Brassica juncea (n ¼ 18)

Hybrids

Table 2.12. (Continued)

Harberd and McArthur 1980 Prakash et al. 2001; Bhasker et al. 2002 Chandra et al. 2004 Mohanty 1996 Chandra et al. 2004

1II þ 16 I–8 II þ 2 I 19 I–4 II þ 11 I 17 I–5 II þ 7 I 18 I–1 IV þ 1 III þ 1 II þ 9 I 19 I–5 II þ 9 I

18 I–1 III þ 6 II þ 3 I 1 II þ 17 I–7 II þ 5 I

18 19 17 18 19 17 17 18 19 27 28

8–20 I þ 4--10 II þ 0--1 IV

Takahata and Hinata 1983 Harberd and McArthur 1980 Inomata 2003

19 I–6 II þ 7 I 1 II þ 25 I–8 II þ 11 I IV þ 02 III þ 8--12 II þ 10--20 I

19 27 36

Bang et al. 1997 Matsuzawa and Sarashima 1984, 1986 McNaughton 1973; Chen and Wu 2008 Mizushima 1950a; Matsuzawa et al. 2000 Rhee et al. 1997 Kerlan et al. 1993

Mohanty 1996 Mohanty 1996

Prakash et al. 1982 Harberd and McArthur 1980 Matsuzawa et al. 1997; Kaneda and Kato 1999 Harberd and McArthur 1980 Mattsson 1988 Prakash and Chopra 1990 Mattsson 1988

Reference

2 II þ 15 I–1 III þ 3 II þ 10 I 13 I þ 7 II–2 III þ 7 II þ 7 I

19 I–4 II þ 11 I

2 II þ 14 I – 6 II þ 6 I 18 I–3 II þ 12 I 1 II þ 16 I–4 II þ 10 I

Chromosome pairing at M1 of meiosis

19 17

17 18 18 18 19 19 19

2n

77

 Coincya pseuderucastrum (n ¼ 12)  Sinapis alba (n ¼ 12)  Orychophragmus violaceus (n ¼ 12)  Moricandia arvensis ( n ¼ 14)  Capsella bursa–pastoris (n ¼ 16) Brassica tournefortii (n ¼ 10)  Brassica fruticulosa (n ¼ 8)  Brassica nigra (n ¼ 8) 10–20 II þ 2--11 I 24 I–5 II þ 14 I 3 I–13 II

18 I–3 II þ 12 I

22 22 23–42 24 29 18 18

1 IV þ 11--15 II þ 4--12 I

20 I–4 II þ 12 I 20 I–8 II þ 5 I

19 I–5 II þ 9 I

(continued)

19 20 20 21

   

19 I–5 II þ 9 I

Prakash et al. 1982 Narain and Prakash 1972

Mizushima 1968 Mizushima 1968 Prakash et al. 1982 Harberd and McArthur 1980 Mithen and Herron 1991 Mizushima 1950a; Harberd and McArthur 1980; Matsuzawa et al. 2000 Mizushima 1950a Mattsson 1988 Prakash and Narain 1971 Mizushima 1950a; Harberd and McArthur 1980 Inomata 2004 Jandurova and Dolezel 1995 Li and Heneen 1999 Takahata and Takeda 1990 Chen et al. 2007

17 I–5 II þ 7 I 18 I–7 II þ 4 I

17 18 18 18 19 19

Sinapis arvensis (n ¼ 9) Brassica barrelieri (n ¼ 10) Brassica tournefortii (n ¼ 10) Eruca sativa (n ¼ 11)

Takahata and Hinata 1983 Takahata and Hinata 1983 Harberd 1972 Harberd and McArthur 1980 Takahata and Hinata 1983

18 I–4 II þ 10 I 18 I–4 II þ 10 I 9 II þ 1 I 19 I–6 II þ 7 I 20 I–6 II þ 8 I

18 18 19 19 20

1 II þ 16 I–5 II þ 8 I

Mizushima 1950a Momotaz et al. 1998 Mathias 1991

18 I–3 II þ 12 I

18 19 28

Sinapis arvensis (n ¼ 9)  Raphanus sativus (n ¼ 9)  Brassica rapa (n ¼ 10)  Brassica napus (n ¼ 19) Brassica barrelieri (n ¼ 10)  Brassica fruticulosa (n ¼ 8)  Brassica nigra (n ¼ 8)  Brassica oxyrrhina (n ¼ 9)  Sinapis pubescens (n ¼ 9)  Brassica rapa (n ¼ 10) Brassica rapa (n ¼ 10)  Hirschfeldia incana (n ¼ 7)  Brassica fruticulosa (n ¼ 8)  Brassica spinescens (n ¼ 8)  Erucastrum leucanthum (n ¼ 8)  Brassica atlantica (n ¼ 9)  Raphanus sativus (n ¼ 9)

78 Harberd and McArthur 1980 Mohanty 1996 Mohanty 1996 Gundimeda et al. 1992 Gundimeda et al. 1992 Harberd and McArthur 1980 Gundimeda et al. 1992 Gundimeda et al. 1992 Harberd and McArthur 1980 Harberd and McArthur 1980 Harberd and McArthur 1980

21 I–4 II þ 13 I 3 III þ 4 II þ 10 I 2 IV þ 1 II 2 II þ 14 I 19 I–1 III þ 4 II þ 8 I 20 I–2 III þ 4 II þ 6 I 6 II þ 14 I–10 II þ 6 I 1 III þ 13 II þ 8 I 29 I–IV þ 1 III þ 6 II þ 10 I 3 II þ 12 I–9 II 1 II þ 17 I–6 II þ 7 I 2 II þ 24 I–6 II þ 12 I

21 27 29 19 20 26 37 29 18 19 28

20 I–5 II þ 10 I

Ahuja et al. 2003 Batra et al. 1990 Batra et al. 1990

20

 Brassica rapa (n ¼ 10)

19 I–5 II þ 9 I

Mizushima 1968; Narain and Prakash 1972 Mattsson 1988 Mizushima 1950a; Harberd and McArthur 1980 Mizushima 1950a; Harberd and McArthur 1980 Sikka 1940; Mizushima 1968; Choudhary and Joshi 2001

Reference

20 I–6 II þ 8 I 28 I–6 II þ 22 I

19

 Sinapis arvensis (n ¼ 9)

19 I–7 II þ 5 I

19 I–3 II þ 32 I

Chromosome pairing at M1 of meiosis

20 28 29

19 19 19

 Brassica oleracea (n ¼ 9)  Brassica oxyrrhina (n ¼ 9)  Raphanus sativus (n ¼ 9)

Diplotaxis siifolia (n ¼ 10)  Brassica rapa (n ¼ 10)  Brassica juncea (n ¼ 18)  Brassica napus (n ¼ 19) Diplotaxis viminea (n ¼ 10)  Diplotaxis tennuifolia (n ¼ 11)  Brassica carinata (n ¼ 17)  Brassica napus (n ¼ 19) Enarthrocarpus lyratus (n ¼ 10)  Brassica oleracea (n ¼ 9)  Brassica rapa (n ¼ 10)  Erucastrum abyssinicum (n ¼ 16)  Brassica carinata (n ¼ 17)  Brassica napus (n ¼ 19) Sinapidendron frutescens (n ¼ 10)  Brassica fruticulosa (n ¼ 8)  Sinapis pubescens (n ¼ 9)  Brassica juncea (n ¼ 18)

2n

Hybrids

Table 2.12. (Continued)

79

 Diplotaxis tennuifolia (n ¼ 11) Sinapis alba (n ¼ 12)  Brassica oleracea (n ¼ 9)  Brassica napus (n ¼ 19) Erucastrum laevigatum (n ¼ 14)  Hirschfeldia incana (n ¼ 7) Moricandia arvensis (n ¼ 14)  Brassica nigra (n ¼ 8)  Brassica oleracea (n ¼ 9)  Raphanus sativus (n ¼ 9)  Brassica rapa (n ¼ 10)  Brassica juncea (n ¼ 18)  Bassica napus (n ¼ 19)  Diplotaxis muralis (n ¼ 21) Moricandia nitens (n ¼ 14)  Brassica napus (n ¼ 19)

Diplotaxis tennuifolia (n ¼ 11)  Erucastrum virgatum (n ¼ 7)  Hirschfeldia incana (n ¼ 7)  Brassica nigra (n ¼ 8)  Brassica oleracea (n ¼ 9)  Raphanus sativus (n ¼ 9)  Brassica rapa (n ¼ 10)  Brassica elongata (n ¼ 11)  Coincya spp. (n ¼ 12)  Brassica juncea (n ¼ 18) Eruca sativa (n ¼ 11)  Brassica oleracea (n ¼ 9)  Raphanus sativus (n ¼ 9)  Brassica rapa (n ¼ 10)

(continued)

Takahata and Takeda 1990 Takahata 1990 Bang et al. 1995 Takahata and Takeda 1990 Takahata et al. 1993 Takahata et al. 1993; Meng et al. 1997 Razmjoo et al. 1996

22 I–1 III þ 5 II þ 9 I 23 I–2 III þ 6 II þ 5 I 1 III þ 0--5 II þ 13--23 I 24 I–5 II þ 14 I

22 23 23 24 32 33 35

Meng et al. 1999

Harberd and McArthur 1980

3 II þ 15 I–9 II þ 3 I

21

33

U et al. 1937; Wei et al. 2007 Ripley and Arnison 1990; Brown et al. 1997

21 I, 21 I–3 II þ 15 I 2 IV þ 6 II þ 11 I

21 31

21 II 22 I–5 II þ 12 I

1980 1980

1980

1980 1980

U et al. 1937; Matsuzawa and Sarashima 1986 Matsuzawa and Sarashima 1986 Agnihotri et al. 1988; Matsuzawa et al. 1999 Takahata and Hinata 1983

Harberd and McArthur Harberd and McArthur Salisbury 1989 Harberd and McArthur Bang et al. 2003 Salisbury 1989 Harberd and McArthur Harberd and McArthur Salisbury 1989

20 I–3 II þ 14 I

22 I–4 II þ 14 I 23 I–5 II þ 13 I

20 I–2 II þ 16 I

18 I–7 II þ 4 I 18 I–6 II þ 6 I

21 20 42 21 22

18 18 19 20 20 21 22 23 29

80 26 33 34

7 II þ 12 I–10 II þ 6I

1980

1980 1980 1980 1980;

 Brassica rapa (n ¼ 10)  Brassica carinata (n ¼ 17)  Brassica juncea (n ¼ 18)

Harberd and McArthur Harberd and McArthur Harberd and McArthur Harberd and McArthur Rao et al. 1996 Harberd and McArthur Rao et al. 1996 Rao et al. 1996

II þ 9 I–9 II þ 5 I II þ 14 I –8 II þ 8 I II þ 14 I–10 II þ 4 I II þ 23 I–9 II þ 7 I

23 24 24 25

7 5 5 1

Harberd and McArthur 1980

13 II þ 9 I–16 II þ 3 I

35

Harberd and McArthur 1980 Harberd and McArthur 1980 Harberd and McArthur 1980 Harberd and McArthur 1980 Takahata and Hinata 1983 Harberd and McArthur 1980 Harberd and McArthur 1980 Harberd and McArthur 1980 Harberd and McArthur 1980 Harberd and McArthur 1980 Harberd and McArthur 1980 Batra et al. 1989 Batra et al. 1989 Snogerup and Persoon 1983 Snogerup and Persoon 1983

II þ 18 I–8 II þ 6 I II þ 20 I–7 II þ 8 I II þ 20 I–7 II þ 8 I II þ 13 I–10 II þ 3 I II þ 9 I–8 II þ 7 I II þ 18 I–8 II þ 8 I II þ 20 I–9 II þ 6 I II þ 16 I–9 II þ 6 I II þ 19 I–8 II þ 9 I II þ 17 I–8 II þ 9 I II þ 17 I–9 II þ 7 I III þ 1 II þ 22 I III þ 6 II þ 17 I

9 II þ 7 I

2 1 1 5 7 3 2 4 3 3 3 3 3

Reference

25 25

24 24 24 25 25 25 33 38

22 22 22 23

Erucastrum gallicum (n ¼ 15)  Diplotaxis erucoides (n ¼ 7)  Erucastrum virgatum (n ¼ 7)  Hirschfeldia incana (n ¼ 7)  Erucastrum nasturtiifolium (n ¼ 8)

Chromosome pairing at M1 of meiosis

 Raphanus sativus (n ¼ 9)  Sinapis arvensis (n ¼ 9)  Sinapis pubescens (n ¼ 9)  Brassica barrelieri (n ¼ 10)  Enarthrocarpus lyratus (n ¼ 10)  Sinapidendron frutescens (n ¼ 10)  Brassica juncea (n ¼ 18)  Brassica napus (n ¼ 19) Brassica balearica (n ¼ 16)  Brassica oleracea. alboglabra (n ¼ 9)  Brassica oleracea var. insularis (n ¼ 9) Brassica cossoneana (n ¼ 16)  Brassica napus (n ¼ 19) Erucastrum abyssinicum (n ¼ 16)  Erucastrum virgatum (n ¼ 7)  Erucastrum leucanthemum (n ¼ 8)  Erucastrum nasturtiifolium (n ¼ 8)  Brassica oleracea (n ¼ 9)

2n

Hybrids

Table 2.12. (Continued)

81

 Sinapis pubescens (n ¼ 9)  Brassica gravinae (n ¼ 10)  Brassica tournefortii (n ¼ 10)  Enarthrocarpus lyratus (n ¼ 10)  Sinapis alba (n ¼ 12)  Orychophragmus violaceus (n ¼ 12)  Crambe abyssinica (n ¼ 45) Brassica napus (n ¼ 19)  Lesquerella fendleri (n ¼ 6)  Diplotaxis erucoides (n ¼ 7)  Hirschfeldia incana (n ¼ 7)

 Sinapis pubescens (n ¼ 9)  Orychophragmus violaceus (n ¼ 12)  Sinapis alba (n ¼ 12)  Erucastrum gallicum (n ¼ 15) Brassica juncea (n ¼ 18)  Diplotaxis virgata (n ¼ 9)  Raphanus sativus (n ¼ 9)  Sinapis arvensis (n ¼ 9)

Brassica carinata (n ¼ 17)  Brassica fruticulosa (n ¼ 8)  Diplotaxis assurgens (n ¼ 9)  Diplotaxis tenuisiliqua (n ¼ 9)  Diplotaxis virgata (n ¼ 9)  Raphanus sativus (n ¼ 9)  Sinapis arvensis (n ¼ 9)

Mixoploids 63 I Irregular 3–10 I I þ 6--20 I 0–4 IV þ 1--7 II þ 10--24 I

25–42 26 26

Du et al. 2008 Harberd and McArthur 1980 Kerlan et al. 1993 (continued)

Gundimeda et al. 1992 Mohapatra and Bajaj 1990 Li et al. 1998a,b Wang and Luo 1998

28 I–7 II þ 14 I

0–6 I I þ 15--27 I 28 I–1 8 II þ 2 I

7–10 I I þ 7--13 I

Inomata 2003 Fukushima 1945 Mizushima 1950a; Harberd and McArthur 1980 Inomata 1991 Nandkumar et al. 1989

Harberd and McArthur 1980 Harberd and McArthur 1980 Harberd and McArthur 1980 Harberd and McArthur 1980 Harberd and McArthur 1980 Mizushima 1950a; Bing et al. 199; Momotaz et al. 1998 Harberd and McArthur 1980 Hua et al. 2006 Momotaz et al. 1998 Harberd and McArthur 1980

0–4 IV þ 0--2 III þ 7--12 II þ 12--20 I

5–12 I I þ 8--22 I

3–17 I I þ 12--20 I Mixoploids, mostly 17 II

3 II þ 19 I–4 II þ 17 I 3 II þ 20 I–10 II þ 6 I 1 II þ 24 I–10 II þ 6 I 4 II þ 18 I–11 II þ 4 I 2 II þ 22 I–9 II þ 8 I 26 I–8 II þ 10 I

27 28 28 28 30 30–36 63

27 27 27

26 17–35 29 32

25 26 26 26 26 26

82

 Diplotaxis harra (n ¼ 12)  Erucastrum gallicum (n ¼ 15)  Brassica cossoneana (n ¼ 16)  Capsella bursa–pastoris (n ¼ 16) Diplotaxis muralis (n ¼ 21)  Brassica rapa (n ¼ 10)  Sinapidendron frutescens (n ¼ 10)  Diplotaxis harra (n ¼ 13)  Erucastrum gallicum (n ¼ 15)  Brassica napus (n ¼ 19) 31 31 34 36 40

26 28 28 28 28 39 30–38 35–37 31 34 35 29

      

Matthiola incana (n ¼ 7) Raphanus raphanistrum (n ¼ 9) Raphanus sativus (n ¼ 9) Sinapis arvensis (n ¼ 9) Sinapis pubescens (n ¼ 9) Brassica gravinae (n ¼ 10) Orychophragmus violaceus (n ¼ 12)

2n

Hybrids

Table 2.12. (Continued)

31 I–5 II þ 21 I 1 II þ 29 I–5 II þ 21 I 34 I–6 II þ 22 I 3 II þ 3010 II þ 16 I

12–16 I I þ 3--11 I 1 III þ 9 II þ 8 I

16–18 II þ 3--1 I 0–2 III þ 2--10 II þ 12--28 I

0–2 III þ 0--10 II þ 6--28 I 0–1 III þ 2--11 II þ 5--24 I 39 I–10 II þ 19 I

0–4 IV þ 4--10 II þ 8--20 I

Chromosome pairing at M1 of meiosis

Harberd and McArthur 1980 Harberd and McArthur 1980 Harberd and McArthur 1980 Harberd and McArthur 1980 Fan et al. 1985; Ringdhal et al. 1987

Peng et al. 2003 Kerlan et al. 1993; Lefol et al. 1997 Takeshita et al. 1980 Kerlan et al. 1993 Inomata 1991, 1994 Nandkumar et al. 1989 Li et al. 1995; Hua and Li 2006; Cheng et al. 2002 Inomata 2005 Lefol et al. 1997 Harberd and McArthur 1980 Chen et al. 2007

Reference

2. BRASSICA AND ITS CLOSE ALLIES: CYTOGENETICS AND EVOLUTION

83

Table 2.13. Mean chromosome number and bivalent frequency at three ploidy levels in the tribe Brassiceae. Hybrid Diploids Triploids Tetraploids

Mean chromosome number, 2n 18.7 25.6 34.3

Mean bivalent frequency 2.9 6.2 10.2

Source: Harberd and McArthur 1980.

Quiros et al. 1988), B. fruticulosa  B. nigra (2n ¼ 16, 7 II, Mizushima 1968), B. nigra  Hirschfeldia incana (2n ¼ 15, 1 III þ 5 II, Quiros et al. 1988), Erucastrum canariense  B. oleracea (2n ¼ 18, 8 II, Harberd and McArthur 1980), E. cardaminoides  B. oleracea (2n ¼ 18, 1 IV þ 1 III þ 1 II, Mohanty 1996), and Enarthrocarpus lyratus  B. rapa (2n ¼ 20, 2 III þ 4 II, Gundimeda et al. 1992). The triploid and tetraploid hybrids where higher associations have been observed include B juncea  Diplotaxis virgata (1 IV/2 III, Inomata 2003), B. napus  Hirschfeldia incana (1 IV, Kerlan et al. 1993), Diplotaxis viminea  B. napus (2 IV, Mohanty 1996), and Diplotaxis erucoides  B. napus (1 IV, Delourme et al. 1989). Hybrids between the diploids were absolutely pollen and seed sterile while triploid and tetraploid hybrids had a little pollen and seed fertility. Bivalents and higher associations may be interpreted to result from archaic homology within the chromosomes of the same genome (autosyndesis) or because of intergenomic homoeology (allosyndesis). However, it is difficult to interpret the pairing precisely in terms of auto- or allosyndesis since there is little information on the extent of autosyndesis observed through chromosome pairing in haploids. Mizushima (1950a, 1968, 1980) made some observations on the extent of allosyndesis between a limited number of genomes; Harberd and McArthur (1980) could not arrive at any definite conclusion. What can be stated safely is that intrageneric homoeology is not always higher than intergeneric homoeology. With the progress in GISH techniques, the degree of autoand allosyndesis can be ascertained precisely in wide hybrids. An interesting cytological phenomenon was observed in hybrids between Brassica species and Orychophragmus violaceus (2n ¼ 24). O. violaceus is cultivated in China as an ornamental plant and has desirable oil quality. Hybrids with all the six crop species have been obtained with O. violaceus always the pollen parent. Chromosomes remain unpaired as univalents in hybrid cells, and separation of parental genomes occurs regularly during mitotic and meiotic

84

S. PRAKASH, S. R. BHAT, C. F. QUIROS, P. B. KIRTI, AND V. L. CHOPRA

divisions (Li et al. 1995, 1996, 1998a,b, 2003; Li and Heneen 1999). During mitosis, any of three situations may occur, and subsequently the chromosomes are doubled following chromosome duplications in daughter cells: 1. Complete separation of parental genomes results into cells with haploid and diploid complements of the two parents. 2. Partial separation leads to inclusion of some chromosomes of one parent with the haploid complement another genome producing hypo- and hyperdiploid cells. 3. During partial separation, chromosomes of either parent are included in the genomes resulting into substitution lines. Hybrids B. oleracea  O. violaceus had the sum of parental chromosomes (2n ¼ 21) in mitotic and meiotic cells. B. rapa  O. violaceus hybrids were mixoploid with somatic chromosome number ranging from 23 to 42 but cells with 2n ¼ 34 predominating. Partial separation of parental genomes occurred during mitosis, leading to the addition of some Orychophragmus chromosome to the B. rapa complement. Hybridization with B. nigra produced a majority of maternal type F1 plants (2n ¼ 16) and some mixoploids. Hybrids with the three tetraploid species showed variable chromosome numbers: B. carinata  O. violaceus (2n ¼ 12--34), B. juncea  O. violaceus (2n ¼ 30--42), and B. napus  O. violaceus (2n¼12–38). Partial and complete separation was more frequent in B. juncea  O. violaceus and B. carinata  O. violaceus hybrids respectively. Somatic cells and PMCs with additional O. violaceus chromosomes often occurred in B. juncea  O. violaceus and not in other two combinations. It was proposed that differences in the duration of somatic cell cycles of two parents cause partial or complete genome elimination. Based on cytological observations, Li and Heneen (1999) and Li et al. (2003) proposed that B genome accounts for complete and partial genome separation in B. carinata; both A and B genomes contribute to this separation in B. juncea; and A genome is more influential than C genome in B. napus during mitosis and meiosis. Genetic information from Orychophragmus has been introgressed into Brassica genomes as demonstrated by GISH (Hua and Li 2006). Employing these hybridizations, it may be possible to produce Brassica aneuploids and haploids and subsequently homozygous lines (see review by Li and Ge 2007). B. Somatic Hybrids The majority of the wild Brassica species belong to the secondary and tertiary gene pool, which renders their genes inaccessible for crop

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85

improvement. As mentioned earlier, barriers to sexual hybridization are easily overcome through the somatic route. Recent years have seen spectacular developments in protoplast fusion technology, particularly in Brassicaceae. Also, Brassica and related genera are very amenable to tissue culture techniques. Cell fusion allows cytoplasmic substitutions and generation of novel cytoplasmic variability through organellar reassortment and DNA recombination, a phenomenon not possible during sexual hybridization. Because of these advantages, cell fusion has become a promising methodology for introgressing desirable alien genes in crop cultivars (see reviews by Glimelius 1999a; Christey 2004; Navra´tilova´ 2004; Liu et al. 2005). The first successful report of cell fusion in Brassicacea was by Kartha et al. (1974) and involved protoplasts of B. napus and Glycine max. A major breakthrough was made by Gleba and Hoffmann (1979, 1980) when, following fusion of B. rapa and Arabidopsis thaliana protoplasts, an intertribal hybrid was successfully regenerated. This event achieved the distinction of first somatic hybrid in Brassicaceae, although no offspring could be obtained from it. Subsequently, a large number of somatic hybrids have been obtained that combine crop species with taxonomically divergent wild germ pools (Tables 2.14, 2.15). These represent interspecific, intergeneric, and a substantial number of intertribal combinations from six different tribes— Sisymbrieae (Arabidopsis thaliana, Camelina sativa); Arabideae (Armoracia rusticana, Barbarea stricta, B. vulgaris); Drabeae (Lesquerella fendleri); Lepidieae (Capsella bursapastoris, Lepidium, Thlaspi caerulescens, T. perfoliatum); Lunarieae (Lunaria annua); and Hesperideae (Matthiola incana)—and a few subtribes— Raphananiae (Raphanus, Trachystoma), and Moricandiineae (Moricandia). Another species, Orychophragmus violaceus, previously included in subtribe Moricandiinae but now excluded from tribe Brassiceae (Go´mez-Campo, personal communication), has also been hybridized. The priorities have shifted to practical utilization and efforts are toward introgressing nuclear and cytoplasmic genes from wild relatives to crop species. The desirable traits targeted include:  C3–C4 intermediate photosynthetic system (from Moricandia arvensis and M. nitens)  Resistance to: club root (from Raphanus sativus); alternaria leaf spot (from Sinapis alba, Camelina sativa, Capsella bursa-pastoris); beet cyst nematodes (from Sinapis alba, C. bursa–pastoris); blackleg (from B. tournefortii, Sinapis arvensis, Arabidopsis thaliana)  High nervonic acid content (from Thlaspi perfoliatum)  High erucic acid content (from Crambe abyssinica)

86

Crambe abyssinica (n ¼ 45) þ B. napus

Armoracea rusticana (n ¼ 16) þ B. oleracea Barbarea vulgaris (n ¼ 8) þ B. oleracea Barbarea vulgaris (n ¼ 8) þ B. rapa Barbarea vulgaris (n ¼ 8) þ B. napus Barbarea stricta (n ¼ 8) þ B. rapa Camelina sativa (n ¼ 20) þ B. oleracea Camelina sativa (n ¼ 20) þ B. carinata Capsella bursa–pastoris (n ¼ 16) þ B. oleracea

Arabidopsis thaliana (n ¼ 5) þ B. rapa Arabidopsis thaliana (n ¼ 5) þ B. juncea Arabidopsis thaliana (n ¼ 5) þ B. napus

Arabidopsis thaliana (n ¼ 5) þ B. oleracea

Arabidopsis thaliana (n ¼ 5) þ B. nigra

Somatic hybrid

Experimental demonstration Phosphinothricin resistance Herbicide resistance, black leg resistance Transposable element Spm/dSpm Clubroot resistance Cold tolerance Cold tolerance Cold tolerance Cold tolerance Alternaria resistance Alternaria resistance Resistance to flea beetles, alternaria blight High erucic acid content, insect resistance

Resistance to flea beetles, cold tolerance, short life cycle Plastome transformation

Desirable trait to introgress

Table 2.14. Intertribal somatic hybrids in Brassiceae.

Wang et al. 2003,* 2004a*

Nitovskaya and Shahkhovskii 1998; Yamagishi and Nakagawa 2004; Nitovskaya et al. 2006a Gleba and Hoffmann 1979, 1980 Ovcharenko et al. 2004 Bauer-Weston et al.1993;* Forsberg et al. 1994, 1998;* Yamagishi et al. 2002* Ovcharenko et al. 2005* Navra´tilova´ et al. 1997 Ryschka et al. 1999 Oikarinen and Ryo¨ppy 1992 Fahleson et al. 1994b Oikarinen and Ryo¨ppy 1992 Hansen 1998; Sigareva and Earle 1999a Narasimhulu et al. 1994 Nitovskaya et al. 1998; Sigareva and Earle 1999b

Siemens and Sacrista´n 1995*

Reference

87

*Asymmetric. **Both symmetric and asymmetric.

Thlaspi perfoliatum (n ¼ 21) þ B. napus Thlaspi caerulescens (n ¼ 7) þ B. napus Thlaspi caerulescens (n ¼ 7) þ B. juncea

Lunaria annua (n ¼ 14) þ B. napus Matthiola incana (n ¼ 7) þ B. oleracea Orychophragmus violaceus (n ¼ 12) þ B. napus

Lepidium meyenii (n ¼ 32) þ B. oleracea Lesquerella fendleri (n ¼ 6) þ B. napus

Descurainia sophia (n ¼ 14) þ B. napus High linolenic acid content, cold tolerance Glucosinolate content High lesquerolic acid content, drought tolerance, Lesquerella chloroplasts High nervonic acid content Oil quality High linoleic and palmitic acid content, phosphinothrin resistance Chlorosis correction High nervonic acid content Zinc and cadmium tolerance High metal accumulation Vasilenko et al. 2003 Fahleson et al. 1994a** Brewer et al. 1999 Dushenkov et al. 2002

Ryschka et al. 2003 Skarzhinskaya et al. 1996,** 1998; Schro¨der-Pontoppidan et al. 1999 Nitovskaya et al. 2006b Craig and Millam 1995 Ryschka et al. 1999; Sheng et al. 2008 Hu et al. 1999; Hu et al. 2000b;* Sakhno et al. 2007

Guan et al. 2007

88

Moricandia arvensis (n ¼ 14) þ B. juncea Moricandia arvensis (n ¼ 14) þ B. napus Moricandia nitens (n ¼ 14) þ B. oleracea Moricandia nitens (n ¼ 14) þ B. rapa Moricandia nitens (n ¼ 14) þ B. napus Raphanus sativus (n ¼ 9) þ B. oleracea

Intergeneric Diplotaxis catholica (n ¼ 9) þ B. juncea Diplotaxis harra (n ¼ 13) þ B. juncea Diplotaxis harra (n ¼ 13) þ B. napus Diplotaxis muralis (n ¼ 21) þ B. juncea Eruca sativa (n ¼ 11) þ B. juncea Eruca sativa (n ¼ 11) þ B. napus Moricandia arvensis (n ¼ 14) þ B. oleracea

Interspecific Brassica spinescens (n ¼ 8) þ B. juncea Brassica tournefortii (n ¼ 10) þ B. napus

Somatic hybrids

3 IV þ 33 II 1 IV þ 27 II

27 II–2 IV þ 23 II 60, 62, 1–4 IV þ 23 II

25, 36, 62 36

64, 3 III þ 15 II þ 25 I 66 46, 74, 92

54, 58, 64 78, 58, 60 46

52, 26 II 58

2n and meiotic pairing

sterile — fertile 0–50 fertile — —

0–56 sterile — sterile 0–8 8–98

sterile 0–31 0–43

sterile — fertile 0–50 fertile very low very low

0–7 sterile — sterile very poor 0–28

sterile poor poor

Seed

Fertility (%) Pollen

Table 2.15. Interspecific and intergeneric somatic hybrids in coenospecies.

Kirti et al. 1995c Begum et al. 1995 Klimaszewska and Keller 1988 Chatterjee et al. 1988 Sikdar et al. 1990 Fahleson et al. 1988, 1997 Toriyama et al. 1987a; Sobrino-Vesperinas 1988; Ishikawa et al. 2003 Kirti et al. 1992b O’Neill et al. 1996 Meng et al. 1999 Meng et al. 1999 Meng et al. 1999 Kameya et al. 1989 Hagimori et al. 1992; Yamanaka et al. 1992

Kirti et al. 1991 Stiewe and Ro¨bbelen 1994 Liu et al. 1995

Reference

89

arvensis (n ¼ 8) þ arvensis (n ¼ 8) þ arvensis (n ¼ 8) þ alba (n ¼ 12) þ B.

B. nigra B. oleracea B. napus oleracea

Trachystoma ballii (n ¼ 8) þ B. juncea

Sinapis alba (n ¼ 12) þ B. juncea Sinapis alba (n ¼ 12) þ B. napus

Sinapis Sinapis Sinapis Sinapis

Raphanus sativus (n ¼ 9) þ B. juncea Raphanus sativus (n ¼ 9) þ B. napus

66–77 sterile

56–90, 30–44 II þ 5 I–10 I 62, 31 II 52, 26 II–2 II þ 22 II þ 2 I

sterile sterile — — low

sterile 43–63 — — 23–58

good sterile

sterile low very low

sterile sterile

sterile 5–7

sterile 0–20 0–3

60, 30 II

56, 26 ii 32 34 54, predominantly 27 II 42

54, 56, 70, 26–30 II

Muller et al. 2001 Sakai and Inamura 1990; Sundberg and Glimelius 1991; Lelivelt and Krens 1992 Muller et al. 2001 Wang et al. 2006b Toriyama et al. 1987b Toriyama et al. 1987b Hu et al. 2002a Bauer 1990; Ryschka et al. 1996; Hansen and Earle 1997; Sigareva et al. 1999 Gaikwad et al. 1996 Primard et al. 1988 Lelivelt et al. 1993 Chevre et al. 1994a Wang et al. 2005b Kirti et al. 1992a

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 High lesquerolic acid content (from Lesquerella fendleri)  High linoleic and palmitic acid content (from Orychophragmus violaceus)  Cold tolerance (from Barbarea vulgaris)  Zinc and cadmium hyperaccumulation (from Thlaspi carulesens)  Cytoplasmic genes for inducing male sterility from a number of wild species Somatic hybrids in several combinations—for example, Camelina sativa þ B. carinata (Narasimhulu et al. 1994), Camelina sativa þ B. oleracea (Hansen 1998) and Barbarea vulgaris þ B. napus (Fahleson et al. 1994b)—could not be established to viable field plants. It appears that although protoplast fusion removes fertilization barriers, genetic incompatibilities due largely to phylogenetic distances still prevail at the somatic level, affecting differentiation, growth, and development of normal plant parts, particularly floral organs, thus leading to sterility. However, other distant hybrids, particularly with Arabidopsis thaliana, probably could be established successfully, due to its small genome and also with little repetitive DNA, which promotes greater homoeology between the partaking genomes (Hansen 1998). Lunaria annua þ B. napus hybrid has been reported only up to callus stage (Craig and Millam 1995). Somatic hybrids have been identified and characterized by a range of techniques including morphological attributes, chromosome number, meiotic behavior, fertility, DNA content estimation, isozyme analysis, RFLP, AFLP, and cytoplasmic constitution. However, there are not many reports on chromosome cytology, and in several studies, the ploidy status has been determined by estimating DNA content. Somatic hybrids, in general, are intermediate in morphology between the fusion partner species. This expression is particularly relevant for leaves and frequently for flower characteristics. Floral abnormalities are also observed and include 6 to 8 petals and multiple carpellike structures in A. thaliana þ B. napus (Bauer-Weston et al. 1993); 1 or 2 petals in Thlaspi perfoliatum þ B. napus (Fahleson et al. 1994a); enlarged, distorted or globular pistils, and reduced or missing stamens in L. fendleri þ B. napus (Skarzhniskaya et al. 1996); stamens with stunted filaments in R. sativus þ B. napus (Lelivelt et al. 1992); and shorter, thicker pistils in D. harra þ B. napus (Klimaszewska and Keller 1988). Chromosome cytology revealed that somatic hybrids either have the full chromosome complements of both parents, which are derived mostly from symmetric fusions, or spontaneous chromosome elimination occurs leading to generation of asymmetric hybrids or

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cybrids (Navara´tilova´ et al. 1997; Hu et al. 2002a). Sometimes the regenerants from the same fusion events have different chromosome numbers (Hoffman and Adachi 1981). Meiotic studies have been carried out in some of these hybrids which include intergeneric and a few intertribal combinations. Besides occurrence of normal bivalents as the sum of parental chromosomes at M1, higher associations such as tri- and quadrivalents, in addition to univalents, were also encountered (Table 2.15). Interestingly, the intersubtribal hybrid Moricandia arvensis þ B. juncea exhibits up to three quadrivalents (Kirti et al.1992b). Such higher associations suggest intergenomic chromosome homoeology. Post-metaphase-1 stages have not been investigated, but it appears that meiosis proceeds normally, as can be inferred from normal pollen formation in many of the hybrids. Intergenomic chromosome recombination due to allosyndesis has been documented in some somatic hybrids, such as Moricandia arvensis þ B. juncea, D. catholica þ B. juncea, Trachystoma ballii þ B. juncea. Genomic in situ hybridization has been used effectively to determine the alien chromosome status at mitosis in some somatic hybrids and their progeny, for example, in Eruca sativa þ B. napus (Fahleson et al. 1988, 1997), Lesquerella fendeleri þ B. napus (Skarzhniskaya et al. 1996), Crambe abyssinica þ B. napus (Wang et al. 2004a,b), and Sinapis alba þ B. napus (Wang et al. 2005a). GISH was also employed to dectect intergenomic homoeologous recombination in these hybrids. A majority of somatic hybrid plants were seed sterile when selfed. However, some fertile hybrids were also obtained, including intertribal hybrids Arabidopsis thaliana þ B. napus (Forsberg et al. 1994), Thlaspi perfoliatum þ B. napus (Fahleson et al. 1994a), Capsella bursa-pastoris þ B. oleracea (Sigareva and Earle 1999b), Orychophragmus violaceus þ B. napus (Hu et al. 2002b), and Moricandia arvensis þ B. oleracea (Ishikawa et al. 2003), and a few intergeneric and interspecific ones. Wherever pollen fertility was observed, it was quite low in A1 generation. With a few exceptions seed fertility was invariably very poor, and seeds could be obtained only on backcrossing. Only in rare instances were seeds obtained on selfing. A correlation between the presence of alien chromosomes and fertility was observed in somatic hybrids Lesquerrella fendleri þ B. napus (Skarzhinskaya et al. 1998) and Crambe abyssinica þ B. napus (Wang et al. 2004b); with the decreasing number of alien chromosomes, the fertility of the hybrids increased and plants possessing the entire alien chromosome complement were absolutely sterile. Similar correlations were obtained in asymmetric hybrids A. thaliana þ B. napus (Forsberg et al.

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1998) and Orychophragmus violaceus þ B. napus (Hu et al. 2002b). In many instances, to produce reasonably fertile hybrids, irradiated protoplasts from wild species have been used, eliminating substantial amounts of alien DNA to obtain asymmetric hybrids that contain varying amounts of alien DNA from donor species. Somatic hybrids present three possibilities with respect to their cytoplasmic genomes: (1) parental genomes segregate to homogeneity during cell division, (2) both the parental genomes occur as a mixed population, and (3) novel genome constitution is generated when parental genomes undergo recombination. Segregation of chloroplasts is independent of mitochondrial segregation. In Brassiceae, mitochondrial recombination has been observed to occur frequently and is very well documented (Glimelius 1999a). In sharp contrast, intergenomic chloroplast recombination is rare. Two chloroplast types occurring in mixture is also rare, and there is no information about whether this mixture persists in subsequent generations, wherever it does occur. It is observed in interspecific, intergeneric, and intertribal somatic hybrids that chloroplast from crop species are generally favored. This biased segregation is attributed to genetic divergence, ploidy level differences between the parental species, and rate of chloroplast division (Sundberg and Glimelius 1991). Also, plastome-genome incompatibility may be a factor. A higher ploidy level of one of the parental species contributes a larger number of chloroplasts per cell (Butterfass 1989). Since in most of the hybrid alloploid species B. napus or B. juncea has been one of the parents, they contribute more chloroplasts to the fusion products than the wild diploid parent. However, it is not possible to predict which parental chloroplast will establish in hybrids. The intertribal somatic hybrid Lesquerella fendleri þ Brassica napus has been reported to have mixed chloroplasts (Skarzhinskaya et al. 1996), as has intergeneric hybrid Diplotaxis catholica þ B. juncea (Mohapatra et al. 1998). A report documented the occurrence of intergeneric chloroplast recombination in the somatic hybrid Trachystoma balli þ B. juncea (Baldev et al. 1998). where the recombination has occurred in a single copy region and remains stable over the generations. Also, it caused no imbalance in the recombinant plastomes in terms of chloroplast-related functions. In addition, choroplast recombination was also indicated in the somatic hybrid B. oleracea þ Raphanus sativus (Kanno et al. 1997). Mitochondrial genomes behave differently from plastid genomes. Besides segregation and mixture of parental types, intergenomic mitochondrial recombination is frequent in Brassiceae somatic hybrids. The recombinant mitochondrial genome contains DNA fragments of both the

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parents or entirely new and unique ones not found in parental types (Belliard et al. 1979). While investigating seven sets of interspecific, intergeneric, and intertribal combinations, Landgren and Glimelius (1994b) observed that 43% to 95% of the hybrids had mt DNA rearrangements. Recombination hot spots have also been found; for example, Mohapatra et al. (1998) suggested that intergenomic recombination is preferred at specific sites in somatic hybrids Diplotaxis catholica þ B. juncea. The cox2 coding region may serve as an active site for inter- or intragenomic recombination (Stiewe and Ro¨bbelen 1994; Liu et al 1995). Conflicting views are reported regarding the segregation of mitochondria in somatic hybrids. According to Landgren and Glimelius (1990, 1994a,b), crop types are favored. In cybrids where CMS line is one of the parents, the mt segregation was slightly biased toward the CMS parent (Mukhopadhyay et al. 1994; Liu et al. 1996). However, many of the somatic hybrids have recombinant mitochondrial genomes, although a lack of recombination has also been documented, for example, S. alba þ B. napus (Lelivelt et al. 1993), Lesqurella fendleri þ B. napus (Skarzhinskaya et al. 1996), and Moricandia arvensis þ B. oleracea (Ishikawa et al. 2003). Somatic hybridization in Brassicaceae has crossed all the intergeneric and intertribal barriers. However, the results are not too encouraging because of a general high degree of sterility or severe intergenomic incompatibilities leading to many abnormalities. Asymmetric hybrids in such instances appear to be more promising as crop species, tolerating only a fraction of alien genetic content rather the whole genome for integrated functioning of the system. Such asymmetric fusions have been obtained by irradiating donor (wild) protoplasts to induce doublestrand DNA breaks. Most of intertribal hybrids are asymmetric and show improved fertility. One of the limiting factors in gene transfer from wild to crop species is the very low level or complete absence of intergenomic chromosome pairing, which implies that overall genome structures interfere with free gene flow across the generic boundaries. Nevertheless, several traits of agronomic importance have been observed in somatic hybrids and in some cases, the genes have been introgressed to crop species, as revealed by progeny plant analysis. Examples include: 1. Raphanus sativus þ B. napus: express resistance to beet cyst nematode—Heterodera schachtii (Lelivelt and Krens 1992) 2. Sinapis alba þ B. napus: possess high level of beet cyst nematode (Heterodera schachtii) resistance (Lelivelt et al. 1993) 3. Thlaspi perfoliatum þ B. napus: contain significantly greater amounts of nervonic acid (Fahleson et al. 1994a)

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S. PRAKASH, S. R. BHAT, C. F. QUIROS, P. B. KIRTI, AND V. L. CHOPRA

4. B. tournefortii þ B. napus: express resistance to blackleg (Phoma lingum) (Liu et al. 1995) 5. Moricandia arvensis þ B. napus: C3-C4 character is expressed at both the physiological and anatomical level (O’Neill et al. 1996) 6. Moricandia nitens þ B. oleracea: C3–C4 character expressed as transition between the parents (Yan et al. 1999) 7. Sinapis alba þ B. oleracea: exhibit resistance to Alternaria brassicicola and Phoma lingam (Ryschka et al. 1996; Hansen and Earle 1997; Sigareva et al. 1999) 8. Capsella bursa-pastoris þ B. oleracea: exhibit high degree of resistance to Alternaria brassicicola (Sigereva and Earle 1999b) 9. Thlaspi caerulescens þ B. napus: accumulate high levels of zinc and cadmium, which would have been toxic to B. napus (Brewer et al. 1999) 10. Camelina sativa þ B. oleracea: possess resistance to Alternaria (Sigareva and Earle 1999a ) 11. Lesquerella fendleri þ B. napus: contain high amount of erucic acid for industrial purpose (Glimelius 1999b; Schro¨der-Pontokpidan et al. 1999) 12. Arabidopsis thaliana þ B. napus: possess resistance to Leptosphaeria maculans (Bohman et al. 2002) 13. Orychophragmus violaceus þ B. napus: contain high content of palmitic and linoleic acid expressed in the progeny plants (Hu et al. 2002b; Ma and Li 2007) 14. Sinapis avensis þ B. napus: possess resistance to blackleg in the hybrids and progeny (Hu et al. 2002a) 15. Crambe abyssinica þ B. napus: progeny contain high amount of seed erucic acid (Wang et al. 2004b) Several CMS systems in B. napus and B. juncea have been obtained following protoplast fusion. These are based on Arabidopsis thaliana, Brassica tournefortii, Diplotaxis catholica, Eruca sativa, Moricandia arvensis, Orychophragmus violaceus, Raphanus, Sinapis arvensis, and Trachystoma ballii. C. Introgression of Genes A number of bacterial, fungal, and viral diseases cause severe infection on Brassica crops and inflict heavy losses to seed yield. Among them, black leg or stem canker (Leptosphaeria maculans/Phoma lingam),

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95

alternaria leaf spot (Alternaria spp.), white rust (Albugo candida), black rot (Xanthomonas campestris pv. campestris), soft rot (Erwinia carotovora), and sclerotinia stem rot (Sclerotinia sclerotiorum) are important. Nuclear genes conferring resistance to these diseases have been transferred from related species and alien wild germplasm. Other desirable traits, particularly the fertility restoration for several CMS systems, have also been incorporated (Table 2.16). These genes have been introgressed, taking advantage of nonhomologous allosyndetic recombination in early backcross generations following sexual/somatic hybridizations and also through generating chromosome addition lines. In recent years, efforts have been made to identify alien introgressions to specific chromosomes through GISH and molecular markers. Results of GISH are not very encouraging primarily due to unusually low copy number of repeat sequences in chromosome arms, which form the basis of GISH signals. Nevertheless, examples of detecting introgressions include B. napus from Lesquerella fendleri (Skarzhinskaya et al. 1998), Raphanus sativus (Voss et al. 2000), Sinapis arvensis (Snowdon et al. 2000b), Crambe abyssinica (Wang et al. 2004b), and Orychophragmus violaceus (Li and Ge 2007).

V. CYTOPLASMIC SUBSTITUTION AND MALE STERILITY During the last 50 years, several investigations have reported the expression of a high degree of heterosis for seed yield in intervarietial hybrids of B. rapa, B. juncea, and B. napus (see Fu and Yang 1998). However, in earlier years, full potential of heterosis could not be exploited in B. juncea and B. napus as these are predominantly selffertilized crops. A suitable pollination control mechanism is required to produce commercial hybrid seed. A cytoplasmic male sterility (CMS) fertility restoration system is an excellent potential means to facilitate hybridization because it is easy to maintain. CMS, a maternally inherited inability to produce fertile pollen, is encoded in the mitochondrial genome and can arise spontaneously due to mutation in the genome (autoplasmy) or can be expressed following cytoplasmic substitutions due to nuclear-mitochondrial incompatibility (alloplasmy). A sterility inducing cytoplasm was identified in a wild population of Raphanus sativus in 1968 (Ogura 1968), and the first alloplasmic in Brassica was developed by introgressing this sterility inducing cytoplasm into B. napus and B. oleracea (Bannerot et al. 1974). Around that time, Pearson (1972) also developed male sterile broccoli

96

Black leg resistance

Club root resistance

Chlorosis correction Beet cyst nematode resistance

B. napus B. rapa B. juncea

Raphanus sativus

Raphanus sativus Sinapis alba

Enarthrocarpus lyratus

Raphanus sativus Raphanus sativus B. juncea B. tournefortii Trachystoma ballii Moricandia arvensis Erucastrum canariense

Fertility restoration

rapa juncea/B. carinata carinata rapa/B. carinata rapa/B. juncea

B. B. B. B. B.

Donor species

Yellow seed coat color

Trait napus napus napus napus napus

B. oleracea var. capitata B. napus B. napus

B. napus

CMS (Ogu?) B. napus CMS (Kosena) B. napus CMS (Polima) B. napus CMS (Tour) B. napus CMS (Trachy) B. juncea CMS (Moricandia) B. juncea CMS (Canariense) B. juncea CMS (Canariense) B. napus CMS (Lyratus) B. rapa CMS (Lyratus) B. juncea/ B. napus CMS (Ogu) B. napus B. napus

B. B. B. B. B.

Recepient species

Table 2.16. Introgression of nuclear genes conferring desirable traits.

Lelivelt and Krens 1992; Voss et al. 2000; Peterka et al. 2004; Budahn et al. 2006 Chiang et al. 1977, 1980 Gowers 1982 Roy et al. 1984; Sacrista´n and GerdemannKno¨rck 1986; Chevre et al. 1997a; Dixelius 1999

Paulmann and Ro¨bbelen 1988 Lelivelt et al. 1993

Chen and Heneen 1992 Rashid et al. 1994 Qi et al. 1995 Meng et al. 1998 Rahman et al. 2001; Potapov and Osipova 2003 Heyn 1978; Rousselle and Dosba 1985 Sakai et al. 1996 Fan et al. 1986 Stiewe and Ro¨bbelen 1994 Kirti et al. 1997 Prakash et al. 1998 Prakash et al. 2001 Banga et al. 2003b Deol et al. 2003 Banga et al. 2003a

Reference

97

Earliness

Altered oil quality

Black rot resistance Soft rot resistance

Alternaria leaf spot resistance

B. napus B. napus B. napus B. napus B. napus B. napus B. napus B. oleracea; B. rapa ssp. pekinensis B. napus B. napus B. napus

Arabidopsis thaliana

Sinapis arvensis

Coincya monensis B. rapa Sinapis alba

Diplotaxis erucoides B. juncea B. oleracea var. italica

Orychophragmus violaceus Lesquerella fendleri B. rapa

B. napus

B. nigra

Hu et al. 2000b; Hu and Li 2006 Du et al. 2008 Shiga 1970; Namai et al. 1980

Klewer et al. 2003 Tonguc and Griffiths 2004 Ren et al. 2000, 2001

Struss et al. 1996; Chevre et al. 1996, 1997b; Plieske et al. 1998; Dixelius 1999 Bohman et al. 2002; Ogbonnaya et al. 2003; Saal et al. 2004 Snowdon et al. 2000b: Hu et al. 2002a: Winter et al. 2003 Winter et al. 2003 Chevre et al. 2003; Leflon et al. 2007 Primard et al. 1988

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S. PRAKASH, S. R. BHAT, C. F. QUIROS, P. B. KIRTI, AND V. L. CHOPRA

(B. oleracea var. italica) by placing its nucleus in B. nigra cytoplasm. A large number of alloplasmics have been reported since. Brassica coenospecies is a rich repository of diverse mitochondrial genomes, as revealed by RFLP studies (Pradhan et al. 1992). By combining these cytoplasms with crop nuclei, a spectrum of alloplasmic lines of diverse origin expressing male sterility has been obtained, particularly in B. juncea (Table 2.17) (see reviews by Delourme and Budar 1999; Prakash 2001; Budar et al. 2004). Cytoplasmic male sterile lines have been developed following backcrossings of either sexually synthesized allopolyploids or somatic hybrids between wild and crop species. Somatic hybridization for the synthesis of an alloplasmic was attempted for the first time by Kameya et al. (1989) when they combined the nucleus of B. oleracea with Raphanus cytoplasm. Subsequently, it has been employed extensively to obtain new combinations. As expected, CMS originating from sexual hybridizations possess unaltered organellar genomes because of exclusive maternal inheritance. Since organelle assortment and intergenomic mitochondrial recombinant is of frequent occurrence in Brassiceae, the cytoplasmic constitution is entirely different in those originating from somatic hybrids, and various possible combinations of mitochondrial and chloroplast genomes have been reported in different CMS lines (Table 2.17). Alloplasmic CMS plants, in general, are similar to euplasmic plants in development and morphology. However, many of them exhibit abnormalities, both developmental and floral, which are manifestations of alien cytoplasms. Varying degrees of leaf chlorosis, ranging from mild to severe, occurs in several systems, such as Raphanus/Ogu, Oxyrrhina, Tournefortii, Moricandia, and Enarthrocarpus. Floral abnormalities are also common in male sterile plants and are expressed in several ways: petaloid anthers (Nigra, Muralis, Trachystoma, Raphanus, Tournefortii, Canariense); poor or absent nectaries (Tournefortii and Raphanus); crooked style (Tournefortii, Raphanus); thick pistil (Raphanus); and low seed fertility (Raphanus, Tournefortii, Enarthrocarpus and Trachystoma). Leaf chlorosis is an expression of incompatible interaction between alien chloroplasts and endogenous nucleus. Agronomically, it is an undesirable feature associated with reduced growth and adversely affects productivity. Similarly, it has clearly been demonstrated at least in Brassica that nuclearmitochondrial interaction is critical for flower development, normal morphology, and subsequent seed fertility (see Glimelius 1999a). Application of biotechnology, particularly protoplast fusion methodology, has created entirely new opportunities for improving these

99

B. rapa B. napus

B. rapa B. juncea B. napus

B. oleracea Somatic hybrid R. sativus var. capitata þ B. oleracea

Diplotaxis muralis

Brassica oxyrrhina

Raphanus sativus

Sexual alloploid B. oxyrrhina  B. rapa

Sexual alloploid D.muralis  B. rapa

Sexual alloploid B. nigra  B. oleracea var. italica

B. olerecea var. italica

Spontaneous mutation in Raphanus mitochondrial genome

Brassica nigra

oleracea rapa juncea napus

Origin

B. B. B. B.

Recepient species

Raphanus sativus (Ogura)

Cytoplasmic donor

Table 2.17. Cytoplasmic substitution lines in Brassiceae.

(continued)

Kameya et al. 1989

Prakash and Chopra 1988, 1990; Kirti et al. 1993

Hinata and Konno 1979; Pellan-Delourme and Renard 1987

Pelletier et al. 1983; Jarl and Bornman 1988; Jarl et al. 1989; Kao et al. 1992; Kirti et al. 1995a Pearson 1972

Abnormalities removed following protoplast fusion.

Leaves green, flowers of two types: (1) petaloid anthers and absent nectaries; (2) rudimentary anthers. Poor seed fertility. cp and mt: B. nigra Flowers with narrow petals and occasionally with petaloid anthers. Nectaries very poorly developed. Female fertility low. cp and mt: D. muralis Flowers normal with nondeshicent anthers and excellent nectaries. Female fertility: 96%. Cp and mt: B. oxyrrhina. Leaf chlorosis in B. juncea rectified following protoplast fusion Plants similar to B. oleracea in floral morphology and female fertility. cp: R. ativus

Ogura 1968; Bannerot et al. 1974; Mathias 1985

Reference

Leaves highly chlorotic. Flowers with petaloid anthers or absent nectaries. Female fertility poor.

Characteristics

100

B. napus

B. napus

B. juncea B. napus

B. juncea

B. juncea B. napus

Brassica tournefortii

Diplotaxis siifolia

Trachystoma ballii

Moricandia arvensis

Recepient species

Raphanus sativus (Kosena)

Cytoplasmic donor

Table 2.17. (Continued) Characteristics

Plants green. Flowers with stunted filaments and aborted anthers. Female fertility normal. mt: recombined Somatic hybrid Leaves green. Flowers with normal B. tournefortii or without petals and reduced þ B. napus empty anthers. Female fertility normal. cp: B. napus, mt: recombined Sexual alloploid Leaves green. Flowers with reduced D. siifolia  B. juncea anthers and well-developed nectaries. Female fertility 95 %. cp and mt: D. siifolia Somatic hybrid Leaves green. Flowers with 2 types T. ballii þ B. juncea; of anthers: petaloid and selender cp: B. juncea mt: reco nondehiscent. Female fertility 30–40%. cp: B. juncea; mt: recombined Somatic hybrid M. ‘arvensis Leaves highly chlorotic almost þ B. juncea yellowish. Delayed flowering. Normal flowers with slender anthers and excellent nectaries. Female fertility 96%. cp and mt: M. arvensis. Chlorosis rectified following protoplast fusion.

Somatic hybrid Raphanus sativus þ B. napus

Origin

Prakash et al. 1998 Kirti et al. 1998

Kirti et al. 1995b

Rao et al. 1994; Rao and Shivanna 1996

Stiewe and Ro¨bbelen 1994; Liu et al. 1996

Sakai and Inamura 1990

Reference

101

B. juncea

B. juncea

B. rapa

B. juncea B. napus

B. juncea

B. juncea

Raphanus sativus

Diplotaxis erucoides

Diplotaxis berthautii

Eruca sativa

Erucastrum canariense

Diplotaxis catholica

Diplotaxis catholica

Moricandia arvensis

Sexual hybrid M. arvensis  R. sativus

Sexual hybrid D. catholica  B. juncea

Somatic hybrid D. catholica þ B. juncea

Sexual alloploid E. canariense  B. rapa

Sexual hybrid E. sativa  B. rapa

Sexual hybrid D. berthautii  B. rapa

Sexual hybrid D. erucoides  B. rapa

Leaves normal green. Flowers with smaller slender anthers and excellent nectaries. Female fertility 96%. cp and mt: D. erucoides Leaves green, flowers with short indehiscent or petaloid anthers. Female fertility 95%. cp and mt: D. berthautii Leaves green. Abnormal flowers with petaloid, partial petaloid or slender anthers. Poorly developed nectaries. Female fertility low. cp and mt: E. sativa Leaves normal green. Flowers with slender anthers and excellent nectaries. Female fertility 95%. cp and mt: E. canariense Leaves green, flowers with small rudimentary anthers and excellent nectaries. Female fertility 96%. cp: B. juncea, mt: recombined Leaves green. Flowers with petaloid/tubular structures. Nectaries poorly developed. Abnormal style. Female fertility 40%. cp and mt: D. catholica Leaves light green. Flowers with well-developed anthers and nectaries. cp and mt: M. arvensis Bang et al. 2002

(continued)

Mohanty 1996; Pathania et al. 2003

Prakash 2001; Pathania et al. 2007

Prakash et al. 2001; Banga et al. 2003b

Matsuzawa et al. 1999

Malik et al. 1999; Bhat et al. 2008

Malik et al. 1999; Prakash 2001; Bhat et al. 2006

102

B. napus

B. napus

Arabidopsis thaliana

Orychophragmus violaceus Sinapis arvensis

B. napus

B. rapa B. juncea B. napus

Recepient species

Enarthrocarpus lyratus

Cytoplasmic donor

Table 2.17. (Continued)

Somatic hybrid O. violaceus þ B. napus Somatic hybrid S. arvensis þ B. napus

Somatic hybrid A. thaliana þ B. napus

Sexual hybrid E. lyratus  B. rapa

Origin Leaves pale green. Delayed flowering. Flowers small with narrow petals having petaloid or rudimentary anthers. Female fertility poor. cp and mt: E. lyraus Plants normal green. Two types of flowers: (1) feminized anthers and abnormal pistils and (2) reduced shrunken anthers. Normal nectaries. Female fertility normal. mt: recombined Plants green. Flowers with reduced anthers. Female fertility normal. Plants green. Flowers normal with slender anthers. Female fertility normal.

Characteristics

Mei et al. 2003; Hu et al. 2004

Mei et al. 2003

Leino et al. 2003

Deol et al. 2003; Banga et al. 2003a; Janeja et al. 2003

Reference

2. BRASSICA AND ITS CLOSE ALLIES: CYTOGENETICS AND EVOLUTION

103

organelle associated traits, which are not likely to obtain through conventional sexual hybridization. Protoplast fusion allows the production of hybrid cytoplasm, commonly referred to as cybrid, that often has new and rearranged novel combinations of mitochondria and chloroplast genomes. Intergenomic mitochondrial recombination in Brassiceae somatic hybrids is a frequent phenomenon. Pelletier et al. (1983) were the first to demonstrate the efficacy of protoplast fusion for rectifying chlorosis through chloroplast substitution while retaining the male-sterility trait in CMS (Raphanus/Ogu) B. napus. Furthermore, some cybrids with better nectaries and atrazine resistance were obtained. This investigation induced researchers to transfer and combine sterility-inducing cytoplasm with chloroplast-encoded atrazine resistance and improve CMS systems through cytoplasmic hybridization in B. rapa, B. oleracea, and B. napus (Chetrit et al. 1985; Barsby et al. 1987; Robertson et al. 1987; Chuong et al. 1988; Jourdan et al. 1989a,b; Christey et al. 1991; Kao et al. 1992; Temple et al. 1992; Earle and Dickson 1994). Leaf chlorosis has been overcome in CMS (Raphanus/Ogu) B. napus (Pelleter et al. 1983; Menczel et al. 1987; Jarl and Bornman 1988; Jarl et al. 1989)), (Raphanus/Ogu) B. oleracea (Kao et al. 1992; Walters et al. 1992: Walters and Earle 1993), (Raphanus/ Ogu) B. juncea (Kirti et al. 1995a), (Oxyrrhina) B. juncea (Kirti et al. 1993; Arumugam et al. 2000), and (Moricandia) B. juncea (Kirti et al. 1998). In all these investigations, chlorosis correction was attributed to substitution of alien chloroplasts with those of crop chloroplasts. The cybrids are completely green; show normal growth and flowering; are fully male sterile and have normal female fertility. Because of maternal inheritance of the chloroplast genome, green character in CMS is stable in subsequent generations. Similarly, cold-tolerant CMS (Ogu) B. oleracea var. capitata was produced following protoplast fusion between cold-tolerant CMS broccoli and fertile cabbage (Sigareva and Earle 1997). In addition to these new cytoplasmic substitution lines, already existing sterile cytoplasms have been transferred following protoplast fusion. Examples include Polima CMS to broccoli (Yarrow et al. 1990), Nigra CMS to broccoli (Christey et al. 1991), and Tournefortii cytoplasm to rapid-cycling B. oleracea (Cardi and Earle 1997). It has been elegantly demonstrated in the Brassiceae that CMS plants with genetically reconstituted mitochondria obtained from protoplast fusion with less alien genetic content and least divergence from cultivated species have minimal floral abnormalities. Systems where such improved floral morphologies have been obtained include (Raphanus/Ogu) B. napus (Pelletier et al. 1983), B. oleracea (Kao et al. 1992), and B. juncea (Kirti et al. 1995a), (Tournefortii) B. napus (Stiewe

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and Ro¨bbelen 1994; Liu et al.1996), (Tournefortii) B. juncea (Arumugam et al. 1996), and (Arabidopsis) B. napus (Leino et al. 2003).

VI. GENOME DISSECTION AND DEVELOPMENT OF CHROMOSOME ADDITION LINES Chromosome addition lines have a major role in revealing genome organization and evolution, identifying gene linkage groups, assigning species-specific characters to a particular chromosome, and comparing gene synteny between related species. Localization of specific markers on individual chromosomes facilitates construction of genetic and cytogenetic maps. Their practical utilization lies in introgressing characters of agronomic value, particularly from alien species to crop cultivars. Several Brassica and related genomes— B. nigra, B. oleracea, B. rapa and B. oxyrrhina, Diplotaxis erucoides, Raphanus sativus, Sinapis alba, S. arvensis, Moricandia arvensis, Crambe abyssinica, Orychophragmus violaceus, and Arabidopsis thaliana—have been dissected using a series of monosomic addition lines (Table 2.18). Disomic additions have also been generated but only in a few instances for a specific chromosome as in (A. thaliana) B. napus–A. thaliana (Leino et al. 2004), B. napus–S. alba (Wang et al. 2005b) and B. napus–C. abyssinica (Wang et al. 2006a). The recently developed full set of nine disomic B. napus–R. sativus addition lines by Budhan et al. (2008) is the first complete disomic alien chromosome addition series in Brassicaceae. B. oleracea was the first genome to be dissected and is the most extensively studied. Addition lines generally do not show specific morphological phenotypes associated with a particular chromosome and are rarely distinguishable from one another, thus requiring additional markers for identification. It may well be that the recipient nuclear background masks the effect of added chromosome, or its effect is negated by homoeologous chromosome as these genomes evolved from a common archetype. Nevertheless, the added chromosomes sometimes exhibit peculiar morphological characters. For example, a radish chromosome addition in B. napus exhibits white flower color (Sernyk and Stefansson 1982). Chromosome 1 of Diplotaxis erucoides in B. napus was distinguished by light yellow color of their flowers (Chevre et al. 1994b). Also all additions of Sinapis alba in B. napus background possessed a long beak characteristic of S. alba (Wang et al. 2005b). Cytological investigations have been carried out in a number of studies. In general, the alien chromosome remained unpaired as a

105

B. rapa B. rapa

B. rapa B. rapa

B. rapa B. rapa B. rapa Raphanus sativus B. napus B. napus

B. oleracea B. oleracea

B. oleracea var. alboglabra B. oleracea var. alboglabra

B. B. B. B.

B. B. B. B. B. B.

nigra nigra nigra nigra nigra nigra

B. nigra B. nigra

napus napus napus napus napus oleracea

B. rapa

B. oleracea

B. B. B. B. B. B.

B. rapa

B. oleracea

oleracea var. alboglabra oleracea var. alboglabra oleracea var. alboglabra oleracea

B. rapa

Background genome

B. oleracea

Genome dissected

Disomics Monosomics, disomics Disomics Monosomics Monosomics Monosomics Monosomics Monosomics

Monosomics Monosomics Monosomics Monosomics

Monosomics Monosomics, disomics Monosomics Monosomics

Monosomics

Monosomics, disomics Monosomics

Nature of additions

Table 2.18. Major articles on chromosome addition lines.

Isozymes, fatty acids, RFLPs Isozymes, RFLPs, RAPDs Black leg resistance Black leg resistance, isozymes Isozymes, RAPDs Isozymes, RAPDs

Cytology Cytology

Cytology, isozymes Chromosome morphology, Isozymes, RAPDs Chromosome morphology Cytology, RAPDs FISH Morphology, RAPDs

Phenotypes, isozymes, RFLPs Chromosome morphology, isozymes RFLPs Cytology

Isozymes, rDNA

Markers

Chevre et al. 1991 Struss et al. 1992 Zhu et al. 1993 Chevre et al. 1996 Struss et al. 1996 Chevre et al. 1997b

Jahier et al. 1989 Struss et al. 1991

(continued)

Cheng et al. 1994a,b, 1995a Heneen and Jrgensen 2001 Hasterok et al. 2005b Kaneko et al. 1987, 1991, 1996, 2000

Chen et al. 1992 Chen et al. 1997a,b

McGrath et al. 1990 McGrath and Quiros 1990; Hu and Quiros 1991 Hosaka et al. 1990 Lee and Namai 1992

Quiros et al. 1987

Reference

106

Alloplasmic (Raphanus) B. napus Alloplasmic (Moricandia) Raphanus sativus B. napus B. napus

B. juncea

Raphanus sativus

Arabidopsis thaliana Crambe abyssinica

Orychophragmus violaceus

Moricandia arvensis

napus napus napus oleracea napus

B. B. B. B. B.

Diplotaxis muralis Sinapis arvensis Sinapis alba Sinapis alba Raphanus sativus

Brassica oxyrrhina

B. rapa

Diplotaxis erucoides

Background genome

Diplotaxis erucoides Raphanus sativus Alloplasmic (oxy) B. rapa B. napus

B. nigra

Genome dissected

Table 2.18. (Continued)

Disomics Monosomics, disomics Monosomics

Monosomics

Monosomics, disomics

Monosomics, disomics Monosomics Monosomics Monosomics Monosomics Monosomics

Monosomics

Monosomics

Monosomics

Nature of additions

GISH

GISH, RFLPs GISH

Morphology, cytology, RAPDs

RAPDs, FISH

Flower color, morphology, RAPDs Male sterility GISH, black leg resistance GISH GISH FISH, RFLPs

Morphology, RAPDs

RAPDs, leaf shape

Isozymes, RFLPs

Markers

Xu et al. 2007

Leino et al. 2004 Wang et al. 2006a

Bang et al. 2002

Fan et al. 1985 Snowdon et al. 2000b Wang et al. 2005b Wei et al. 2007 Peterka et al. 2004; Budahn et al. 2006 Budhan et al. 2008

Chevre et al. 1994b

Srinivasan et al. 1998

Kaneko et al. 2001, 2003

This et al. 1990

Reference

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univalent at metaphase 1 of meiosis. However, it underwent pairing also and formed a trivalent as in R. sativus–B. oleracea (Kaneko et al. 1987), B. rapa–B. oleracea (Chen et al. 1992; Heneen and Jrgensen 2001; Hasterok et al. 2005b), B. rapa–B. oxyrrhina (Srinivasan et al.1998), B. napus–S. alba (Wang et al. 2005b) and B. napus–Crambe abyssinica (Wang et al. 2006a). These associations reflected intergenomic homoeology between the added chromosome and recipient genome chromosomes. Using GISH, Wang et al. (2005b) observed homoeologous associations between S. alba and B. napus chromosomes, and in some cases recombinant chromosomes could clearly be identified. Hasterok et al. (2005b) identified B. oleracea chromosomes undergoing pairing with B. rapa chromosomes including chromosome C5 with an intercalary 5s rDNA locus and chromosomes C8 and C9 involving the regions occupied by 18S-5.8S–25S rRNA genes. On the contrary, in B. nigra additions on B. napus, only occasional chromosome pairing was observed (Jahier et al. 1989; Struss et al. 1991), reflecting the genetic distance between B. nigra and B. napus (AC) genomes as proposed earlier by several investigators. Transmission frequency of added chromosomes through male and female gametes does not follow a Mendelian pattern. Many factors, such as meiotic behavior of added chromosomes and their integrity (intact or recombined), genotype, and ploidy level of the donor and recipient species, affect transmission. Transmission frequency is generally far higher through the ovules than the pollen. Reduction in transmission frequency of added chromosomes due to competition with normal gametes was a common feature, leading to production of normal euploid type. Transmission of B. nigra additions was assessed using isozyme markers carried by different chromosomes. It was on an average 14% to 23% through ovules while the male transmission values ranged from 27% to 39% (Chevre et al. 1997b) and 8% to 30% (This et al. 1990). However, in Oxyrrhina addition lines, there was a decrease in ovule transmission frequency (Srinivasan et al. 1998). Addition lines Raphanus sativus–B. rapa, R. sativus–B. oleracea and R. sativus–Moricandia arvensis were generally stable, and predominant formation of gametes with added chromosomes might explain these observations (Kaneko et al. 1991; Bang et al. 2002; Kaneko et al. 2003). The male and female fertility of B. nigra additions were highly variable. Pollen fertility in general was high. B. nigra chromosome 2 in B. oleracea induced complete male sterility but not in B. napus (Chevre et al. 1997b). A similar behavior was also observed by This et al. (1990), where two B. nigra addition lines were totally male sterile

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in D. erucoides background. Monosomic R. sativus additions to alloplasmic (R. sativus) B. napus showed disturbed stamen development with very poor pollen production (Budhan et al. 2008). Some of the monosomic additions have been observed to restore fertility to alloplasmics, and four such examples are reported. Synteny group 6 of B. oxyrrhina to (B. oxyrrhina) B. rapa (16% pollen fertility, Srinivasan et al. 1998), an unspecified chromosome of Moricandia arvensis to (M. arvensis) B. juncea (53% pollen fertility, Prakash et al. 1998), chromosome c of Moricandia arvensis to (M. arvensis) R. sativus (85.6 pollen fertility, Bang et al. 2002), chromosome III of Arabidopsis thaliana to (A. thaliana) B. napus (Leino et al. 2004), and Raphanus chromosome f to (R. sativus) B. napus (Budhan et al. 2008). Due to small size of chromosomes and nonavailability of precise cytological landmarks in the earlier years, addition lines were characterized either through rare association with specific morphological characters, such as flower color, male sterility, or disease resistance; in recent years, isozyme and DNA markers are widely employed to characterize them. Markers employed include RFLP, RAPD, SSR, and GISH and FISH. By making use of these techniques, substantial information has been accumulated. Isozymes were initially used to characterize addition lines. B. rapa-oleracea was identified using such enzyme systems, such as 6PGD, PGI, LAP, and PGM (Quiros et al 1987); 6 PGD-1, PGM-1, and GOT-5 (McGrath and Quiros 1990); and PGM-2, PGDH-1, and PGDH-2 (Hu and Quiros 1991). B. nigra chromosome additions in the background of B. napus genome were characterized extensively using a large number of isozymes, such as MDH, IDH, LAP, 6-PGDH, ACO, PGI, TPI, GOT, PGM, and ADH (Chevre et al. 1991; Struss et al. 1996). Monosomic additions of Diplotaxis erucoides–B. nigra revealed synteny associations for loci coding for isozyme markers GOT-2, 6PGD-2, MDH-2, LAP-2, and TPI-1 (Quiros et al. 1987). This et al. (1990) located these synteny associations on four different B. nigra chromosomes using six isozyme loci and confirmed the observations of Quiros et al. (1987). Also a Diplotaxis erucoides chromosome was observed to carry three isozyme alleles (Chevre et al. 1994b). RAPD analysis have been carried out extensively in brassicas. Use of this system has been demonstrated in B. nigra (Quiros et al 1991; Struss et al. 1992, 1996; Chevre et al. 1994a), B. oxyrrhina (Srinivasan et al. 1998), Diplotaxis erucoides (Chevre et al. 1994b), and Moricandia arvensis (Bang et al. 2002) additions. Similarly, addition lines in several instances have been characterized through RFLPs. Using these markers, many chromosomes of the different genomes could precisely

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be identified. The markers revealed extensive intergenomic recombination, presence of duplicated loci, and synteny rearrangements of chromosomes. GISH has been one of the major tools to identify alien chromosomes and was employed in addition lines for Sinapis arvensis (Snowdon et al. 2000b), Arabidopsis thaliana (Leino et al. 2004), S. alba, Crambe abyssinica (Wang et al. 2005b, 2006a), and Orychophragmus violaceus chromosomes (Li and Ge 2007) in the background of the B. napus genome. Recently FISH has been used to identify the addion lines. For example, Peterka et al. (2004) identified chromosome d of Raphanus sativus carrying a gene imparting resistance to beet cyst nematode. Hasterok et al. (2005b) characterized three of the nine B. oleracea var. alboglabra chromosome additions using double target FISH. Chromosome addition lines as such are commercially unacceptable because of their unstable nature, reduced fertility, and expression of undesirable traits due to alien chromosomes. However, these lines are of academic interest and important genetic stocks for introgressing alien genetic material that might ultimately confer agronomic or horticultural advantages. For achieving gene introgression, homoeologous recombination between the alien chromosome and its homologous counterpart of the recipient genome should occur. Location of genes for horticultural traits such as curd in cauliflower, heading in cabbage, root enlargement in turnip, and axillary bud enlargement in Brussels sprouts are envisaged (Quiros et al. 1987; This et al. 1990). Chromosomal location of various genes controlling important agronomic traits have been accomplished by dissecting the respective genomes in various nuclear background; these include determining black-seeded phenotype and erucic acid content on a B. oleracea var. alboglabra chromosome (Chen et al. 1997b); a male sterility factor on a Diplotaxis muralis chromosome (Fan et al. 1985); self-incompatibility locus in B. nigra on chromosome 3 (Chevre et al. 1997b); resistance genes to black leg (Phoma lingam) on B. nigra (Zhu et al. 1993; Chevre et al. 1996; Struss et al. 1996); and Sinapis arvensis chromosomes (Snowdon et al. 2000b); B. oleracea chromosome possessing genes conferring resistance to turnip mosaic virus (TuMV) (Kaneko et al. 1996); a fertility-restoring gene on chromosome III of Arabidopsis (Leino et al. 2004); resistance to beet cyst nematode on chromosome d of Raphanus sativus (Peterka et al. 2004); and high erucic acid content on a Crambe abyssinica chromosome (Wang et al. 2006a). A dominant fertility restoration gene was introgressed to CMS (Moricandia) B. juncea following generation of Moricandia chromosome addition on CMS (Prakash et al. 1998), as also from

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Enarthrocarpus lyratus to CMS (Lyratus) B. rapa (Deol et al. 2003) and B. juncea (Banga et al. 2003a). RAPD markers linked with the genes for erucic acid and seed color on B. oleracea var. alboglabra chromosomes have been established (Jrgensen et al. 1996; Chen et al. 1997b). It was obsereved that chromosome 4 carries the gene for seed color and exerts its control embryonically. Chromosome 1 carries a gene that controls seed color maternally (Heneen and Brismar 2001). Hasterok et al. (2005b) are of the view that precise identification of extra chromosome in addition lines could be accomplished by using chromosome-specific or even arm-specific sets of BAC clone-based probes, as has been demonstrated by Howell et al. (2002), Ziolkowski and Sadowski (2002), and Koo et al. (2004).

VII. MITOCHONDRIAL GENOME A. Organization Plant mitochondrial genomes are relatively large as compared with other eukaryotes and range in size from 200 kb to 2000 kb (Fauron et al. 2004). Among the angiosperm plants examined so far, members of Brassica and related genera have the smallest mitochondrial genome. For example, the mt-genome of Sinapis alba (syn. B. hirta) is 208 kb while the values for B. rapa, B. oleracea, and B. napus are 218, 219, and 222 kb, respectively (reviewed in Palmer 1985; Pring and Lonsdale 1985). Arabidopsis thaliana, the model eudicot plant with the smallest nuclear genome, has a mitochondrial genome of 367 kb (Unseld et al. 1997). The circular nature of the mitochondrial genome of plants was first deduced from physical mapping studies of B. rapa and B. oleracea (Chetrit et al. 1984; Palmer and Shields 1984). This is now confirmed from complete mitochondrial genome sequencing exercises in several organisms including fungi, plants, and animals. However, such single circular molecules have not been physically visualized from electron microscopic examinations. Instead, small circular molecules of varying sizes have been observed in plant mitochondrial preparations. Based on RFLP analysis, Palmer and Shields (1984) demonstrated a tripartite structure for the mitochondrial genome of B. rapa. They identified two 2 kb direct repeats in the master circular genome, which were found to undergo recombination, giving rise to subgenomic circles of 83 and 135 kB. Similar organization of mt-genome has now been found in several other plants. The repeats ranging in size from 50–100 bp to 1–10 kb have been recorded in mitochondria of several species. For

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example, the B. napus mt genome has two large repeats of 1 to 10 kb and 37 of 0.1 to 1.0 kb. A. thaliana has four repeats of 1 to 10 kb and 90 repeats of 0.1 to 1.0 kb. These repeats are involved in homologous recombination. Intra- and intermolecular recombination in the repeat region is believed to generate multipartite subgenomic circular molecules. Such recombination events have been implicated in substoichometric shift in mitochondrial genome in different tissues and accessions, and creation of novel open reading frames (orfs). S. alba carries only a single copy of the repeat found in B. rapa and thus is the only species known to lack any large direct repeats (Palmer and Herbon 1987). The repeat sequences contain protein coding sequences; hence such genes are duplicated. The repeat sequences including the protein coding genes are different in different species. For example, in B. napus, a part of the cox2 gene is found in the repeat region whereas in A. thaliana atp6 gene is duplicated (Handa 2003). Detailed restriction profiles of mitochondrial genomes of Brassica species have revealed very limited intraspecific variation within species. Intraspecific variations in the form of two short deletions (100 and 700 bp in B. nigra) and one inversion (in S. alba) were detected (Palmer 1988). Considerable variation is found among species in both mt-DNA restriction and RFLP patterns (Palmer and Herbon 1987; Palmer 1988; Pradhan et al. 1992). However, most of the variation appears to be restricted to noncoding regions (Palmer and Herbon 1986, 1987). Based on comparative restriction analysis of different mt-genomes, it was found that inversions and small deletions are mainly responsible for the observed variation in mt-genomes among species. For example, mt-genome restriction profiles of S. alba and B. rapa differ significantly. However, most of the mt-genome can be divided into 11 regions; sequences within each region have the same arrangement in the two genomes, but the relative orientation and order of these regions differ between the species (Palmer and Herbon 1987). Similarly, B. rapa and B. oleracea differ by three large inversions whereas B. rapa and Raphanus differ by 14 inversions (Palmer 1988). B. Gene Content Despite large variation in size, gene content in mitochondrial genomes of angiosperms appears to be comparable and limited. Thus much of the observed difference in mt-genome size among different species is attributable to the insertion of foreign DNA and the presence of large repeated regions. This conclusion is further strengthened

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following the availability of complete mitochondrial genome sequences of A. thaliana and B. napus (Handa 2003). Plant mitochondrial genomes contain about 50 genes coding for various functions such as transcription, protein synthesis and transport, oxidative phosphorylation, and so on. In addition, dozens of orfs of unknown function are also found in sequenced mitochondrial genomes of plants. The overall G þ C content of the B. napus genome is 45.2%, which is comparable to other plant mitochondrial genomes. The gene content of mitochondrial genomes of B. napus and A. thaliana is summarized in Table 2.19. The only major difference in gene content between mitochondrial genomes of B. napus and A. thaliana is with respect to rps14 gene, which is a nuclear gene in A. thaliana (Figueroa et al. 1999). Although A. thaliana contains 22 tRNA species (five more than B. napus), both the species can specify only 15 amino acids. Thus a complete set of t-RNA genes is lacking in Brassica and Arabidopsis mitochondrial genomes. Some of the sequences (about 3.6%) present in the B. napus mt-genome appears to be of plastid origin, including some tRNA species. Mitochondrial genes of B. napus share many features, such as the presence of introns and RNA editing with mt-genes of other species. Despite wide evolutionary divergence between A. thaliana and B. napus, there is a high degree of conservation at the functional level. The size and number of introns are identical between the two species. Similarly, the RNA editing sites (441 in Arabidopsis versus 427 in B. napus) are highly conserved (Handa 2003).

Table 2.19. Number of genes in mitochondrial genomes of B. napus and A. thaliana. Genes Respiratory Complex I Complex II Complex III Complex IV Complex V Cytochrome biogenesis Transcription Translation Transport t-RNA r-RNA orfs ( > 100 amino acids) Total (excluding orfs)

B. napus

A. thaliana

9 — 1 3 5 4 1 9 1 17 3 45 54

9 — 1 3 5 4 1 7 1 22 3 85 59

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It is now clear that during the course of evolution, much of the mitochondrial genome has been transferred to the nucleus. The complete genome sequencing of A. thaliana and rice have shown this more clearly. A 620-kb segment of mt genome is found on chromosome 2 of A. thaliana (Stupar et al. 2001). Similarly, a 190-kb sequence of rice mitochondrial genome is present on chromosome 12 (Ueda 2005). Therefore, it is not unexpected that other large segments of mt DNA will be found in nuclear genomes of Brassica species. C. Mitochondrial Plasmids Small autonomously replicating linear plasmids are also found in some accessions of Brassica. Palmer et al. (1983b) observed a 11.3-kb plasmid in B. rapa whose copy number varied 100-fold among accessions containing the plasmid. Its nucleotide sequence was found to differ from other known sequences. Further, the presence of plasmid was associated with cytoplasmic male sterility. Since this plasmid was absent in the cytoplasm donor species (R. sativus), its transmission from the male side was suspected. Handa et al. (2002) also reported a 11.6-kb linear plasmid in B. napus, which was capable of transmission through both maternal and paternal route. This plasmid contains six orfs (two coding for phage-type DNA polymerase and one coding for phage-type RNA polymerase). All six orfs were found to be transcribed, and proteins of at least three orfs are found at high levels in flower buds of B. napus.

VIII. PLASTID GENOME To date, complete plastid genome sequences are available for 55 species including A. thaliana. These studies have shown that cp-genomes of angiosperm plant species are highly conserved for both gene order and sequence. In general, plant cp-genomes are circular and contain two large (22–25 kb) inverted repeats. These features are found to be shared by members of genus Brassica. For example, Link et al. (1981) observed uniform 158-kb circular cp-DNA molecules of Sinapis alba by electron microscopic examination and later confirmed it through restriction analysis. Further, two inverted repeats accounting for nearly 30% of the cp-genome were identified. Most of the studies on cp-genomes in Brassica are focused on variation for restriction enzyme sites (Erickson et al. 1983; Palmer et al. 1983a; Vedel and Mathieu 1983; Warwick and Black 1991, 1993, 1994, 1997; Warwick et al. 1992).

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Palmer et al. (1983a) compared restriction patterns of six U-triangle species along with S. alba and R. sativus. Small insertions or deletions (indels, 50–400 bp) seem to be the cause of most of the variations observed among species. Total sequence variation among Brassica species was estimated to be about 2.4%. Low level (0–0.01%) of intraspecific variation was also reported by Warwick’s lab based on cp-DNA RFLP and restriction analyses. A majority (53–80%) of restriction site mutations recorded were found between species. These studies have been extremely useful in identifying the maternal parents of the allotetraploid species. Availability of the complete cpDNA sequence of A. thaliana (Sato et al. 1999) may provide further opportunity for more incisive investigation of cp-genome evolution in Brassiceae.

IX. POTENTIAL ROLE OF ARABIDOPSIS THALIANA IN BRASSICA IMPROVEMENT A. A. thaliana as a Model Crucifer The fact that Arabidopsis and Brassica are in the same family is of great advantage to Brassica researchers who are benefiting from the information generated by the completed Arabidopsis thaliana genome sequence. Although the taxonomic distance between the two genera is large, with approximate divergence of 15 to 20 million years (Yang et al. 1999; Wroblewski et al. 2000), there is a great deal of conservation. The genomes of diploid brassicas are three to four times larger than that of Arabidopsis (157 Mb, Bennett et al. 2003), ranging from 468 Mb for B. nigra to 662 Mb for B. oleracea (Arumuganathan and Earle 1991). In spite of these differences, sequence conservation and synteny are large enough in most cases to use the genome of A. thaliana as a guide to find genes of interest in Brassica species. B. Cytology and Possible Origin of the A. thaliana Genome Similar to Brassica, mitotic chromosomes of A. thaliana are too small and lack distinctive landmarks; thus developing a physical cytogenetical map is not possible. Its chromosome number (2n ¼ 10) was determined by Laibach in 1907. Since 1960s, several researchers reported mitotic and meiotic and a giemsa C-banded karyotypes. However, a major breakthrough in its cytogenetics was achieved with the introduction of FISH technology using repeat DNAs such as

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ribosomal (18S, 26S, and 5S rDNAs), pericentromeric, centromeric, and telomere repeats (reviewed in Koornneef et al. 2003; Lysak et al. 2003). Both mitotic and meiotic chromosomes have been investigated, but better resolution was achieved with meiotic prophase complements, and heterochromatic and centromeric regions could be clearly differentiated. Using BAC contigs as probes in FISH, Fransz et al. (1998) presented a comprehensive pachytene bivalents karyotype. Accordingly, the mean total length of pachytene bivalents is 331 mm. The major part is euchromatin, with heterochromatin regions comprising of only 7.1 %, confined mostly in pericentromeric regions and NOR. Chromosomes 1 and 5 are the longest and metacentric with average length of 80.76 and 76.32 mm respectively. Chromosome 5, the second largest, carries a major and a minor 5S rDNA loci. The major locus is in the pericentromeric heterochromatin region of the upper arm and the minor locus is in the opposite arm. Chromosomes 2 and 4 are acrocentric and carry NOR. Their average length is 52.12 and 52.65 mm respectively. Chromosome 4 contains a 5S rDNA locus in the pericentromeric heterochromatin region of the short arm. Chromosome 3, a submetacentric with an average length of 69.34 mm, contains a major 5S rDNA in the middle of the long arm. Polymorphism for 5S rDNA loci was also observed in different ecotypes. However, all of them possess chromosomes 4 and 5 in the short arms. Earlier investigations documented 45S rDNA on NOR of chromosomes 2 and 4 and 5S rDNA on chromosomes 4 and 5 and polymorphic sites on chromosome 3 (Murata et al. 1997). Chromosome painting has been fairly successful due to the high resolution of FISH on pachytene chromosomes using BAC contig pools as probes. Chromosomes 1, 2, and 4 were entirely painted (Fransz et al. 2000; Lysak et al. 2003). Of these, chromosome 4 became the first entirely painted chromosome of a euploid plant karyotype. Earlier, identifying Arabidopsis individual chromosomes was difficult; now FISH technique has made it possible to unambiguously identify chromosomes or specific chromosome regions. Sequencing of the A. thaliana genome has disclosed extensive duplication, indicating a paleotetraploid origin possibly resulting from three rounds of genome duplications (Henry et al. 2006). The first two rounds took place very early on in the evolution of the angisoperms, between 90 and 135 million years ago. According to this tentative hypothesis, the most recent polyplodization event took place 24 to 40 million years ago, before the divergence of the Arabidopsis and Brassica lineages, when an ancestral species of n ¼ 4 originated a tetraploid n ¼ 8 species. Therefore, the basal chromosome number of the family is considered to

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be n ¼ 4. A. thaliana evolved from the hypothetical tetraploid species approximately 5 million years ago by reduction in chromosome number caused mostly by chromosome fusions and also by translocations and inversions (Henry et al. 2006; Schranz et al. 2006). These chromosomal rearrangements were accompanied by substantial DNA losses in A. thaliana (Town et al. 2006), when compared to A. lyrata and other related species (Schranz et al. 2006). C. Synteny Conservation The first attempt to compare synteny between Arabidopsis and Brassica based on RFLP markers was reported by Kowalski et al. (1994), who found ‘‘islands of conserved gene organization.’’ This report was confirmed by subsequent studies using techniques of increasing levels of sophistication, such as construction of maps using Arabidopsis ESTs (Lan et al. 2000; Babula et al. 2003) and a transcriptome map based on cDNA polymorphisms (Li et al. 2003). The EST maps revealed colinear regions of 20 cM between B. oleracea and A. thaliana (Babula et al. 2003). Transcriptome mapping allowed a global gene for gene alignment of the genomes of B. oleracea and A. thaliana by construction of a map based on mRNA from leaf tissue. The resulting map consisted of 247 cDNA markers obtained by the cDNA-SRAP technique. After sequencing 190 of the polymorphic cDNA fragments, 169 of these sequences had similarity to genes reported in Arabidopsis, which allowed alignment to the physical map of this species. Extensive colinearity was found between the two genomes for large chromosomal segments (but not whole chromosomes), albeit often inversions and deletions/insertions were shown. This work has been expanded by adding genomic markers and sequences corresponding to known genes to construct a high-density map of B. oleracea (Gao et al. 2007). Lukens et al. (2003) compared positions of Arabidopsis BAC sequences with B. oleracea sequences from RFLP probes previously located on a genetic map. Complex level of duplications prevented identification of orthologous segments between the two species. Hence, Lukens et al. (2003) developed an algorithm to facilitate identification of collinear loci. Using this statistical tool, the authors found 34 regions in Arabidopsis that were colinear with over 28% of the Brassica map, a result very similar to the result of empirical genetic mapping by Kowalski et al. (1994). Various approaches have discerned at least 20 to 30 chromosome structural rearrangements between Brassica and Arabidopsis, and a smaller number among the diploid Brassica spp. (Kowalski et al. 1994;

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Truco et al. 1996; Lan et al. 2000; Schranz et al. 2002; Lukens et al. 2003). Some of these rearrangements may have been in association with large-scale chromatin duplication. However, it remains controversial how many such events have occurred, how much chromatin they involved, and whether they occurred before or after the divergence of the various branches of the Brassica genus. Parkin et al. (2005) expanded the EST-RFLP comparative genomic study by mapping over 1,000 of these markers in B. napus. They found more than 21 conserved genomic units in Arabidopsis of approximately 9 Mb that could generate the B. napus genome through rearrangements and duplications. Centromeric regions are also conserved. These authors were able to construct alignments with each of the A and C genome chromosomes of B. napus and the five chromosomes of A. thaliana. As progress is made sequencing the Brassica genomes (Ayele et al. 2006), this new information enables researchers to compare genome organization of B. napus with A. thaliana, whose genome is now completely sequenced (Arabidopsis Genome Initiative 2000). This will add to the previous studies on comparative genomics for these species and reveal general conservation of gene content and colinearity. However, this conservation is incomplete, due to extensive chromosomal rearrangements (Kowalski et al. 1994; Cavell et al. 1998; Lagercrantz 1998; Lan et al. 2000; O’Neill and Bancroft 2000; Bancroft 2001; Quiros et al. 2001; Ryder et al. 2001; Babula et al. 2003; Li et al. 2003; Lukens et al. 2003). Town et al. (2006) sequenced BAC contigs covering a 2.2-MB region of the B. oleracea genome, which is partially triplicated in this species but duplicated in A. thaliana. They found 177 conserved colinear genes between the two species, and to explain absence of perfect triplication they inferred that 35% of the genes were lost in B. oleracea, most likely by deletions. Other factors breaking colinearity were gene duplication by tandem arrays in B. oleracea of single-copy A. thaliana genes, insertions, and dispersal of gene fragments. Additionally, the sequencing of three complete B. oleracea BAC clones corresponding to two different chromosomes in A. thaliana revealed less gene density in the former species and frequent accumulation of transposable elements (Gao et al. 2004, 2005, 2006). Genome enlargement by TE has occurred mostly by accumulation of tandem repeats and retrotransposons largely centered in centromeric and peri-centromeric heterochromatic regions (Lim et al. 2007). Therefore, during the evolution of these two Brassicaceae species, changes have occurred not only in chromosome number but also in chromosomal structure, causing frequent breaks in synteny (Gao et al. 2005, 2006).

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D. Synteny-Based Gene Discovery and Cloning Based on genomic shotgun sequences covering close to half of the B. oleracea genome, Ayele et al. (2006) estimated that 84% of the A. thaliana genes have a match in B. oleracea. They called these regions CAGs (conserved Arabidopsis genome sequences) and found that the highest sequence alignments occur near the centromeres of the Arabidopsis chromosomes. Sequence conservation is high in exons, ranging from 70% to 90% with the majority having similarities higher than 80%, whereas for introns it is < 70%. Protein similarity or orthologs is often above 95% (Gao et al. 2006). These high similarity values along with synteny conservation make it possible, in most cases, to find Brassica orthologs based on A. thaliana gene models with ease. Sadowski et al. (1996) exploited the genetic map of A. thaliana (Hauge et al. 1993) to probe the Brassica genomes with an A. thaliana gene complex carrying five genes within a 20-kb span (Gaubier et al. 1993). This complex comprises a well-characterized Em-like protein coding gene and other four flanking genes on chromosome 3. Although the fivegene complex array from A. thaliana was conserved on a single chromosome of each Brassica genome, additional copies for most of the genes were found in one or two other chromosomes. A similar situation was observed for a six-gene complex on A. thaliana chromosome 4, including the disease resistance gene RPS2 (Sadowski and Quiros 1998). In this case, besides the conserved array in one Brassica chromosome, four other chromosomes contained copies for some of the genes. The benefit of synteny conservation for gene discovery in Brassica is well demonstrated in studies on genes coding for glucosinolates (GSL), which are secondary metabolites synthesized by many species of the order Capparales, including Brassica and Arabidopsis. Breakdown products of GSLs, particularly isothiocynates, have been found to be anticarcinogenic (Talalay and Zhang 1996). Therefore, consumption of some of the brassica crops, such as broccoli, has been reported to exert cancer-protecting effects due to the formation of sulforaphane, an aliphatic glucosinolate-derived ITC (Fahey et al. 1997). The pattern of GSL accumulation and variability has been well described during the life cycle of A. thaliana (Brown et al. 2003). Furthermore, the genetic control GSL content is best understood in this model species, which provides a starting point to genetically improve GSL content and profiles in related Brassica crops. At least 10 genes related to the GSL pathway have been identified in A. thaliana

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(Mikkelsen et al. 2002; Wittstock and Halkier 2002), and knockout mutants for most of these genes are available. The colinearity between A. thaliana and B. oleracea has been explored for three chromosomal regions carrying three glucosinolate genes. Two of them are involved in side-chain elongation and belong to a gene family of major genes encoding methylthioalkylmalate synthase enzymes (MAM). In A. thaliana, three loci are duplicated in tandem (MAM1, MAM2 and MAM-L) on chromosome 5, and their presence depends on the ecotype; MAM-L is always present, but MAM1 and MAM2 are dispensable. A functional allele of MAM1 results in the presence of GSL with side chains containing four carbons (4C-GSL), whereas the presence a MAM2 in the absence of MAM1 results in the presence GSL with side chains containing three carbons (3C-GSL). The function of MAM1 is dominant to that of MAM2, because when both are present, the plants produce 4C-GSL (Kryomann et al. 2003). It was found that in B. oleracea, the BoGSL-ELONG gene corresponds to MAM1 in A. thaliana, which results in plants with 4CGSL (Li and Quiros 2002). Comparing the sequence of a 96.7-kb-long BAC clone (B19N3) from Brassica oleracea (broccoli) harboring the BoGSL-ELONG gene with its equivalent regions in A. thaliana disclosed these breaks in synteny:  B19N3 contains eight genes and six TEs.  The first two genes in this clone, Bo1 and Bo2, have its corresponding region at the end of chromosome 5 of Arabidopsis (24 Mb).  The third gene, Bo3, corresponds to an ortholog at the opposite end (2.6 Mb) of the same chromosome.  The other five genes, Bo4 to Bo8, also have a equivalent region on the same chromosome but at 7.7 Mb. Bo5 is a tandem duplicate of BoGSL-ELONG (Bo4) and was named BoGSL-ELONG-L, which is equivalent to MAM-L in A. thaliana. These five genes are colinear with those found in the corresponding region of Arabidopsis, which contains, however, 15 genes. Therefore, a cluster of 10 genes is missing in B. oleracea clone B19N3. All five genes have the same order and orientation in the genomes of both species. Their 36 exons constituting the eight homologous genes have high conservation in size and sequence identity in both species. The presence of duplicate member MAM2 has not been surveyed in B. oleracea, although it is known that it is not present in the broccoli variety used to study the segment harboring the BoGSL-ELONG gene.

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The region was further expanded by constructing a contig primer walking and BAC-end sequencing, revealing general gene colinearity beyond the segment harboring the BoGSL-ELONG gene (Gao et al. 2005). Two other B. oleracea BAC clones were surveyed for colinearity. The second BAC clone contained gene BoGSL-PRO, which is also a homolog of the MAM A. thaliana gene family. This gene has its homolog at the top of chromosome I in A. thaliana (At1g18500, MAM4). A duplicate member of this gene is located in the opposite arm of the same chromosome (At1g74040, MAM3). This gene is likely orthologous to BoGSLPRO-L, another member in the family also at a different location in B. oleracea. Twelve protein-coding genes and 10 TEs were found in this clone. The corresponding region in A. thaliana chromosome I has 14 genes and no TEs. No synteny breaks was observed for this clone, except for the two missing genes absent in B. oleracea (Gao et al. 2006). Compared to its homologous A. thaliana sequence, the third B. oleracea BAC clone harbors the side-chain modification gene BoGSL-ALK and has 101 kb. It is involved in desaturation, addition of a double bond, and removal of the methylsulfinyl residue in both 3C- and 4C-GSL. It corresponds to the gene AOP2 in A. thaliana, which is a member of another gene family. These genes code for 2-oxoglutarate-dependent dioxygenases, forming part of the AOP gene family located on chromosome 4. It is found that the sequenced B. oleracea BAC clone harbors 23 genes, including BoGSL-ALK, while the equivalent region in Arabidopsis contains 37 genes. All 23 common genes have the same order and orientation in both Brassica and Arabidopsis. The 16 missing genes in the broccoli BAC clone constitute two major blocks of five and seven contiguous genes, two singletons and a twosome. The 118 exons corresponding to these 23 genes have high conservation between the two species. The arrangement of the AOP gene family in A. thaliana is: AOP3 (GS–OHP)   AOP2 (GS– ALK) pseudogene AOP1. In contrast, in B. oleracea (broccoli and collard), two of the genes are duplicated and the third gene, AOP3, is missing, and genes are arranged as follows: Bo–AOP2.1 (BoGSL–ALKa)  pseudogene—AOP2.2 (BoGSL–ALKb)—AOP1.1—AOP1.2 (Gao et al. 2004). The general conclusion from these studies is very much the same that Kowaltski et al. (1994) reached earlier. There is general gene colinearity between A. thaliana and Brassica, but this is incomplete, and a break in synteny is common due to chromosomal rearrangements. Furthermore, these differences are accentuated by the presence of tandem duplicates for some genes in Brassica. Another difference that can be inferred but not generalized, because of the small portion of the

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genome surveyed so far, is the lower gene density found in the three BAC B. oleracea clones. This is due mostly to the insertion of TE in intergenic spacers and introns. As a consequence of these changes and breaks in colinearity, especially the frequent absence of genes in corresponding segments of A. thaliana, using this species as a guide to find a corresponding Brassica gene is not a trivial task. The tandem duplicates often found in the latter species require further experimentation to determine the correct gene based on its functionality and expression. E. Arabidopsis Knowledge-Based Gene Discovery and Brassica Improvement Brassica and Arabidopsis genomes share a high degree of homology ( > 80%), particularly in the exon regions, and most of the genes present in Brassica are represented in Arabidopsis. Hence knowledge gained from Arabidopsis is highly transferable to Brassica, and is providing valuable insights into various aspects of Brassica, including domestication and speciation, growth and development, and metabolism. Various approaches and resources currently are being employed to accomplish the goal of assigning functions to all the genes in Arabidopsis by 2010. Brassica improvement is expected to get a boost from the availability of complete functional genomic information of Arabidopsis. Once the key genes responsible for expression of a given trait are identified in Arabidopsis, they can be used to engineer the trait in Brassica. The examples discussed next highlight the significance of Arabidopsis functional genomics to Brassica. 1. Understanding Domestication. Kempin et al. (1995) identified a gene mutation in Arabidopsis, which together with a mutation in another gene, AP1 conferred cauliflower phenotype. Further, the same gene was found mutated in cultivated cauliflower suggesting its role in cauliflower phenotype. Subsequent studies have demonstrated that mutations in the cauliflower orthologue gene (BoCAL) are, indeed, responsible for cauliflower phenotype and have been selected during domestication of cauliflower (Lowman and Purugganan 1999; Purugganan et al. 2000). Similalry, nonshattering siliques is a characteristic feature of crop domestication. Liljegren et al. (2000) identified that mutations in two redundant genes SHP1 and SHP2 lead to non-shattering siliques in Arabidopsis. SHP1 and SHP2 orthologues are present in cultivated Brassica species and are expected to play similar function. Additional genes FRUITFUL and REPLUMLESS have also been identified in

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Arabidopsis, which negatively regulate SHATTERPROOF (Roeder et al. 2003). Based on this information, stergaard et al. (2006) developed nonshattering B. napus lines. Genes governing vernalization response and flowering time have also been well characterized in Arabidopsis. Robert et al. (1998) isolated four orthologues of Arabidopsis CONSTANS gene from B. napus lines differeing in flowering time and showed that their function is conserved between Arabidopsis and Brassica. FLC is a major gene responsible for suppression of flowering in Arabidopsis and is downregulated upon exposure to cold temperature. Kole et al. (2001) found that the major QTL, VFR2 responsible for winter type B. rapa cosegregated with FLC orthologues. These studies illustrate how Arabidopsis could serve as a reference for Brassica improvement. 2. Understanding Metabolism. Fatty acid metabolism has been extensively studied in Arabidopsis, and genes encoding key enzymes involved in fatty acid synthesis, elongation, and modification have been cloned and characterized. Analysis of QTLs for oil quality in Brassica crops have revealed that, in a majority of cases, these QTLs correspond to the known Arabidopsis genes involved in fatty acid metabolism. For example, FAE1 gene encodes the enzyme responsible for erucic acid biosynthesis in Arabidopsis. Mutations in the othrologues of the FAE1 gene have been found to be responsible for low-erucic acid in seed oils of B. rapa and B. oleracea (Das et al. 2002). Similarly, in B. juncea, FAE1.1 and FAE1.3 genes have been shown to cosegregate with QTLs, which account for 60% and 38% varaince for erucic acid content (Mahmood et al. 2003). Vitamin E (a-tocopherol) synthesis is restricted to photosynthetic organisms. Molecular analysis of Arabidopsis mutants has helped unravel the genes involved in tocopherol biosynthesis. Shintani and Della P