Sunflowers 2014

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BOTANICAL RESEARCH AND PRACTICES

SUNFLOWERS GROWTH AND DEVELOPMENT, ENVIRONMENTAL INFLUENCES AND PESTS/DISEASES

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BOTANICAL RESEARCH AND PRACTICES

SUNFLOWERS GROWTH AND DEVELOPMENT, ENVIRONMENTAL INFLUENCES AND PESTS/DISEASES

JUAN IGNACIO ARRIBAS EDITOR

New York

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Copyright © 2014 by Nova Science Publishers, Inc. All rights reserved. No part of this book may be reproduced, stored in a retrieval system or transmitted in any form or by any means: electronic, electrostatic, magnetic, tape, mechanical photocopying, recording or otherwise without the written permission of the Publisher. For permission to use material from this book please contact us: Telephone 631-231-7269; Fax 631-231-8175 Web Site: http://www.novapublishers.com NOTICE TO THE READER The Publisher has taken reasonable care in the preparation of this book, but makes no expressed or implied warranty of any kind and assumes no responsibility for any errors or omissions. No liability is assumed for incidental or consequential damages in connection with or arising out of information contained in this book. The Publisher shall not be liable for any special, consequential, or exemplary damages resulting, in whole or in part, from the readers‘ use of, or reliance upon, this material. Any parts of this book based on government reports are so indicated and copyright is claimed for those parts to the extent applicable to compilations of such works. Independent verification should be sought for any data, advice or recommendations contained in this book. In addition, no responsibility is assumed by the publisher for any injury and/or damage to persons or property arising from any methods, products, instructions, ideas or otherwise contained in this publication. This publication is designed to provide accurate and authoritative information with regard to the subject matter covered herein. It is sold with the clear understanding that the Publisher is not engaged in rendering legal or any other professional services. If legal or any other expert assistance is required, the services of a competent person should be sought. FROM A DECLARATION OF PARTICIPANTS JOINTLY ADOPTED BY A COMMITTEE OF THE AMERICAN BAR ASSOCIATION AND A COMMITTEE OF PUBLISHERS. Additional color graphics may be available in the e-book version of this book.

Library of Congress Cataloging-in-Publication Data Sunflowers : growth and development, environmental influences and pests/diseases / editor: Juan Ignacio Arribas (Electrical Engineering Department, Univ. Valladolid, Spain). pages cm Includes index. ISBN:  (eBook)

1. Sunflowers. I. Arribas, Juan Ignacio. QK495.C74S87 2014 583'.99--dc23 2014003599

Published by Nova Science Publishers, Inc. † New York

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CONTENTS Preface

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Chapter 1

An Introduction to the Sunflower Crop Fabián Fernández-Luqueño, Fernando López-Valdez, Mariana Miranda-Arámbula, Minerva Rosas-Morales, Nicolaza Pariona and Roberto Espinoza-Zapata

Chapter 2

Floral Biology of Sunflowers: A Histological and Physiological Analysis Basudha Sharma, Rashmi Shakya and Satish C. Bhatla

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Development of Female Reproductive Structures and Apomixis in Sunflowers Olga N. Voronova

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Chapter 3

Chapter 4

Genetics and Genomics Applied to Sunflower Breeding Carla Filippi, Jeremías Zubrzycki, Verónica Lía, Ruth A. Heinz, Norma B. Paniego and H. Esteban Hopp

Chapter 5

Sunflower Genetic Resources: Interspecific Hybridization and Cytogenetics in Prebreeding Jovanka Atlagić and Sreten Terzić

Chapter 6

Functional Genomics and Transgenesis Applied to Sunflower Breeding Sebastian Moschen, Laura M. Radonic, Guillermo F. Ehrenbolger, Paula Fernández, Verónica Lía, Norma B. Paniego, Marisa López Bilbao, Ruth A. Heinz and H. Esteban Hopp

Chapter 7

Disease Management in Sunflowers Regina M. V. B. C. Leite

Chapter 8

Recent Advances for Developing Resistance against Plasmopara halstedii in Sunflowers Osman Radwan

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vi Chapter 9

Contents Effects of Crop Management on the Incidence and Severity of Fungal Diseases in Sunflowers P. Debaeke, E. Mestries, M. Desanlis and C. Seassau

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Chapter 10

Insect Pests of Sunflowers in Africa Hannalene du Plessis

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Chapter 11

Soil Amendments and Their Effects on Sunflower Growth Fernando López-Valdez, Fabián Fernández-Luqueño, Perla Xóchitl Hernández-Rodríguez, Minerva Rosas-Morales and Silvia Luna-Suárez

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Chapter 12

Nutrition and Fertilization of Sunflowers in Brazilian Cerrado C. de Castro, F. A. Oliveira, A. Oliveira Junior and N. P. Ramos

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Chapter 13

Environmental Issues in the Sunflower Crop of Midwestern Brazil: Diversification and Complementarities in the Biodiesel Chain N. P. Ramos, A. M. M. Pires, C. C. A. Buschinelli, H. B. Vieira, C. de Castro and G. S. Rodrigues

Chapter 14

Micro and Macro-Morphological Variation of Cosmos bipinnatus and Cosmos bipinnatus var. Albiflorus in Sympatric Zones in Central Mexico M. Paniagua-Ibañez, A. Zepeda-Rodríguez, P. Mussali-Galante, R. Ramírez-Rodríguez and E. Tovar-Sánchez

Index

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To Juan Ignacio, Jr., Elena, Jr. and Elena

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PREFACE We are all well aware that the importance of the sunflower (Helianthus Annuus) as a crop has increased significantly in recent years, not only in the food industry but also as a natural energy resource in oil production. I am, thus, very pleased to be able to present this comprehensive monograph on a wide range of important issues regarding sunflowers, with an emphasis on environmental influences, pests and diseases in order to maximise production whilst minimising costs. Contributors where selected based on their proven experience in the field of sunflowers. Contributors submitted an extended abstract that was assessed for relevance. They were then invited to contribute draft chapters. Each chapter underwent a stringent and thorough peer review process by other experts in the field, with final approval by the editor who, thus, was able to balance the topics from all contributors. The book contains important original results. Each chapter deals with a different topic, and draws, where appropriate, from studies and results previously published by the authors. Authors were encouraged to complement their writing with original and high quality graphs, charts, tables, figures, pictures and photographs. It‘s my honour and pleasure to acknowledge the rigorous work carried out by all authors in this book, and at the same time I am very grateful to them for trusting me in leading this project in the role of the editor of their work. My thanks also go to the anonymous reviewers who contributed their time so generously to this book, and without whom it would not exist. I am also very grateful to Nova Science Publishers for inviting me to lead this book, and thank them for the help and coverage provided during the whole time that this project lasted. I really do hope that you find this book of interest and wish you enjoy its reading as much as I have done through the whole editing process and as much I am sure all authors have done while writing it. The book is structured as follows: Chapter 1 introduces sunflowers. Chapters 2 and 3 detail the biology of a sunflower. Chapters 4, 5 and 6 explore the important topics of sunflower production: genetics, genomics and the breeding of sunflowers. Chapters 7, 8, 9 and 10 address with other important aspects of sunflower production, pests and diseases. Chapters 11 and 12 deal with sunflower nutrition and growth. Finally, Chapter 13 presents environmental sunflower crop issues and Chapter 14 studies sunflower morphological Cosmos variations.

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In Chapter 1, entitled An Introduction to the Sunflower Crop, Dr. Luqueno and colleagues from the Natural Resources and Energy Group, Cinvestav-Saltillo, Mexico, present an enjoyable historical perspective on sunflowers. Sunflower (Helianthus annuus L.) belongs to the family Asteraceae. The sunflower plant originated in eastern North America. It is thought to have been domesticated around 3000 B.C. by Native Americans. In the late 1800s the sunflower was introduced in the Russian Federation where it became a food crop and Russian farmers made significant improvements in the way that the sunflower was cultivated. Since 3000 B.C. a wide range of uses of sunflower have been reported throughout the world. Sunflower is well known by its phytoremediation potential and by its seed oil content. Because the sunflower has several potential markets, it is a good choice for growers on both small and large scales. However, it has to be remembered that scientific, technical or agricultural projects linked with sunflower have to include side effects elsewhere in order to shape a sustainable future. In Chapter 2, entitled Floral Biology of Sunflower - A Histological and Physiological Analysis, Dr. Bhatla and colleagues from the Laboratory of Plant Physiology and Biochemistry, Department of Botany, University of Delhi, India, introduce a meticulous approach to the development of sunflower inflorescence as considered under three phases listed next: inflorescence initiation, floret development and anther formation. Anthesis of disc florets is a phytochrome-mediated response and is also modulated by phytohormones, such as auxins and gibberellic acid. Dr. Bhatla and colleagues focus on the role of various biomolecules, like glycoproteins, calcium, nitric oxide, reactive oxygen species, and associated scavenging enzymes in relation to stigma maturation. Specific expression of lignoceric acid (24:0) in the pollen coat and localization of lipase in pollen and stigma are likely to have possible roles during pollen-stigma interaction. The phenomenon of selfincompatibility and pseudo self-compatibility in sunflower has been discussed. The initial processes accompanying pollen-stigma interaction and their regulation, especially the adhesion of pollen on the stigma surface, hydration, formation of an "attachment foot" and pollen tube germination in sunflower with respect to self-and cross-pollinated situations, has also been dealt with in detail. In Chapter 3, entitled Development of Female Reproductive Structures and Apomixis in Sunflowers, Dr. Voronona from the Department of Embryology and reproductive biology, Komarov Botanical Institute of RAS, Saint-Petersburg, Russia, presents an scrupulous visual analysis of archesporial cells which are formed and gave rise to megaspore mother cells. The meiotic divisions produced a linear tetrad of haploid megaspores and from one chalazal megaspore a Polygonum-type embryo sac is formed. Under natural conditions the apomixis phenomenon was hardly observed in genus Helianthus L. In addition, author shows how on plants of CMS-lines a number of anomalies in development of female reproductive system were detected, including such phenomena as total absence of embryo sac, apospory and integumentary embryony. Lack of the main embryo sac and formation of additional aposporous embryo sacs could be observed in the same ovule. Finally, investigation of the early stages of ovule development showed that aposporous embryo sacs originated from the same ovule subepidermal cells as a normal embryo sac. In Chapter 4, entitled Genetics and Genomics Applied to Sunflower Breeding, Dr. Hopp and colleagues from the Instituto de Biotecnologia, Centro de Investigaciones Veterinarias y Agronomicas, Instituto Nacional de Tecnologia Agropecuaria, Hurlingham, Argentina, present a thoughtful study about new breeding strategies based on molecular markers, like

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quantitative trait loci mapping, association mapping and genomic selection that are currently being developed for commercial crop improvement. The need to increase efficiency and precision, and save time, resources and efforts, has motivated the application of Marker Assisted Selection (MAS) in sunflower breeding programs. Furthermore, nowadays, the focus is on tolerance improvement to biotic and abiotic stresses and oil quality and yield increasing, in order to reduce the gap between potential and actual sunflower production. In Chapter 5, entitled Sunflower Genetic Resources – Interspecific Hybridization and Cytogenetics in Prebreeding, Drs. Atlagic and Terzic present a rigorous description of the genus Helianthus by reviewing genetic resources, cytogenetic research and application in breeding. Besides the review of available literature, research results of the Institute of Field and Vegetable Crops are presented in detail since the establishment of its collection in 1980. Experience collected during this period indicates the difficulties in collection maintenance, interspecific crosses and isolation of desired genes. Nevertheless, genus Helianthus proved to be a good source of material for the constant improvement of cultivated sunflower. In Chapter 6, entitled Functional Genomics and Transgenesis Applied to Sunflower Breeding, Dr. Hopp and colleagues from the Instituto de Biotecnologia, Centro de Investigaciones Veterinarias y Agronómicas, Instituto Nacional de Tecnologia Agropecuaria, Hurlingham, Argentina, introduce an interesting chapter where they analyze different strategies which have been developed in the last decade from functional genomics and post genomics disciplines to contribute to the elucidation of gene regulation and identification of key metabolic pathways involved in the response to biotic and abiotic stresses in sunflower. The state of the art of strategies for gene function, studies in silico and in planta, by stable gene transfer or agroinfiltration in sunflower as well as in the model system for Asteraceae species, lettuce, are discussed within the frame of their application in sunflower breeding. In Chapter 7, entitled Disease Management in Sunflowers, Dr. Leite from Embrapa Soybean, Brazil, presents an interesting approach regarding the most important sunflower diseases and strategies for disease management. Sunflower can be affected by the presence of diseases, which may, depending on climatic conditions that favor the occurrence of pathogens and the infective process, lead to a significant reduction on yield and quality of product. Disease management should be based on an integrated program, in order to give support for the sustainability and competitiveness of the sunflower crop. In Chapter 8, entitled Recent Advances for Developing Resistance against Plasmopara Halstedii in Sunflowers, Dr. Radwan from the Department of Natural Resources and Environmental Sciences, University of Illinois at Urbana-Champaign, Urbana, IL, USA, presents an interesting visual approach to Downy Mildew disease, as one of the most important diseases of sunflower, which leads to an economic yield loss. In last two decades, different approaches of genetics and genomics have significantly contributed to better understand sunflower-Plasmopara halstedii. This progress directed to development of sunflower lines carrying resistance to different races of this pathogen. In Chapter 9, entitled Effects of Crop Management on the Incidence and Severity of Fungal Diseases in Sunflowers, Dr. Debaeke and colleagues from the Institut National de la Recherche Agronomique (INRA), Toulouse, France, dissected the effects of crop management on the incidence and severity of major fungal diseases in sunflower, including downy mildew, phoma, phomopsis and sclerotinia. They deeply reviewed and discussed the influence of sowing date, plant population, N fertilization, and irrigation on sunflower diseases from numerous experiments conducted in France during the last twenty years. They

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proposed indicators of canopy development and nutritional status that could be useful when developing crop management systems with reduced chemical applications. In Chapter 10, entitled Insect Pests of Sunflowers in Africa, Dr. du Plessis from the Unit for Environmental Sciences and Management, North-West University, Potchefstroom, South Africa, present a complete study regarding a number of insect species which have adapted to cultivated sunflower as a source of food and have consequently become economically important pests. However, although many insect species is associated with sunflower in Africa, only few are considered to be of potential economic importance. Insects most commonly reported as injurious to this crop, occur sporadically, but usually in high numbers. The families Noctuidae, Tenebrionidae, Curculionidae, Pentatomidae and Orsillidae are the most important. Various types of damage are caused to sunflower seedlings, but the damage symptoms are specific to a particular pest species. Dr. du PLessis argues that these seedling pests mainly constitute dusty surface beetles (Gonocephalum simplex), greater false wire worms (Somaticus spp.), cutworms (Agrotis spp.) and ground weevils (Protostrophus spp.). Total defoliation can be incurred by the Plusia looper, Trichoplusia orichalcea. Hemipterans and the African bollworm, Helicoverpa armigera are the most important insect pests of sunflower during the heading stages of crop development. Intensive feeding by hemipterans during this development stage results in deformed heads, which delay flower opening. The occurrence of the false chinch bug, Nysius natalensis on sunflower during the heading stage onwards in South Africa, is similar to that of N. stali in Nigeria. Since high summer temperatures prevail throughout the sunflower production area of South Africa and the most favourable temperature range for N. natalensis development is between 26°C and 38 °C, the potential for rapid population build-up by this pest during the sunflower production season is good. It is likely that N. natalensis can become important in sunflower production in other African countries with similar weather conditions too. The insect is polyphagous and a variety of wild host plants, mainly weed species as well as crops such as grain sorghum play an important role in sustaining its populations. Sunflower is not the preferred host but N. natalensis lays its eggs on sunflower when its preferred host plants are removed or dead. This behaviour explains the insects‘ injuriousness to late-planted sunflower because weed species have already senesced before the sunflower. This period often coincides with seed fill of lateplanted sunflower, providing an alternative for the insect for moisture, as well as seeds that are necessary for reproduction of the pest. Weeding in and around sunflower during seed fill of the crop, therefore results in destruction of the preferred host plants of N. natalensis, and they consequently move to sunflower where they feed and cause damage. When considering application of insecticides for control of this pest, it should take into consideration that N. natalensis is highly mobile and continuous re-infestations could occur. Timing of insecticide application is therefore important. African bollworm (H. armigera) is frequently present during the reproductive stage of cultivated sunflower in Africa. The attractiveness of sunflower to this pest is demonstrated by the trap‘s use as a trap crop in and around organic cotton fields in Tanzania. Larvae occur from the budding stage onwards. Levels of infestation vary between localities and seasons, sporadically reaching epidemic proportions. Sunflower has, however, the ability to compensate for head damage and along with the fact that preferential feeding sites of H. armigera are not the achenes, a significant number of larvae could be tolerated without any significant effect on yield. Actual damage is, therefore, the only criterion that could be used in the determination of economic injury levels for control of African bollworm on sunflower crop.

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In Chapter 11, entitled Soil Amendments and Their Effects on Sunflower Growth, Dr. Lopez and colleagues from the Centro de Investigacion en Biotecnologia Aplicada, Instituto Politecnico Nacional, Tlaxcala, Mexico, present a novel approach to several forms of amend or fertilize sunflowers as alternative to improves this important cultivar. In particular authors are interested in the study of organic materials that could be applied to soil in order to improve their properties and plant growth, keeping in mind that we must recycle or reuse this kind of materials. Finally, the organic amendments could be a beneficial disposal approach that must be considered. In Chapter 12, entitled Nutrition and Fertilization of Sunflowers in Brazilian Cerrado, Dr. Castro and colleagues present an interesting chapter centered in a particular region in Brazil, which authors argue that is recognized as a major global food producer and despite the fact that its agriculture occupies only less than 5 % of the national territory, the estimated grain production for 2013/2014 growing season is 195 million tons. The Brazilian Cerrado is the main agriculture expansion region in the country, driven by soybean cultivation in an area of 13 million ha. In this tropical agricultural region, sunflower has great potential for expansion and consolidation as an important component of sustainable crop rotation production systems. This chapter addresses the major limiting soil fertility factors which hinder the crop development and discusses fertilization management practices related to the main limiting nutrients, like the macronutrients nitrogen, phosphorus and potassium and micronutrients such as boron and molybdenum. Adequate management of soil acidity and fertilization has been proved as a powerful tool to improve the natural conditions of acidic and chemically poor arable tropical soils. In addition to the soil fertility assessment, leaf analysis is essential for the proper interpretation of plant nutritional status, thereby enabling better refinement of the crop nutritional management. In Chapter 13, entitled Environmental Issues in the Sunflower Crop of Midwestern Brazil – Diversification And Complementarities in the Biodiesel Chain, Dr. Ramos and colleagues from Embrapa Environment, Brazil, present an interesting study regarding the increase in global demand for renewable energy, the production of oilseeds, including sunflower, as feedstock for biodiesel. The increase in global demand for renewable energy has encouraged, both directly and indirectly, the production of oilseeds, including sunflower, as feedstock for biodiesel. In this scenario, authors argue that Brazil stands out for its excellent agronomic and climatic conditions for growing these crops throughout its territory. Sunflower is considered an interesting option and your production in the mid-western region of Brazil has done valuable contributions as a second, and especially the production practices adopted by the reference farmers of the country, rendering complementarities and diversification to both the food and the bioenergy sectors. In Chapter 14, entitled Micro and Macro-Morphological Variation of Cosmos Bipinnatus and Cosmos Bipinnatus Var. Albiflorus in Sympatric Zones in Central Mexico, Dr. Paniagua and colleagues from the Centro de Investigacion en Biodiversidad y Conservacion, Univ. Autonoma Estado de Morelos, Morelos, Mexico, present a concise but at the same time precise analysis of the morphological variations in various sunflower species in central Mexico area. Mexico is considered one of the centers of diversification of the Asteraceae family, which contains the greatest richness of flowering plants. Particularly, the TransMexican Volcanic Belt (TMVB) is a heterogeneous mountain belt, located in the central part of the country in an east–west direction, which has been considered a diversification site for many genera due to the vast number of species that it contains, as well as its high degree of

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endemism. Cosmos bipinnatus is present in various types of vegetation along the TMVB which is favored by disturbances. Taxonomic studies have documented that this sunflower species presents white to lilac ligules. However, horticulturists have considered the white variety as C. bipinnatus var. albiflorus. Still, there is no scientific evidence to support their observations. Therefore, the goal of this study was to compare the micro and macro morphology characters between C. bipinnatus individuals with white and lilac ligules to determine a possible morphological differentiation between both phenotypes in sympatric zones in the central Mexico region. Principal Component Analysis and Non-Metric Multidimensional Scaling showed a clear morphological differentiation between both groups; this pattern was consistent even when the ligule‘s color was not considered for the statistical analyses. Dr. Paniagua and colleagues results sign a possible speciation between these phenotypes and support a taxonomic shift for the Mexican sunflower with white ligules to C. bipinnatus var. albiflorus.

Juan Ignacio Arribas, PhD Associate Professor of Electrical Engineering University Valladoild, Spain Valladolid, November 2013 Tel: +34 983423000 Fax: +34 983423667 [email protected]

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In: Sunflowers Editor: Juan Ignacio Arribas

ISBN: 978-1-63117-347-9 © 2014 Nova Science Publishers, Inc.

Chapter 1

AN INTRODUCTION TO THE SUNFLOWER CROP Fabián Fernández-Luqueño1,, Fernando López-Valdez2, Mariana Miranda-Arámbula2, Minerva Rosas-Morales2, Nicolaza Pariona1 and Roberto Espinoza-Zapata3 1

Natural Resources and Energy Group, Cinvestav-Saltillo, Coahuila, México CIBA - Instituto Politécnico Nacional, Tepetitla de Lardizábal, Tlaxcala, México 3 Crop Breeding Department, UAAAN, Saltillo, Coahuila, México

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ABSTRACT Sunflower (Helianthus annuus L.) belongs to the family Asteraceae. The Helianthus genus contains 65 different species of which 14 are annual plants. The sunflower plant originated in eastern North America. It is thought to have been domesticated around 3000 B.C. by Native Americans. In the late 1800s the sunflower was introduced in the Russian Federation where it became a food crop and Russian farmers made significant improvements in the way that the sunflower was cultivated. Since 3000 B.C. a wide range of uses of sunflower have been reported throughout the world such as ornamental plant, medicinal, alimentary, feedstock, fodder, dyes for textile industry, body painting, decorations, and so on. Sunflower species are allelopathic in nature and this crop appears to have a bright future, especially if the scientists can translate the cutting-edge research into technologies that will reduce the reliance on synthetic herbicides, pesticides, and crop protection chemicals. On the one hand sunflower is well known by its phytoremediation potential, thus it can be speculated that the good tolerance of sunflower towards pollutants coupled with an increased accumulation/degradation capacity might contribute to an efficient removal of pollutants from soil and water; on the other hand sunflower possesses the potential to develop bioenergy systems that allow for synergies between food and energy production. Because the sunflower has several potential markets, it is a good choice for growers on both small and large scales. However, it has to be remembered that scientific, technical or agricultural projects linked with sunflower have to include side effects elsewhere in order to shape a sustainable future. *

Corresponding author: F. Fernández-Luqueño, Natural Resources and Energy Group, Cinvestav-Saltillo, Coahuila. C. P. 25900, México Tel.: +52 844 4389625; Fax: +52 844 4389610. E-mail address: cinves.cp.cha.luqueno@ gmail.com.

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F. Fernández-Luqueño, F. López-Valdez, M. Miranda-Arámbula et al.

Keywords: Allelopathy, biodiesel, phytoremediation, renewable energy, sustainable development, symbiosis

1. INTRODUCTION Sunflower (Helianthus annuus L.) belongs to the family Asteraceae. Helianthus genus contains 65 different species (Andrew et al., 2013). The name Helianthus, being derived from helios (the sun) and anthos (a flower), has the same meaning as the English name Sunflower, which has been given these flowers from a supposition that they follow the sun by day, always turning towards its direct rays. The sunflower that most people refer to is H. annuus, an annual sunflower. In general, it is an annual plant which possesses a large inflorescence (flowering head), and its name is derived from the flower's shape and image, which is often used to depict the sun. The plant has a rough, hairy stem, broad, coarsely toothed, rough leaves and circular heads of flowers (Khaleghizadeh, 2011). The heads consist of many individual flowers which mature into seeds on a receptacle base (Seghatoleslami et al., 2012). Sunflower is the world‘s fourth largest oil-seed crop and its seeds are used as food and its dried stalk as fuel. It is already been used as ornamental plant and was used in ancient ceremonies (Harter et al., 2004; Muller et al., 2011). Additionally, medical uses for pulmonary afflictions have been reported. In addition, parts of this plant are used in making dyes for the textile industry, body painting, and other decorations. Sunflower oil is used in salad dressings, for cooking and in the manufacturing of margarine and shortening (Kunduraci et al., 2010). Sunflower is used in industry for making paints and cosmetics. A coffee type could be made with the roasted seeds. In some countries the seed cake that is left after the oil extraction is used as livestock feed. In the Soviet Union the hulls are used for manufacturing ethyl alcohol, in lining for plywood and growing yeast. The dried stems have also been used for fuel. The stems contain phosphorous and potassium which can be composted and returned to soil as fertilizer. Sunflower meal is a potential source of protein for human consumption due to its high nutritional value and lack of anti-nutritional factors (Fozia et al., 2008). Sunflower was a common crop among American Indian tribes throughout North America. Evidence suggests that the plant was cultivated by natives in present-day Arizona and New Mexico about 3000 B.C. Some archaeologists suggest that sunflower may have been domesticated before corn (NSA, 2013). Although the scientific consensus had long been that sunflower was domesticated once in eastern North America, the discovery of pre-Columbian sunflower remains at archaeological sites in Mexico led to the proposal of a second domestication center in southern Mexico. However, evidences from multiple evolutionary important loci and from neutral markets support a single domestication event for extant cultivated sunflower in eastern North America (Blackman et al., 2011). The objective of this chapter is to present and discuss a summary about the huge amount of information in which the sunflower is the main subject. The chapter aims to assist people involved in all aspects of sunflower management, including conservation, agriculture, mining, energy, food production, health and other industries, to obtain a broad knowledge of sunflower and of its ecosystem services.

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An Introduction to the Sunflower Crop

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2. BOTANICAL AND MORPHOLOGICAL DESCRIPTION Sunflowers are botanically classified as Helianthus annuus L. (Table 1). They are large plant and are grown throughout the world because of their relatively short growing season. Sunflower is an annual herb, with a rough, hairy stem, 3 to 12 feet high, broad, coarsely toothed, rough leaves, 3 to 12 inches long and circular heads of flowers, 3 to 6 inches wide in wild specimens and often a foot or more in cultivation. The flower-heads are composed of many small tubular flowers arranged compactly on a flattish disk: those in the outer row have long strap-shaped corollas, forming the rays of the composite flower. Each sunflower head, or inflorescence, is actually composed of two types of flowers. What appears to be yellow petals around the edge of the head are actually individual ray flowers. The face of the head is comprised of hundreds of disk flowers, which each form into a seed (achene). The basic chromosome number for the Helianthus genus is 17. Diploid, tetraploid and hexaploid species are known. There are only 14 annual species of Helianthus. Plant breeders have made interspecific crosses within the genus and have transferred such useful characters as higher oil percentage, cytoplasmic male sterility for use in production of hybrids, and disease and insect resistance to commercial sunflower. Table 1. Scientific classification of H. annuus L.; this genus counts 65 different species Taxa Kingdom Plantae Subkingdom Viridaeplantae Infrakingdom Streptophyta Division Tracheophyta Subdivision Spermatophytina Infradivision Angiospermae Class Magnoliopsida Superorder Asteranae Order Asterales Family Asteraceae Subfamily Helianthoideae Tribe Heliantheae Genus Helianthus Specie annuus The taxonomic classification has been in place since 1753.

3. PRODUCTION In recent years, the sunflower cultivated area has been steadily increasing due to the breeding of dwarf high yielding hybrids that also facilitate mechanization and the emphasis given to polyunsaturated acids for human consumption. Global production grew steadily in last 25 years (PSD-USDA, 2011), and FAO expect a total world output close to 60 million tons towards 2050. The four largest producers (Russia, Ukraine, European Union and Argentina) account for 70% of global volume, with an exponential growth of production in

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the last ten years in the Black Sea region, with increased acreage an higher yields achieved by the replacing old varieties by hybrid seeds. According to data from FAOSTAT (FAOSTAT, 2011) Russia Federation ranked first producing ca. 9.7 millions of tons of sunflower seeds or 26% of the world total. Ukraine and Argentina ranked second and third place with 8.6 and 3.6 tons of sunflower seeds, respectively. France, Romania, China, Bulgaria, Hungary, Turkey, and Spain produced between 1.0 and 1.9 millions of tons of sunflower seeds (Table 2). The United States produced ca. 1.0 millions of tons of sunflower seeds, or 5% of the world‘s total production. That is enough to make the United States rank eleventh in that category. South Africa ranked twelfth producing ca. 0.9 millions of tons of sunflower seeds. Table 2. The highest twelve sunflower seed producing countries in the world during 2011 Place Countries Production (tons) 1 Russia Federation 9,696,450 2 Ukraine 8,670,500 3 Argentina 3,671,750 4 France 1,882,450 5 Romania 1,789,330 6 China 1,700,000 7 Bulgaria 1,439,700 8 Hungary 1,374,780 9 Turkey 1,335,000 10 Spain 1,084,300 11 United States of America 924,550 12 South Africa 860,000 Russia followed by Ukraine are harvesting almost half of the world sunflower seed production. The total sunflower seed production is reaching ca. 35 millions of tons Data source: data obtained from FAOSTAT (2011).

According to FAO (FAO, 2010), there are some key production parameters which have to be known by farmers throughout the world: 

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Sunflowers are grown in warm to moderate semi-arid climatic regions of the world from Argentina to Canada and from central Africa to the Commonwealth of Independent States (Esmaeli et al., 2012; Onemli, 2012). Frost will damage sunflowers at all stages of growth. The plant grows well within a temperature range of 20-25°C; temperatures above 25°C reduce yields and oil content of the seeds (Thomaz et al., 2012). Plants are drought-resistant, but yield and oil content are reduced if they are exposed to drought stress during the main growing and flowering periods. Sunflowers will produce moderate yields with as little as 300 mm of rain per year, while 500-750 mm are required for better yields (Gholamhoseini et al., 2013; Ghaffari et al., 2012). Sunflowers adapt to a wide variety of soil, but perform best on good soils suitable for maize or wheat production (Radanielson et al., 2012).

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Sunflower plant density of 5-8 plants per m2 is required to form the optimum leaf area for plant photosynthesis. Kernel weight (40-80 g per 1000 kernels) and the average number of kernels in a sunflower head (1200-1500) are the other most important yield component (Seassau et al., 2012; Emami-Bistghani et al., 2012). Sunflower growth depends more on nitrogen than any other nutrient. Due to its deep rooting system, sunflower is able to use nitrogen from soil layers that are inaccessible to wheat, corn or other field crops. The plant requires a maximum of 150 kg of nitrogen per hectare to produce a three tons ha-1 yield. Over fertilization may lead to sunflower lodging. Phosphorous, potassium, boron, magnesium and molybdenum are also needed to achieve the best yields (Jabeen and Ahmad, 2012; Babaeian et al., 2011). The average fatty acid composition of oil from temperate sunflower crops is 55-75% linoleic acid and 15-25% oleic acid. Protein content is 15-20% (Aznar-Moreno et al., 2013; Ali and Ullah, 2012). Planting in the Western Balkan countries, Eastern Europe and countries of the Former Soviet Union takes place during March and April (Zheljazkov et al., 2012; Saleem et al., 2008). Sunflower has one of the shortest growing seasons of the major economically important crops of the world. Early maturing varieties are ready for harvesting 90 to 120 days after planting, and late maturing varieties 120 to 160 days after planting. Delayed harvesting causes unwelcome changes in oil quality, with an increase in free fatty acid content. The seeds are ready to harvest when the heads turn black or brown and the seed moisture content reaches 10-12%. Grain combines are fairly easily adapted for the harvesting of sunflower by the addition of a head snatcher (Borbely et al., 2008). Depending on climatic and cultivation conditions, yields can vary from as much as 600 to 3000 kg ha-1; irrigation is a key factor for obtaining high yields (Chigeza et al., 2013; Khan et al., 2013; Akhtar et al., 2012).

Table 3 shows the oil yields in gallons per acre of oil producing crops, the yields will vary in different agroclimatic zones. Sunflower produces 98 Gal oil acre-1. That is enough to make the sunflower rank twenty-third in that category. Additionally, higher-yielding oil crops like safflower, mustards and sunflower have significant rotational benefits. For example, deep safflower and sunflower roots help break up hardpan and improve soil tilth.

4. GROWTH AND DEVELOPMENT Sunflower is a broadleaf plant that emerges from the soil with two large cotyledons (Rawat et al., 2010). The emergence will take four to five days when planted an inch deep in warm soil, but will take a few days longer in cooler soils or when planted deeper. Soil crusting can make it difficult for the large seedlings to push out of the soil. Sunflowers grow rapidly, producing large and rough leaves. Current sunflower varieties reach an average height of six feet, varying between five and seven feet depending on planting date and soil conditions (Saensee et al., 2012). After reaching their full height and blooming, heads on

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commercial cultivars turn downwards, designed to make it harder for birds to eat the seed. Commercial sunflowers have flowers that are self-compatible for pollination, meaning they do not require a pollinating insect, although some studies have shown bee pollinators providing a slight yield boost (de Carvalho and de Toledo, 2008). Some farmers prefer sowing their rows from north to south so that the capitula can lean into the row space, rather than bumping against an adjacent plant, causing some seed to fall (Olowe and Adeyemo, 2009). Table 3. Oil producing crops Number Crop Scientific name Yield (Gal oil acre-1) 1 Oil palm Elaeis guineensis Jacq. 610 2 Macauba palm Acrocomia aculeata Jacq. 461 3 Pequi Caryocar brasiliense Camb. 383 4 Buriti palm Mauritia flexuosa L. 335 5 Oiticia Licania rigida Benth 307 6 Coconut Cocos nucifera L. 276 7 Avocado Persea americana Mill. 270 8 Brazil nut Bertholletia excelsa Humb & Bonpl. 245 9 Macadamia nut Macadamia ternifolia F.V. Muell. 230 10 Jatrofa Jatropha curcas L. 194 11 Babassu palm Orbignya martiana Mart. 188 12 Jojoba Simmondsia chinensis Link 186 13 Pecan Carya illinoensis Wangenh. 183 14 Bacuri Platonia insignis Mart. 146 15 Castor bean Ricinus communis L. 145 16 Ghoper plant Euphorbia lathyris L. 137 17 Pissava Attalea funifera Mart. 136 18 Olive tree Olea europea L. 124 19 Rapessed Brassica napus L. 122 20 Opium poppy Papaver somniferum L. 119 21 Peanut Arachis hypogea L. 109 22 Cocoa Theobroma cacao L. 105 23 Sunflower Helianthus annuus L. 98 24 Tung oil tree Aleurites fordii Hemsl. 96 Yields of common energy crops are associated with biodiesel production. This is not related to ethanol production, which relies on starch, sugar, and cellulose content instead of oil yields.

Experiments have been carried out to improve the growth and development of sunflower under natural or stress conditions (Gerardo et al., 2013; Nasim et al., 2011; Da Silva et al., 2012). Naz and Bano (2013) reported that the adverse effects of salt stress on sunflower growth could be alleviated by foliar application of salicylic acid alone or in combination with Azospirillum and Pseudomonas inoculations (Table 4). Gholamhoseini et al. (2013) shown that the application of Glomus musseae and Glomus hoi could be critical in the cultivation of sunflowers under arid and semi-arid conditions, where water is the most important factor in determining plant growth and yield. Additionally, Akbari et al. (2011) reported that inoculating the sunflower seeds with plant-growth promoting rhizobacteria increased the

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qualitative and quantitative properties of sunflower significantly, as compared to the control treatment. Table 4. Recent uses of the sunflower during the last years; main or alternative uses make evident the diversity of sunflower Area Food

Description

References

Blends of high linoleic sunflower oil with selected cold pressed soils. Production of florets of sunflower. Tocopherols and phytosterols for the human food market. Sunflower flour as a rich source of high quality proteins. Protein hydrolysis using proteases.

(Ramadan, 2013)

Sunflower products fed to finishing pigs.

(González-Vega and Stein, 2012) (Agy et al., 2013)

(Liang et al., 2013) (Fernández-Cuesta et al., 2012) (Levic et al., 2012) (Tavano, 2013)

Animal Feed

Ingestive behavior and physiological responses of goats fed with sunflower cake. Nutritional value of sunflower meal on broiler (Moghaddam et al., 2012) chickens. Potential nutritive value as source of feed for (Osuga et al., 2012) ruminants in Kenya. Energy Methane production. Biodiesel production. Bioenergy: biotechnology progress and emerging possibilities. Anaerobic digestion of sunflower oil cake. Oil production.

(Fernández-Cegrí et al., 2013; Todorovic et al., 2013) (Iriarte and Villalobos, 2013; Iglesias et al., 2012) (González-Rosas et al., 2013) (De la Rubia et al., 2013) (Spinelli et al., 2012)

Sustainability Of sunflower cultivation within the EU (Spugnoli et al., 2012) Renewable Energy Directive. Sustainable sunflower processing. (Weisz et al., 2013) Economic sustainability of sunflower (Keskin and Dellal, 2011) production. Symbiosis and Plant-Growth Promoting Rhizobacteria Effect of arbuscular mycorrhizal inoculation (Naz and Bano, 2013; Audet on sunflower. and Charest, 2013; Gholamhoseini et al., 2013)

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Area Description Symbiosis and Plant-Growth Promoting Rhizobacteria Bacterial inoculation speeds zinc release from ground tire rubber. A strain of Bacillus subtilis stimulates sunflower growth. Remediation Biodegradation of PAHs. Plant response to lead. Metal accumulation on sunflower. Fertilization, pesticides and environment Foliar fertilization with molybdenum. Fertilization affects the agronomic traits of high oleic sunflower hybrid. Gas exchange in sunflower plants. Effect of different nitrogen level on yield components. Biological control Encrusting offers protection against phytotoxic chemicals. Biological control of Macrophomina phaseolina on sunflower. Allelopathic effects On growth of rice and subsequent wheat crop. On seed germination and seedling growth of Trianthema portulacastrum. Health In vivo evaluation of an oral health toothpaste with sunflower oil. Health benefits of the sunflower kernel.

References (Khoshgoftarmanesh et al., 2012) (López-Valdez et al., 2011)

(Tejeda-Agredano et al., 2013) (Doncheva et al., 2013) (Mahmood et al., 2013; Hao et al., 2012) (Skarpa et al., 2013) (Mohammadi et al., 2013) (Da Silva et al., 2013) (Rafiei et al., 2012)

(Szemruch and Ferrari, 2013) (Ullah, 2010)

(Bashir et al., 2012) (Rawat et al., 2012)

(Schafer et al., 2007) (Holliday and Phillips, 2001)

5. SUNFLOWER ALLELOPATHY Sunflower species are allelopathic in nature; as well cultivated sunflower has great allelopathic potential and inhibits weed-seedling growth of velvet leaf, thorn apple, morning glory, wild mustard and other weeds (Macías et al., 1998a). Two members of the genus Helianthus contain a great quantity of allelopathic compounds. H. annuus is well known for its allelopathic compounds, including sesquiterpene lactones, heliespirones A, annoionones, helibis-abonols and heliannols (Macías et al., 1998b). Heliannols A, D and E have special relevance due to high phytotoxic activity (Macías et al., 1999).

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Figure 1. Some molecular structures of allelopathic compounds presents in sunflower cultivars. A) Annuithrin (sesquiterpene lactone) or Niveusin C, a growth inhibitor. B) Furanoheliangoline, a biologically active molecule. C) Germacranolide, a toxic sesquiterpene lactone (a potent feeding deterrents).

Helianthus tuberosus contains helian-gine and H. annuus contains a sesquiterpene lactone; a heliangolide [Annuithrin or Niveusin C (Figure 1A)] (a growth inhibitor); furanoheliangolide [(Figure 1B) a biologically active]; three additional sesquiterpene lactones: the known compound niveusin B, a germacranolide (Figure 1C) (the tifruticin-type); a 3-ethoxy-niveusin B; an ethoxyheliangolide (Spring et al., 1982) and coumarins (only accumulate in healthy sunflower plants as a response to the variation in environmental conditions that affect field-grown plants). In sunflower, it was reported that the concentrations of scopolin exceeded those in both infected and uninfected plants (Gutiérrez-Mellado et al., 1996). Scopoletin have been described as phytoalexins and allelopathic compounds, being accumulated in response to fungal and parasitic plant infection, insect attack, mechanical injury and treatment with abiotic elicitors such as sucrose and CuCl2, and plant hormones; besides scopoletin has also been shown to have a physiological activity, including the promotion of stomatal closure in sunflower and inhibition of bud growth in pea at very low concentrations (Gutiérrez-Mellado et al., 1996). Annuithrin was tested using a bioassay with Avena straight growth test. The addition of a concentration range from 50 to 180 μM resulted in a linear reduction of growth between 10 and 90%. In fact, annuithrin was shown to have antibacterial qualities. However, fungi and yeast were either less inhibited or not inhibited (minimal inhibitory concentration, MIC 45 μg mL-1 on Bacillus brevis; MIC 90 μg mL-1 on Proteus vulgaris; MIC 90 μg mL-1 on Eremothecium ashbyi; Macías et al., 1996). In addition, in vivo DNA and RNA synthesis in cells of the ascitic form of Ehrlich carcinoma was drastically reduced by annuithrin (at an annuithrin concentration of 20 μg mL-1 about 50% inhibition of DNA synthesis and about 75% inhibition of RNA synthesis) (Spring et al., 1981). It is well known that there are examples of allelopathic cover crops being used for weed management in other crops, as well as other cultural methods to employ allelopathy (Duke, 2010). However, there are still no cultivars of crops being sold with allelopathic properties as a selling point (Cheema and Khaliq, 2000; Tesio and Ferrero, 2010). Enhancement or impartation of allelopathy in crops through the use of transgenes could eventually be used to

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produce such a cultivar. The study of allelopathic crops appears to have a bright future, especially if the scientists can translate the cutting-edge research into technologies that will reduce the reliance on synthetic herbicides, pesticides, and crop protection chemicals. Tesio and Ferrero (2010) reported that the use of allelopathic traits from crops or cultivars with important weed inhibition qualities, together with common weed control strategies, can play an important role in the establishment of sustainable agriculture. It has to be noted that allelopathy may also be another component of desired improved weed management. It will not solve all weed problems in any field, but may help considerably to reduce the population of weeds in the fields (Labrada, 2008).

6. PHYTOREMEDIATION WITH SUNFLOWER Phytoremediation consists of mitigating pollutant concentrations in contaminated soils, water, or air, with plants able to contain, degrade, or eliminate contaminants and its derivatives (Malaviya and Singh, 2012). H. annuus is a plant with not only food and energy values, but also with phytoremediation potential (Seth et al., 2011; Mukhtar et al., 2010). It is one of the most widely studied plants for heavy metal phytoremediation (Kara et al., 2013). However, it is well known that sunflower is able to contain, degrade or eliminate metals (Chen et al., 2012; Ker and Charest, 2010; Lee and Yang, 2010), polycyclic aromatic hydrocarbons (Tejeda-Agredano et al., 2013; Gan et al., 2009) and polychlorinated biphenyls (Fiebig et al., 1997) from soil or water. Investigations with H. annuus have revealed that several heavy metals, including lead, cadmium, copper, zinc and cobalt, accumulate at high concentrations in shoots as well as in roots. Heavy metal uptake is minor in seeds than in roots and shoots. However, few attempts have been made to use plant-growth promoting rhizobacteria to facilitate phytoextraction and cadmium uptake in H. annuus planted in cadmium-contaminated soil (Prapagdee et al., 2013). Sunflower is a documented metal accumulator and its growth on contaminated soil for simultaneous remediation and further energy production has been studied (Marques et al., 2013; Madejon et al., 2003). The good tolerance of sunflower toward pollutants coupled with an increased accumulation/degradation capacity might contribute to an efficient removal of pollutants from soil and water. Clearly it is not an easy job, thus scientists of multidisciplinary areas have to work hard. Additionally, there is a lack of knowledge concerning the pollutants accumulation and antioxidant responses during the growth and development of sunflowers.

7. SUNFLOWER AS A RENEWABLE ENERGY SOURCE Thousands of years ago, people in many regions throughout the world began to process vegetable oils, utilizing whatever food stuffs they had on hand to obtain oils for a variety of cooking purposes. The Chinese and Japanese produced soy bean oil as early as 2000 B.C., while southern Europeans had begun to produce olive oil by 3000 B.C. In Mexico and North America, sunflower seeds were roasted and beaten into a paste before being boiled in water; the oil that rose to the surface was skimmed off (FAO, 2010). During the last decade, an

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increased attention would be observed being paid on the use of sunflower as renewable energy source. Oilseed sunflower is quickly gaining popularity as a feedstock crop for biodiesel because it shares several positive agronomic features with other common oil crops such as canola and soybean; yields well in a variety of conditions, and can be grown easily and profitably at both small farm and large field scales. It is well known that a number of crops can be used for both food and bioenergy production such as sunflower (Kibazohi et al., 2012). Under some circumstances, the potential exist to develop bioenergy systems that allow for synergies between food and energy production. Integrated food and energy systems could produce food crops while simultaneously addressing energy needs (Bogdanski et al., 2010). There is a trend world-wide to grow crops in short rotation or in monoculture (such as sunflower), particularly in conventional agriculture (Bennett et al., 2012). This practice is becoming more prevalent due to a range of factors including economic market trends, technological advances, government incentives, and retailer and consumer demands. Landuse intensity will have to increase further in future in order to meet the demands of growing crops for both bioenergy and food production, and long rotations may not be considered viable or practical. Notwithstanding, evidence indicates that crops grown in short rotations or monoculture often suffer from yield decline compared to those grown in longer rotations or for the first time (Zambrano-Navea et al., 2012). Numerous factors have been hypothesized as contributing to yield decline, including biotic factors such as plant pathogens, deleterious rhizosphere microorganisms, mycorrhizas acting as pathogens, and allelopathy or autotoxicity of the crop, as well as abiotic factors such as land management practices and nutrient availability (Sun et al., 2011). This section identifies gaps in our understanding about the energy production of biomass and the interaction of the ecosystems. Additionally, it has to be remembered that each bioenergy development projects have to include side effects elsewhere in order to shape a sustainable future.

CONCLUSION Sunflower was domesticated in eastern North America and since 3000 B.C. this crop was bred by natives. Thenceforth a wide range of uses of sunflower have been reported throughout the world. Sunflowers are a permanent source of food, oilseed and biofuels because they are well adapted to a variety of conditions and often require fewer agricultural inputs than other more common crops, while under some circumstances, the potential exist to develop bioenergy systems that allow for synergies between food and energy production. Because the sunflower has several potential markets, it is a good choice for growers in both small and large scales. However, scientific, technical or agricultural projects linked with sunflower have to include environmental side effects such as pollution, greenhouse gases emissions, salinization, or energy consumption elsewhere in order to shape a sustainable future.

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employing wetland macrophyte vetiver and energy plant sunflower. Desalin Water Treat 45(1-3): 144-152. Chigeza, G., Mashingaidze, K. & Shanahan, P. (2013). Advanced cycle pedigree breeding in sunflower. I: Genetic variability and testcross hybrid performance for seed yield and other agronomic traits. Euphytica 190(3): 425-438. Da Silva, A. R. A., Bezerra, F. M. L., de Lacerda, F. C. F., Pereira F. J. V. & de Freitas, C. A. S. (2013). Gas exchange in sunflower plants subjected to water deficit at different stages of growth. Rev Cienc Agron 44(1): 86-93. da Silva, A. R. A., Bezerra, F. M. L., Freitas, C. A. S., Pereira, J. V., Andrade, R. R. & Feitosa, D. R. C. (2012). Morphology and biomass of sunflower plants grown under water deficits in different development stages. Rev Bras Eng Agr Amb 16(9): 959-968. de Carvalho, C. G. P. & de Toledo, J. F. F. (2008). Extracting female inbred lines from commercial sunflower hybrids. Pesqui Agropecu Bras 43(9): 1159-1162. de la Rubia, M., Fernández-Cegrí V, Raposo F & Borja R (2013). Anaerobic digestion of sunflower oil cake: a current overview. Water Sci Technol 67(2): 410-417. Doncheva, S., Moustakas, M., Ananieva, K., Chavdarova, M., Gesheva, E., Vassilevska, R. & Mateev, P. (2013). Plant response to lead in the presence or absence EDTA in two sunflower genotypes (cultivated H. annuus cv. 1114 and interspecific line H. annuus x H. argophyllus). Environ Sci Pollut Res 20(2): 823-833. Duke, S. O. (2010). Allelopathy: Current status of research and future of the discipline: A Commentary Allelopathy J 25(1): 17-30. Emami-Bistghani, Z., Siadat, S. A., Torabi, M., Bakhshande, A., Alami, S. K. & Hiresmaeili, H. (2012). Influence of plant density on light absorption and light extinction coefficient in sunflower cultivars Res Crop 13(1): 174-179. Esmaeli, M., Javanmard, H. R., Nassiry, B. M. & Soleymani, A. (2012). Effect of different plant densities and planting pattern on sunflower (Helianthus annuus L.) cultivars grown under climatic conditions of Isfahan region of Iran. Res Crop 13(2): 517-520. FAO (2010). Sunflower crude and refined oils. Food and Agriculture Organization of the United Nations. Retrieved from: http://www.responsibleagroinvestment. org /sites/ responsibleagroinvestment.org/files/FAO_Agbiz%20handbook_oilseeds_0.pdf. (Verified June 5, 2013). FAOSTAT (2011). Production of sunflower seed throughout the world. Retrieved from: http://faostat3.fao.org/home/index.html#DOWNLOAD. (Verified May 6, 2013). Fernández-Cegrí, V., Raposo, F., de la Rubia, M. A. & Borja, R. (2013). Effects of chemical and thermochemical pretreatments on sunflower oil cake in biochemical methane potential assays. J Chem Technol Biot 88(5): 924-929. Fernández-Cuesta, A., Nabloussi, A., Fernández-Martínez, J. M. & Velasco, L. (2012). Tocopherols and phytosterols in sunflower seeds for the human food market. Grasas y Aceites 63(3): 321-327. Fiebig, R., Schulze, D., Chung, J.-C. & Lee, S.-T. (1997). Biodegradation of polychlorinated biphenyls (PCBs) in the presence of a bioemulsifier produced on sunflower oil. Biodegradation 8(2): 67-75. Fozia A., Muhammad AZ., Muhammad A. & Zafar MK. (2008). Effect of chromium on growth attributes in sunflower (Helianthus annuus L.). J Environ Sci (China) 20(12): 1475-1480.

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Gan, S., Lau, E. V. & Ng, H. K. (2009). Remediation of soils contaminated with polycyclic aromatic hydrocarbons (PAHs). J Hazard Mater 172(2-3): 532-549. Gerardo, R., Boem, F. H. G. & Fernandez, M. C. (2013). Severe phosphorus stress affects sunflower and maize but not soybean root to shoot allometry. Agron J 105(5): 12831288. Ghaffari, M., Toorchi, M., Valizadeh, M. & Shakiba, M. R. (2012). Morpho-physiological screening of sunflower inbred lines under drought stress condition Turk J Field Crop 17(2): 185-190. Gholamhoseini, M., Ghalavand, A., Dolatabadian, A., Jamshidi, E. & Khodaei-Joghan, A. (2013). Effects of arbuscular mycorrhizal inoculation on growth, yield, nutrient uptake and irrigation water productivity of sunflowers grown under drought stress. Agr Water Manage 117: 106-114. González-Rosas, A., Miranda-Gómez, J. M., Padmasree, K. P. & Fernández-Luqueño, F. (2013). How Green is bioenergy? A review on myths, challengues, biotechnology progress and emerging possibilities. Sci Res Essays 8: 532-542. González-Vega, J. C. & Stein, H. H. (2012). Amino acid digestibility in canola, cottonseed, and sunflower products fed to finishing pigs. J. Anim. Sci. 90(12): 4391-4400. Gutiérrez-Mellado, M. C., Edwards, R., Tena, M., Cabello, F., Serghini, K. & Jorrín, J. (1996). The production of coumarin phytoalexins in different plant organs of sunflower (Helianthus annuus L.). J. Plant Physiol 149(3-4): 261-266. Hao, X. Z., Zhou, D. M., Li, D. D. & Jiang, P. (2012). Growth, cadmium and zinc accumulation of ornamental sunflower (Helianthus annuus L.) in contaminated soil with different amendments. Pedosphere 22(5): 631-639. Harter, A. V., Gardner, K. A., Falush, D., Lentz, D. L., Bye, R. A. & Rieseberg, L. H. (2004). Origin of extant domesticated sunflower in eastern North America. Nature 430(6996): 201-205. Holliday, R. & Phillips, K. (2001). Health benefits of the sunflower kernel. Cereal Foods World. 46(5): 205-208. Iglesias, L., Laca, A., Herrero, M. & Díaz, M. (2012). A life cycle assessment comparison between centralized and decentralized biodiesel production from raw sunflower oil and waste cooking oils. J Clean Prod 37: 162-171. Iriarte, A. & Villalobos, P. (2013). Greenhouse gas emissions and energy balance of sunflower biodiesel: Identification of its key factors in the supply chain. Resour Conserv Recy 73: 46-52. Jabeen, N. & Ahmad, R. (2012). Improvement in growth and leaf water relation parameters of sunflower and safflower plants with foliar application of nutrient solutions under salt stress Pak J Bo 44(4): 1341-1345. Kara, Y., Koca, S., Vaizogullar, H. E. & Kuru, A. (2013). Studying phytoremediation capacity of jojoba (Simmondsia chinensis) and sunflower (Helianthus annuus) in hydroponic systems. Curr Opin Biotech 24(1): S34. Ker, K. & Charest, C. (2010). Nickel remediation by AM-colonized sunflower. Mycorrhiza 20(6): 399-406. Keskin, G. & Dellal, I. (2011). Economic sustainability of sunflower production in thrace region of turkey. J Environ Prot Ecol 12(1): 245-250. Khaleghizadeh, A. (2011). Effect of morphological traits of plant, head and seed of sunflower hybrids on house sparrow damage rate. Crop Prot 30(3): 360-367.

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Khan, A., Lang, I., Amjid, M., Shah, A., Ahmad, I. & Nawaz, H. (2013). Inducing salt tolerance on growth and yield of sunflower by applying different levels of ascorbic acid. J Plant Nutr 36(8): 1180-1190. Khoshgoftarmanesh, A., Behzadan, H., Sanaei, A. & Chaney, R. (2012). Bacterial inoculation speeds zinc release from ground tire rubber used as Zn fertilizer for corn and sunflower in a calcareous soil. Plant Soil 361(1-2): 71-81. Kibazohi O, Rincon-Perez L. E, Felix E & Cardona-Alzate C.A (2012). Technical and economical analysis for biofuel production from sunflower. In Bioenergy and food security: The BEFS analysis for Tanzania-Sunflower biodiesel, water, and household food security, 108 (Ed FAO). Tanzania: FAO. Kunduraci, B. S., Bayrak A., & Kiralan, M. (2010). Effect of essential oil extracts from oregano (Origanum onites L.) leaves on the oxidative stability of refined sunflower oil. Asian J Chem 22(2): 1377-1386. Labrada, R. (2008). Allelopathy as a tool for weed management. Allelopathy J 22(2): 283287. Lee, M. &Yang, M. (2010). Rhizofiltration using sunflower (Helianthus annuus L.) and bean (Phaseolus vulgaris L. var. vulgaris) to remediate uranium contaminated groundwater. J Hazard Mater 173(1-3): 589-596. Levic, J., Ivanov, D., Sredanovic, S., Jovanovic, R., Colovic, R. & Vukmirovic, D. (2012). Sunflower flour as a rich source of high quality proteins. Agro Food Industry HI-TECH 23(4): 13-15. Liang, Q., Cui, J., Li, H., Liu, J. & Zhao, G. (2013). Florets of sunflower (Helianthus annuus L.): potential new sources of dietary fiber and phenolic acids. J Agric Food Chem 61(14): 3435-3442. López-Valdez, F., Fernández-Luqueño, F., Ceballos-Ramírez, J. M., Marsch, R., OlaldePortugal, V. & Dendooven, L. (2011). A strain of Bacillus subtilis stimulates sunflower growth (Helianthus annuus L.) temporarily. Sci Hort 128(4): 499-505. Macías F. A., Torres A., Molinillo J. M. G., Varela R. M., & Castellano D. (1996). Potential allelopathic sesquiterpene lactones from sunflower leaves. Phytochemistry 43(6): 12051215. Macías F. A., Varela R. M., Torres A. & Molinillo J. M. G (1998b). Heliespinone A, The first member of a novel family of bioactive sesquiterpenes. Tetrahedron Lett 39: 427-430. Macías F. A., Varela R. M., Torres A., Oliva R. M. & Molinillo J. (1998a). Bioactive norsesquiterpenes from Helianthus annuus with potential allelopathic activity. Phytochemistry 48: 631-636. Macías F. A., Varela R. M., Torres A. & Molinillo, J. M. G. (1999). New bioactive plant heliannuols from cultivar sunflower leaves. J Nat Prod 62(12): 1636-1639. Madejon, P., Murillo, J., Maranon, T., Cabrera, F. & Soriano, M. (2003). Trace element and nutrient accumulation in sunflower plants two years after the Aznalcollar mine spill. Sci Total Environ 307(1-3): 239-257. Mahmood, S., Ishtiaq, S., Malik, M. I. & Ahmed, A. (2013). Differential growth and photosynthetic responses and pattern of metal accumulation in sunflower (Helianthus annuus L.) cultivars at elevated levels of lead and mercury. Pak J Bot 45(1): 367-374. Malaviya, P. & Singh, A. (2012). Phytoremediation Strategies for Remediation of UraniumContaminated Environments: A Review. Crit Rev Env Sci Tec 42(24): 2575-2647.

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Marques, A. P. G. C., Moreira, H., Franco, A. R., Rangel, A. O. S. S. & Castro, P. M. L. (2013). Inoculating Helianthus annuus (sunflower) grown in zinc and cadmium contaminated soils with plant growth promoting bacteria - Effects on phytoremediation strategies. Chemosphere 92(1): 74-83. Moghaddam, H. N., Salari, S., Arshami, J., Golian, A. & Maleki, M. (2012). Evaluation of the nutritional value of sunflower meal and its effect on performance, digestive enzyme activity, organ weight, and histological alterations of the intestinal villi of broiler chickens. J Appl Poult Res 21(2): 293-304. Mohammadi, K., Heidari, G., Javaheri, M., Rokhzadi, A., Nezhad, M. T. K., Sohrabi, Y. & Talebi, R. (2013). Fertilization affects the agronomic traits of high oleic sunflower hybrid in different tillage systems. Ind Crops Prod 44: 446-451. Mukhtar, S., Bhatti, H. N., Khalid, M., Anwar-ul-Haq, M. & Shahzad, S. M. (2010). Potential of sunflower (Helianthus annuus L.) for phytoremediation of nickel (Ni) and lead (Pb) contaminated water. Pak J Bot 42(6): 4017-4026. Muller, M. H., Latreille, M. & Tollon, C. (2011). The origin and evolution of a recent agricultural weed: population genetic diversity of weedy populations of sunflower (Helianthus annuus L.) in Spain and France. Evol Appl 4(3): 499-514. Nasim, W., Ahmad, A., Wajid, A., Akhtar, J. & Muhammad, D. (2011). Nitrogen effects on growth and development of sunflower hybrids under agro-climatic conditions of Multan. Pak J Bot 43(4): 2083-2092. Naz, R. & Bano, A. (2013). Influence of exogenously applied salicylic acid and plant growth promoting rhizobacteria inoculation on the growth and physiology of sunflower (Helianthus annuus L.) under salt stress. Pak J Bot 45(2): 367-373. NSA (2013). All about sunflower. National Sunflower Association. Retrieved from: http:// www.sunflowernsa.com/all-about/history/. (Verified June 6, 2013). Olowe, V. I. O. & Adeyemo, A. Y. (2009). Enhanced crop productivity and compatibility through intercropping of sesame and sunflower varieties. Ann Appl Biol 155(2): 285-291. Onemli, F. (2012). Impact of climate changes and correlations on oil fatty acids in sunflower. Pak J Agri Sci 49(4): 455-458. Osuga, I. M., Abdulrazak, S. A., Muleke, C. I. & Fujihara, T. (2012). Potential nutritive value of various parts of wild sunflower (Tithonia diversifolia) as source of feed for ruminants in Kenya. J Food Agric Environ 10(2): 632-635. Prapagdee, B., Chanprasert, M. & Mongkolsuk, S. (2013). Bioaugmentation with cadmiumresistant plant growth-promoting rhizobacteria to assist cadmium phytoextraction by Helianthus annuus. Chemosphere 92(6): 659-666. PSD-USDA (2011). Production, Supply & Distribution On Line:- United State Department of Agriculture. Retrieved from: http://www.fas.usda.gov/psdonline/psdquery.aspx. (Verified June 9, 2013). Radanielson, A. M., Lecoeur, J., Christophe, A. & Guilioni, L. (2012). Use of water extraction variability to screen for sunflower genotypes well adapted to soil water limitation. Funct Plant Biol 39(12): 999-1008. Rafiei, H., Soleymani, A. & Golparvar, A. R. (2012). Effect of different nitrogen levels on yield components and seed yield of three sunflower (Helianthus annuus L.) cultivars. Res Crop 13(3): 996-1000.

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Ramadan, M. F. (2013). Healthy blends of high linoleic sunflower oil with selected cold pressed oils: Functionality, stability and antioxidative characteristics. Ind Crops Prod 43: 65-72. Rawat, L. S., Narwal, S. S., Kadiyan, H. S., Maikhuri, R. K., Negi, V. S. & Pharswan, D. S. (2012). Allelopathic effects of sunflower on seed germination and seedling growth of Trianthema portulacastrum. Allelopathy J 30(1): 11-22. Rawat, L. S., Negi, V. S. & Maikhuri, R. K. (2010). Allelopathic effect of sunflower (Helianthus annuus) rhizosphere soil on germination and seedling growth of kharif crops and weeds. Natl Acad Sci Lett (India) 33(9-10): 271-278. Saensee, K., Machikowa, T. & Muangsan, N. 2012. Comparative performace of sunflower synthetic varieties under drought stress. Int J Agric Biol 14(6): 929-934. Saleem, M. F., Ma, B. L., Malik, M. A., Cheema, M. A. & Wahid, M. A. (2008). Yield and quality response of autumn-planted sunflower (Helianthus annuus L.) to sowing dates and planting patterns. Can J Plant Sci 88(1): 101-109. Schafer, F., Adams, S. E., Nicholson, J. A., Cox, T. F., McGrady, M. & Moore, F. (2007). In vivo evaluation of an oral health toothpaste with 0.1% vitamin E acetate and 0.5% sunflower oil (with vitamin F). Int Dent J 57(2): 119-1123. Seassau, C., Dechamp-Guillaume, G., Mestries, E. & Debaeke, P. (2012). Low plant density can reduce sunflower premature ripening caused by Phoma macdonaldii. Eur J Agron 43: 185-193. Seghatoleslami, M. J., Bradaran, R., Ansarinia, E. & Mousavi, S. G. (2012). Effect of irrigation and nitrogen level on yield, yield components and some morphological traits of sunflower. Pak J Bot 44(5): 1551-1555. Seth, C., Misra, V., Singh, R. R. & Zolla, L. (2011). EDTA-enhanced lead phytoremediation in sunflower (Helianthus annuus L.) hydroponic culture. Plant Soil 347(1-2): 231-242. Skarpa, P., Kunzova, E. & Zukalova, H. (2013). Foliar fertilization with molybdenum in sunflower (Helianthus annuus L.). Plant Soil Environ 59(4): 156-161. Spinelli, D., Jez, S. & Basosi, R. (2012). Integrated Environmental Assessment of sunflower oil production. Process Biochemistry 47(11): 1595-1602. Spring, O., Albert, K. & Gradmann, W. (1981). Annuithrin, a new biologically active germacranolide from Helianthus annuus. Phytochemistry 20(8): 1883-1885. Spring, O., Albert, K. & Hager, A. (1982). Three biologically active heliangolides from Helianthus annuus. Phytochemistry 21(10): 2551-2553. Spugnoli, P., Dainelli, R., D'Avino, L., Mazzoncini, M. & Lazzeri, L. (2012). Sustainability of sunflower cultivation for biodiesel production in Tuscany within the EU Renewable Energy Directive. Biosyst Eng 112(1): 49-55. Sun, Y. Y., Jiang, G. Y., Wei, X. C. & Liu, J. G. (2011). Autotoxicity effects of soils continuously cropped with tomato. Allelopathy J 28(2): 135-144. Szemruch, C. L. & Ferrari, L. (2013). Encrusting offers protection against phytotoxic chemicals and maintains the physiological quality of sunflower (Helianthus annuus) seeds. Seed Sci Technol 41(1): 125-132. Tavano, O. L. (2013). Protein hydrolysis using proteases: An important tool for food biotechnology. J Mol Catal B: Enzym 90: 1-11. Tejeda-Agredano, M. C., Gallego, S., Vila, J., Grifoll, M., Ortega-Calvo, J. J. & Cantos, M. (2013). Influence of the sunflower rhizosphere on the biodegradation of PAHs in soil. Soil Biol Biochem 57: 830-840.

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Tesio, F. & Ferrero, A. (2010). Allelopathy, a chance for sustainable weed management. Int J Sust Dev World 17(5): 377-389. Thomaz, G. L., Zagonel, J., Colasante, L. O. & Nogueira, R. R. (2012). Yield of sunflower and oil seed content as a function of air temperature, rainfall and solar radiation. Cienc Rural 42(8): 1380-1385. Todorovic, Z. B., Stamenkovic, O. S., Stamenkovic, I. S., Avramovic, J. M., Velickovic, A. V., Bankovic-Ilic, I. B. & Veljkovic, V. B. (2013). The effects of cosolvents on homogeneously and heterogeneously base-catalyzed methanolysis of sunflower oil. Fuel 107: 493-502. Ullah, H. (2010). Biological control of Macrophomina phaseolina on sunflower in Pakistan. Phytopathology 100(6): s128. Weisz, G. M., Carle, R. & Kammerer, D. R. (2013). Sustainable sunflower processing -II. Recovery of phenolic compounds as a by-product of sunflower protein extraction. Innov Food Sci Emerg Technol 17: 169-179. Zambrano-Navea, C. L., Bastida, F. & Gonzalez-Andujar, J. L. (2012). Herbicidal strategies to control Phalaris brachystachys in a wheat-sunflower rotation: a simulation approach. Span J Agric Res 10(4): 1101-1106. Zheljazkov, V. D., Vick, B. A., Baldwin, B. S., Buehring, N., Astatkie, T. & Johnson, B. (2012). Effect of planting date, nitrogen rate, and hybrid on sunflower. J Plant Nutr 35(14): 2198-2210.

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In: Sunflowers Editor: Juan Ignacio Arribas

ISBN: 978-1-63117-347-9 © 2014 Nova Science Publishers, Inc.

Chapter 2

FLORAL BIOLOGY OF SUNFLOWERS: A HISTOLOGICAL AND PHYSIOLOGICAL ANALYSIS Basudha Sharma, Rashmi Shakya and Satish C. Bhatla* Laboratory of Plant Physiology and Biochemistry, Department of Botany, University of Delhi, Delhi, India

ABSTRACT The development of sunflower inflorescence can be considered under three phases, namely inflorescence initiation, floret development and anther formation. Floret primordia appear at the rim of the receptacle where ray or disc florets are generated. Disc florets are arranged in Fibonacci series whereby a spiral pattern emerges as new florets arise in rows of bumps consisting of a bract and a floret. Floral morphogenesis in sunflower occurs according to the ABC model, whereby genes of the MADS box are activated. Anthesis of disc florets is a phytochrome-mediated response and is also modulated by plant hormones, such as auxins. The disc florets are hermaphrodite and protandrous in nature, whereas the ray florets are sterile, incomplete and have an attractive, fused and flag-like corolla. Stigma in sunflower is semi-dry in nature, producing lipid rich exudates in the crevices of the adjacent papillae. Stigma undergoes physiological maturity with the passage of development from bud, staminate and, finally to the pistillate stage. The production of extracellular lipid rich secretions is initiated at the staminate stage of stigma development and increases at the receptive stage through the availability of elaioplasts and endoplasmic reticulum network in the basal regions of the papillae. Transfer cells, earlier identified only in the wet type of stigma, are also present in the transmitting tissue of sunflower stigma. Neutral esters and triacylglycerols (TAGs) are the major lipidic constituents in pollen grains and stigma, respectively. Lignoceric acid (24:0) and cis-11-eicosenoic acid (20:1) are specifically expressed only in the pollen coat. Similar long-chain fatty acids have earlier been demonstrated to play a significant role during the initial signalling mechanism leading to hydration of pollen grains on the stigma surface. Lipase activity is expressed both in the pollen grains and stigma papillae. Stigma exhibits a better expression of acyl-ester hydrolase activity the pollen grains. Specific expression of lignoceric acid (24:0) in the pollen coat and *

Corresponding author: Professor S.C.Bhatla; E mail: [email protected].

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Basudha Sharma, Rashmi Shakya and Satish C. Bhatla localization of lipase in pollen and stigma are likely to have possible roles during pollenstigma interaction. During the course of stigma development in sunflower, a correlation is evident in the accumulation of reactive oxygen species (ROS), nitric oxide (NO) and the activities of ROS scavenging enzymes [superoxide dismutase (SOD) and peroxidase (POD)]. Mn-SOD (mitochondria localized) and Cu/Zn-SOD (cytoplasmic) exhibit differential expression during the staminate stage of stigma development. An increase in total SOD activity at the staminate stage is followed by a peak of POD activity during the pistillate stage of stigma development, indicating the sequential action of the two enzymes in scavenging ROS in maturing stigma. The number of POD isoforms increases with the passage of stigma development and two POD isoforms are unique to pistillate stage. This highlights their role in ROS scavenging mechanism. ROS and NO accumulation exhibit reverse trends during pollen-stigma interaction. All these recent findings indicate the modulation of floral development in sunflower by an array of biomolecular signalling components which influence development through a series of cross-talk mechanisms.

Keywords: Lipids, nitric oxide, non-specific esterase, pollen, peroxidase, reactive oxygen species, superoxide dismutase, stigma, pollen-stigma interaction

INTRODUCTION A large diversity of floral structures of varying complexities are evident in plants for the attraction of pollinators. Cross-pollinated plants exhibit a kind of synchrony among themselves and also with their pollinators in order to bring about optimal seed set. It is necessary that plants must flower at the correct time of the year for optimal reproductive fitness. Such a strong correlation with the environment for the onset of flowering poses many questions about the mechanisms of sensing of the environmental signals by the plants and also about the sequence of biochemical events which ultimately bring about flowering in response to the environmental signals. A vegetative shoot bud exhibits noteworthy differences when compared with a floral bud, in terms of the constituent forms and types of cells. A change in the fate of cells at the shoot apex is governed by the expression of a set of genes, leading to various biochemical events in the shoot apex which bring about floral evocation, i.e the ability of the apical meristem to produce flowers. Floral evocation is regulated by endogenous factors, such as circadian rhythms and hormones, and exogenous factors such as photoperiod and temperature. The initiation of four types of floral organs from the floral meristem is observed in whorls around the flanks of the meristem. The floral primordial start as small bumps of cells and their further development into reproductive structures is governed by the environmental signals, various metabolic events and also by the activation of specific genetic programs. Broadly, an attempt has been made in the following chapter to understand the histological, physiological and biochemical changes taking place in the shoot apex of sunflower during this process of phase change, i.e., transition from adult vegetative phase to adult reproductive phase. The initiation of whorls of disc florets in the inner core and development of peripheral ray florets during capitulum development in sunflower is a gradual process, showing various stages of development of florets in a capitulum. The present chapter provides detailed information on the ultrastructural changes associated with stigma development in sunflower. These features have been analyzed in relation with the

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biochemical events accompanying stigma maturation. Likewise, a detailed structural analysis of pollen (intact and germinating) has been discussed. Finally, the mechanism of pollenstigma interaction has been analyzed under natural and experimental conditions.

PATTERN OF FLORAL DEVELOPMENT AND ITS MODULATION BY LIGHT, HORMONES AND GENETIC FACTORS Members of Asteraceae maximize their reproductive output by condensing inflorescence and forming a capitulum. Vegetative apex is indeterminate, domed and densely meristematic whereas reproductive apex broadens, flattens and becomes determinate (Teeri et al., 2006). Sunflower inflorescence is a disc-shaped capitulum located at the shoot tip and its shape and size vary according to the cultivar, season and agricultural conditions (Weiss, 2000). The capitulum is surrounded by three rows of ovate to ovate-laceolate involucral bracts or phyllaries which function as sepals and protect the capitulum during its development (Figure 1). Various phases of reproductive development in sunflower have earlier been categorized under nine stages (Schneiter and Miller, 1981). Beginning from the initiation of floral bud (R1) to the attainment of physiological maturity (R9), R5 marks the beginning of flowering. This stage (R5) is further subdivided from R5.1 to R5.2 and so on, representing the percent of disc florets which have completed or are flowering. The florets in the capitulum are arranged in a spiral and geometric pattern (Hernández and Green, 1993). Floret primodia appear at the rim of the receptacle where ray or disc florets are generated. Disc florets are arranged in Fibonacci series leading to the emergence of a spiral pattern as new florets arise in rows of bumps consisting of a bract and a floret (Hernández, 1997). Ray florets (outer) are sterile, incomplete and have an attractive, fused and flag-like corolla whereas disc florets (inner) are complete and exhibit centripetal maturation pattern. Each disc floret consists of an inferior ovary, two pappus scales (modified sepals) and a tubular corolla, which is fused, except at the tip (Figure 2). Flowering begins with the unfolding of the ray florets in the capitulum. Disc florets gradually open in whorls towards the centre of the head, as a consequence exhibiting different stages of floret maturation in a single capitulum. Such a pattern of development also increases flowering time of a capitulum, thereby attracting insects for pollination. The maturation stages of disc florets are referred as bud, staminate, transitional and pistillate (Figure 2). The disc florets are protandrus and are cross-pollinated by insects, particularly bees. At the bud stage, disc florets exhibit the development of corolla, androecium and gynoecium. Stigma is clasped and the pollen grains inside the anther lobes have a well developed exine. Anthesis begins in the morning at the staminate stage when staminal filaments elongate and black syngenesious stamens are exposed through the tubular corolla. Protandry in sunflower is induced by photoperiod and corolla has been suggested to be the site of light perception, stimulating the growth of anther filament (Lobello et al., 2000). Elongation of the antheridial filaments is initiated after the dark period and occurs for about 2-6h. The pollen grains are then released inside the anther tube as three-celled structures. At the transitional stage, the stigma elongates through the anther lobe and hairy pseudopapaillae are observed at the apex of the anther tube. Members of Asteraceae exhibit the phenomenon of secondary pollen presentation whereby pollen grains are relocated from the anther to another floral organ, which then presents pollen for pollination. The pollen grains left in the

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anther tube adhere to the sweeping hair (pseudopapillae) of the stigma and they are mechanically forced out and exposed to the pollinators (Hong et al., 2008). Stigma exhibits biphasic growth kinetics at the pistillate stage during which it first elongates, detaches along the median, and the tip curls outward (Sammataro et al., 1985).

Figure 1. Stages of the development of capitulum in sunflower.

Figure 2. Various stages of disc floret development in sunflower. A: Young bud, B: Mature bud, C: Staminate stage, D: Transitional stage, E: Pistillate stage.

Floret maturation is reported to be under the control of phytochrome and plant hormones (Baroncelli et al., 1990; Koning, 1983). The development of capitulum from R2 to R4 is affected by photoperiod (Rezadoust et al., 2010). Although sunflower is considered to be a day neutral plant, the development of floral buds and their maturation is known to be affected

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by daylength. Light leads to an enhancement of photosynthesis, causing growth of the tissue for the formation of floral bud. Short days are known to modulate anthesis by promoting postinitiation development of the floral buds (Marc and Palmer, 1981). Depending on the influence of photoperiod in green house plantations (from the period of emergence to floral bud development), different sunflower genotypes have been classified as long-day, short-day or day-neutral plants. In addition, some genotypes have been observed to be ambiphotoperiodic, a condition in which the floral buds can develop in long or short day conditions but their further development is delayed in intermediate day length (Goyne and Schneiter, 1987). The young capitula (heads) exhibit heliotropism which is marked by the eastward movement of head in the morning and its westward turning along the direction of sun. As the capitulum matures, the opened heads are locked in the eastward direction (Weiss, 2000). An eastwardly direction of the capitulum dries the night dew in the morning hours and decreases the possibility of fungal attack. It also prevents overheating of the developing stigmas and preserves pollen viability, consequently enhancing the efficiency of fertilization. It has been proposed that the heliotropic movement of the young capitulum is related to auxin distribution in the actively growing parts of the plants (Weiss, 2000). Growing regions of plants contain relatively higher concentration of IAA than as compared to the fully developed plant parts resulting in the accumulation of assimilated substances (Duca, 2006). The content of gibberellic acid also increases, particularly in the cytoplasmic male sterile lines. Gibberellic acid is involved in floral induction and the process of sexual differentiation (Duca et al., 2003; Duca, 2006). The role of phytochrome in favouring protandry, and hence cross pollination, has also been established (Baroncelli et al., 1990; Lobello et al., 2000). Variations in photoperiod and relatively higher concentrations of gibberellic acid are known to cause a deviation in floral development (Blackman et al., 2011). The elongation of antheridial filaments is stimulated by auxins (IAA and NAA) or light. Auxins are known to be involved in the light-regulated expansion of cells. In vitro experiments have confirmed that auxins can reduce the inhibitory action of red or dichromatic treatment (far red + red light) on the elongation of antheridal filaments. Filament elongation caused by light and dark cycles or auxins, is also known to be dependent on the critical phase of growth of the florets (Lobello et al., 2000). High concentrations of gibberellic acid (GA3) are known to inhibit filament and style elongation in favourable photoperiodic conditions (Lobello et al., 2000). Light, thus, plays an important role in altering the availability of gibberellic acid which is essential for cell expansion. The development of floral organs in each whorl is regulated by the differential activities of various genes encoding MADS-box transcription factors (Dezar et al., 2003). The identification of the genes responsible for sunflower morphogenesis has highlighted two types of floral differentiation. Reproductive meristem follows the ABC model of flower development which refers to the class of genes that are required for the development of whorls of sepals, petals, stamens or carpels. The genes corresponding to ABC encode the MADS-box transcription factors which are conserved motifs controlling transition from vegetative to reproductive growth, thus determining the identity of floral meristem and organs. Floral homeotic mutants of Arabidopsis thaliana, Antirrhinum majus and Petunia hybrida have extensively been used to understand the controlling factors for the initiation of floral buds by the ABC model (Coen and Meyerowitz, 1991; Angenent et al., 1995). Seven full length cDNAs of HAM genes (Helianthus annuus MADS) have been isolated and their control in the development of pistil, stamen and petals has been established in sunflower (Shulga et al., 2008). Blot hybridization from different parts of sunflower has further revealed

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that HAM 75 and HAM 92 genes are expressed in petals, seed coat HAM 45 is expressed in the ovule and HAM 59 is expressed in the ovules, stamens and pistil. HAM 59 is expressed essentially in the disc florets and is absent in ray florets, causing sterility in the ray florets. HAM 59 expression during ray floret initiation seems to be important for the structural and functional differences in the developing inflorescence. A correlation between the structure and function of these proteins is evident. Since HAM genes code for proteins that belong to different subfamilies of MADS-box, a duplication of the sunflower genome during evolution has been proposed. Antimicrobial proteins are known to be produced by plants which contribute to resistance. Among the antimicrobial peptides, cysteine-rich thionine, lipidtransfer proteins, defensins and snakin have been described. In sunflower, defensins have been reported to accumulate as florets mature. The defensins are known to be localized mainly in cell wall or vacuoles (Urdangarín et al., 2000).

Figure 3. Structural analysis of receptive stigma and pollen in sunflower. A: Scanning electron micrograph of receptive stigma surface (65X); B: Transverse section of mature stigma showing the presence of papillae (P), vascular strand (VS) and secretory canal (SC) (400X); C: Localization of proteins in the papillae and transmitting tissue (TT) after staining with mercuric bromophenol solution (400X); D: Electron micrograph from the basal region of the papillae showing the accumulation of extracellular secretions (1,150X); E: Transmission electron micrograph showing cluster of mitochondria at the base of papillae at the staminate stage of stigma development (8000X); F: Transmission electron micrograph showing transfer cells in the transmitting tissue below the papillae in the receptive stigma; G: Transmission electron micrograph of cells of transmitting tissue showing plasmodesmatal connections (1,150X); H and I: Transmission electron micrograph of the pollen wall showing spinular region (4,600X) and the inter-spinular region (8,400X). Abbreviations: P, Papillae; PP, Pseudopapillae; E, Extracellular secretions; PD, Plasmodesmata; N, Nucleus; V,Vacuole; M, Mitochondria; WI, Wall Ingrowths; TT, Transmitting tissue; VS, Vascular strand.

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STRUCTURAL ANALYSIS OF DEVELOPING STIGMA AND TRANSMITTING TISSUE Mature stigma in sunflower is forked, bifid and consists of two parts- the peripheral brush-like pseudopapillae and the inner, thin, finger-like papillae, which are raised and densely arranged outgrowths of the peripheral cells (Figure 3A). The papillate surface of stigma increases the pollen capturing area and ensures the proper interaction of pollen with stigma surface (Heslop-Harrison and Shivanna, 1977). A transverse section of stigma shows the presence of a four-layered transmitting tissue immediately below the papillae where the cells are surrounded by an intercellular matrix. Beneath the transmitting tissue is the ground tissue, in the centre of which is a vascular canal and a large secretory canal (Figure 3B). Papillae and the transmitting tissue are abundant in proteins (Figure 3C). In order to attain receptivity, stigma undergoes many structural and physiological changes which allows it to become competent for the directional growth of pollen tubes (Kandasamy et al., 1994; Yi et al., 2006). Transmission electron microscopic analysis has revealed that the papillae in the bud stage of developing stigma are densely cytoplasmic as compared to the ones in mature stigma, in which they are elongated and vacuolated. The papillae have a large nucleus with a prominent nucleolus and abundant mitochondria. Nature of stigma surface in sunflower has remained controversial over the years. It has earlier been described by some investigators as dry and lacking secretions on the surface (Heslop-Harrison and Shivanna, 1977; Vithanage and Knox, 1977; Gotelli et al., 2010). Recently, it has also been described as semi-dry in sunflower (Shakya and Bhatla, 2010), and in some other members of Asteraceae, namely Senecio squalidus (Hiscock et al., 2002a), Lessingianthus grandiflorus and Lucilia lycopodioides (Teixeira et al., 2011). Semi-dry stigma possesses a surface cuticle on the papillae similar to the dry stigma which, however, is not continuous at the base of the papillae. Like the wet stigma, mature semi-dry stigma possesses a small amount of extracellular secretion at the base of the papillae (Allen et al., 2010). The initiation of secretory activity in Helianthus annuus is observed at the staminate stage which leads to an accumulation of lipid-rich extracellular secretion at the base of the papillae during the pistillate stage of stigma development (Figure 3D; Sharma, 2012). The secretory activity coincides with the presence of endoplasmic reticulum and elaioplasts in the basal region of the papillae. These secretions accumulate in the intercellular and subcuticular gaps, causing a disruption of the cuticle and release of exudates on the stigma surface. Dry stigma in Brassica rapa, Arabidopsis thaliana and Raphanus sp. do not show extracellular secretions and the cuticle extends to the base of the papillae (Hiscock et al., 2002a). Lipids are known to be the major components of the exudates in some wet stigmas and are responsible for pollen hydration (Cresti et al., 1986). During the course of evolution, the function of hydration has been taken over by the pollen coat in dry stigmas (Sage et al., 2009). Probably, the pollen coat and lipids on the stigma surface of semi-dry stigma aid during the processes of adhesion and hydration. The cells of transmitting tissue are loosely arranged below the papillae. The secretory products of the transmitting tissue in sunflower are rich in pectins and other polysaccharides, as has also been observed in Tibouchina sp. (Ciampolini et al., 1995), Vitis vinifera (Ciampolini et al., 1996), Passiflora edulis (Souza et al., 2006). The extracelluar secretions of the transmitting tissue increase as stigma attains maturity, showing that it acts as a source of

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nutrition for the growing pollen tube. Cells of the transmitting tissue are polyhedral to spherical, with a prominent nucleus, plastids and endoplasmic reticulum. Cells of the transmitting tissue can be differentiated into three types. Type I cells are vacuolated with parietal cytoplasm and few mitochondria. Type II cells have a large nucleus, several mitochondria and a vacuole smaller than that in Type I cells. Type III cells have a dense cytoplasm (Figure 3G). Some cells of the transmitting tissue show internal ramification (Figure 3F). The finger-like projections into the cytoplasm formed by secondary wall ingrowths increase the surface-volume ratio for enhanced metabolic activities across the cells. The involvement of these cells in secretory activity has also been reported in the members of Rosaceae (Heslop-Harrison and Shivanna, 1977) and watermelon (Sedgley, 1981). Cells of the transmitting tissue possess numerous plasmodesmic connections which are involved in the transfer of metabolic signals from the ovary (Figure 3G). Some plasmodesmic connections are also present between the basal region of papillae and cells of the transmitting tissue, indicating their involvement in the symplastic pathway for the transport of metabolites from the transmitting tissue to the papillae (Figure 3F).

ACCUMULATION OF INTRACELLULAR AND EXTRACELLULAR LIPIDS AND ASSOCIATED ENZYMES IN RELATION WITH STIGMA MATURATION During pollen-stigma interaction, lipids are known to play a role in pollen hydration, germination and pollen tube penetration into the style (Wolters-Arts et al., 1998). Lipids prevent evaporation of stigmatic tissue in dry stigma and prevent desiccation of exudates in wet stigma (Shivanna, 2003). Some lipids in the exudates serve as attractants and are of nutritional value for the pollinators (Lord and Webster, 1979). In wet stigmas, lipids are present in the stigmatic exudates, as observed in Phaseolus vulgaris (Lord and Webster, 1979), Nicotiana tabacum (Cresti et al., 1986), Olea europaea (Serrano et al., 2008), while in the dry stigmas, lipids are present as a continuous layer of cuticle beneath the pellicle, as demonstrated in Zephyranthus sp. (Ghosh and Shivanna, 1984). In semi-dry stigmas, such as those in Helianthus annuus and Senecio squalidus, a small amount of lipid-rich extracellular secretion is evident in the crevices of the papillae, and cuticle is not continuous (Shakya and Bhatla, 2010; Allen et al., 2010; Sharma, 2012). Lipid content in sunflower stigma increases with the attainment of stigma receptivity, as observed in wet stigmas of Forsythia intermedia and Nicotiana tabacum (Matsuzaki et al., 1985; Dumas, 1977). Triacylglycerol (TAGs) content decreases from bud to staminate stage in sunflower and shows an increase at the pistillate stage. It is probable that TAGs at the bud stage are degraded during the growth of the stigma and are synthesized during the pistillate stage of stigma development. In addition, terpenes have also been detected at the pistillate stage of stigma development. Cis-unsaturated fatty acids have been reported to be essential components of stigma secretions among wet stigmas and are required for restoring stigma fertility (Wolters-Arts et al., 2002). Fatty acid composition in Helianthus annuus shows the abundance of some saturated and unsaturated fatty acids, as has also been observed in Nicotiana tabacum and Forstythia intermedia (Shakya and Bhatla, 2010; Matsuzaki et al., 1983; Dumas, 1977). Palmitic acid (16:0) is the major saturated fatty acid at all the stages of stigma development and its content increases as

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the stigma attains maturity (Figure 4A; Shakya and Bhatla, 2010; Sharma, 2012). Stearic acid (18:0) content, however, decreases with stigma maturation. Linoleic acid is the major unsaturated fatty acid in the lipids of sunflower stigma (Figure 4B). Its content decreases on maturity, indicating its role in stigma maturation. In contrast, linolenic acid increases as the stigma attains maturity. Linolenic acid is required as a substrate for octadecanoid pathway which results in the production of signalling molecules, such as jasmonic acid (McConn and Browse, 1996).

Figure 4. Relative content of major saturated (A) and unsaturated (B) free fatty acids from stigma at different maturation stages as resolved by gas liquid chromatography. Each value is a mean of three independent values (±standard error). Zymographic detection of fatty acyl esterase isoforms in different fractions of pollen (C) and developmental stages of stigma (D) following treatment with α-napthyl acetate and fast blue B.

The cuticle on the surface of stigmatic papillae in sunflower has an outer proteinaceous pellicle (Figure 3C). Cytochemical studies have indicated the presence of glycoproteins and some enzymes, predominantly esterases, peroxidases and acid phosphatase, as the major components of the pellicle (Vithanage and Knox, 1977; Shakya and Bhatla, 2010). Nonspecific esterase activity has earlier been implicated in the attainment of stigma receptivity and has a role to play in the metabolism of fatty acids and in host-pathogen interaction (Bilková et al., 2009; Shivanna and Rangaswamy, 1992). Non-specific esterases include carboxylesterase (EC 3.1.1.1), arylesterase (EC 2.1.1.2) and acetyl esterase (EC 3.1.1.6). The activity of these enzymes has been detected in dry stigmas [Linum grandiflorum (Ghosh and

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Shivanna, 1980), Pennisetum americanum (Reger, 1989) and Brassica (Mattsson et al., 1974)], wet stigmas [Impatiens sp. (Kulloli et al., 2010), Moringa oleifera (Bhattacharya and Mandal, 2004) and Nicotiana sylvestris (Kandasamy and Kristen, 1987)] and semi-dry stigmas, as in Helianthus annuus and Senecio squalidus (Shakya and Bhatla, 2010; Hiscock et al., 2002a). Non-specific esterase activity is evident at all stages of stigma development in sunflower and it increases as stigma attains maturity. As in Helianthus annuus, esterase activity has also been detected in the early stages of stigma development in Nicotiana sylvestris (Kandasamy and Kristen, 1987) and Linum grandiflorum (Ghosh and Shivanna, 1980), which can be correlated with the initiation of cell differentiation (Bílková et al., 1999). Qualitative analysis of stigma proteins has revealed a change in the isoform pattern of esterases as stigma approaches maturity (Figure 4D). An increase in the number of isoforms indicates their correlation with stigma maturation (Bhattacharya and Mandal, 2004). Nonspecific esterases present on the stigma surface have been reported to be involved in forming a cutinase complex (Knox et al., 1976; Hiscock et al., 2002b). The cutinase complex interacts with esterases from pollen grain wall and pellicle which allows the breakdown of cuticle, facilitating the penetration of pollen tube into the stigmatic tissue. Removal of stigma surface components by chemical treatment or using an inhibitor of serine esterases, reduces the ability of pollen tubes to penetrate the stigma (Knox et al., 1976). This suggests that serine esterases are associated with cutinase complex formation needed for pollen tube penetration in dry stigmas (Hiscock et al., 2002b). Non-specific esterases have the ability to hydrolyze crossbonds of cell wall polysaccharides and are, therefore, important in the establishment and reorganization of cell wall. Lipase- (Triacylglycerol acyl hydrolase; EC 3.1.1.3) like proteins have also been reported in mature stigma papillae of sunflower (Shakya and Bhatla, 2010), Petunia and Nicotiana (Beisson et al., 2003). Lipases act on the triacylglycerols (TAG) and might be involved in altering the lipidic composition of the stigmatic surface and have been located on the stigmatic papillae and pseudopapillae of the receptive stigma (Shakya and Bhatla, 2010).

REACTIVE OXYGEN SPECIES ACCUMULATION DURING STIGMA DEVELOPMENT AND ASSOCIATED SCAVENGING MECHANISMS Recent reports have suggested that during stigma receptivity, angiosperms exhibit an accumulation of high levels of reactive oxygen species (ROS), principally H2O2 (McInnis et al., 2006a,b; Allen et al., 2011; Losada and Herrero, 2012). ROS are the byproducts of aerobic metabolism which are removed by enzymes and antioxidants. In recent years ROS have been shown to act as signalling molecules in various phases of growth and development, response to environmental stress and during pollen germination and tube growth (McInnis et al., 2006a; Hiscock et al., 2007; Zafra et al., 2010). ROS-mediated signalling is controlled by a fine balance between the production and scavenging of ROS. The main sites of ROS production are located in the plasma membrane, NADPH-oxidases, plastids and peroxisomes (Karuppananpandian et al., 2011; Apel and Hirt, 2004). An increase in ROS accumulation is observed in sunflower stigma accompanying the attainment of stigma receptivity (Figure 5 I; Sharma and Bhatla, 2013). Stigmas are a source of nutrients for the pollen grains and may be prone to microbial attack. It is suggested that ROS play a role in defence mechanisms since

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high levels of ROS have been detected in the floral nectar which never experience microbial attack (Carter and Thornburg, 2004). ROS may directly be toxic to the pathogen or may trigger hypersensitive reaction and programmed cell death at the site of pathogen attack (De Rafael et al., 2001). It has also been demonstrated that ROS is required for the polarized growth of the pollen tube (Potocký et al., 2007). Superoxide dismutases (SOD; EC 1.5.1.1) form the first line of defence against the accumulation of superoxide anion (O2-) and lead to its conversion into H2O2 and dioxygen. SOD activity is associated with redox cycle in the nectar in Nicotiana (Carter and Thonburg, 2004; Carter et al., 2007). The SOD in stigma activity is mainly due to Mn-SOD (mitochondria localized) which may be associated with enhanced nectar secretion at the base of florets during pollen producing stage and also when stigma is receptive (Figure 5 III; Tripathi and Singh, 2008; Sharma and Bhatla, 2013). Coinciding with the increased SOD activity, a cluster of mitochondria is evident at the basal region of papillae cell cytoplasm, showing the increased metabolic activity accompanying stigma elongation and dehiscence of anther (Figure 3E; Sharma, 2012). A slight reduction in SOD activity at the pistillate stage of stigma development leads to a reduction in O2- form of ROS and an increase in H2O2 in the stigmatic papillae. Peroxidases (POD; EC 1.11.7) are heme-containing glycoproteins and their activity has been reported both in wet and dry types of stigmas (McInnis et al., 2005). Peroxidase activity in the stigmatic tissue increases as the floret development reaches its maximum at receptivity. In Arabidopsis thaliana, Petunia hybrida and Senecio squalidus, POD activity increases as stigma attain receptivity (Dafni and Maués, 1998; McInnis et al., 2006b). Younger stages of developing stigma show reduced POD activity in Peduclaris canadensis, Clintonia borealis and Helianthus annuus which has been correlated with poor pollen adhesion and germination (Figure 5 IV; Galen and Plowright, 1987; Sharma and Bhatla, 2013). Recent investigations have reported peroxidase isoforms specific to stigma. Expression of stigma-specific peroxidases is developmentally regulated, their activity being maximally observed during stigma receptivity (McInnis et al., 2005). Among the stigmaspecific peroxidases, three isoforms have earlier been identified in Arabidopsis, one in Senecio squalidus, and one in hazelnut (Beltramo et al., 2012). Stigma-specific expression of POD indicates the key role of peroxidases in the loosening of cell wall components of stigma to allow pollen tube growth into the stigma. They might be involved (through H2O2 metabolism) in signalling network, mediating species-specific pollen recognition (McInnis et al., 2005). Some peroxidases are also known to be induced/upregulated in association with hypersensitive response or stress, thereby indicating their role in defense mechanism of stigma (McInnis et al., 2005; Beltramo et al., 2012). Nitric oxide (NO) is a gaseous signalling molecule known to be involved in different plant processes related to growth and development and in responses to stress. Recent investigations have also revealed that NO is likely to be involved in plant reproductive processes (Hiscock et al., 2007; Seligman et al., 2008; Yadav et al., 2013). In sunflower, NO accumulation increases as stigma attains maturity (Figure 5 II; Sharma and Bhatla, 2013). Similar increase in NO has been reported in the developing stigma of olive and Arabidopsis (Zafra et al., 2010; Seligman et al., 2008). NO is important in imparting immunity to the receptive stigma and it also increases thermotolerance by activating ROS scavenging enzymes, thereby playing a role in ROS-mediated signalling processes (Piterková et al., 2013; Sharma and Bhatla, 2013). Upon pollination, a crosstalk between pollen-localized NO and stigmatic ROS has been proposed which may have a role in pollen recognition and signalling

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between the stigmatic papillae and pollen grains (Hiscock et al., 2007; Sharma and Bhatla, 2013).

Figure 5. Accumulation of reactive oxygen species (ROS) and expression of associated scavenging enzymes during stigma development. I: Localization of ROS on the surface of developing stigma after treatment with fluorescent probe-dichlorodihydrofluorescein diacetate (DCFH-DA). Magnification: 100X. Inset shows intense ROS accumulation in the papillae. II: Localization of nitric oxide (NO) on surface of developing stigma after treatment with MNIP-Cu {Copper derivative of (4-methoxy-2-(1Hnapthol [2,3-d] imidazol-2-yl) phenol)}. Magnification: 200X. III: Zymographic detection of superoxide dismutase (SOD) isoforms in developing stigma, after treatment with nitro blue tetrazolium. IV: Zymographic detection of peroxidase (POD) isoforms in developing stigma following benzidine treatment.

LIPIDS IN POLLEN COAT AND THEIR ROLE IN FOOT FORMATION Mature pollen grains are released in a highly desiccated condition and are metabolically inactive. Pollen grains are suboblate, echinate, tectate and tricolporate (Gotelli et al., 2008). The innermost layer of pollen wall (intine) is thin as compared to the outer layer (exine)

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which is spinulate and has pollen coat substances embedded on it. The spines of pollen are conical and have spinular microperforations, indicating their entomophilous nature (Figure 3H; Harry et al., 1978; Shakya, 2005; Coutinho and Dinis, 2007). The exine is differentiated into ektexine and endexine, both of which are separated by a space designated as cavus 2. Ektexine is formed of spinules, tectum, internal foramina (openings), columella, large internal spaces (cavus 1) and foot layer (Figure 3I). Proteins originating from tapetal cytoplasm attach to the cavae and internal foramina of exine, which are known to act as allergens and recognition substances for interspecific compatibility (Horner and Pearson, 1978). The exine pattern is of ‗Helianthoid type‘, referring to abundant internal foramina in the columella and tectum, equal length of columella having basally fused regions and presence of cavae and a thin foot layer (Skvarla and Turner, 1966). Connected basal region of columella, internal formina in columella and enlarged cavae in sunflower (as in other members of Asteraceae, such as Pallenis maritiama, Jasonia tuberose and Astericus aquaticus) allows easy communication between pollen surface and cavae, facilitating the exchange of water and physiologically active substances between them (Coutinho and Dinis, 2007). Pollen coat is rich in lipids which originate from the tapetal cytoplasm (Horner and Pearson, 1978). Pollen capture is exine-dependent and at a later stage it involves the formation of ―attachment foot‖ at the point of its contact with the stigmatic papillae. Pollen coat contains essential components required for adhesion and cell to cell interaction between the stigmatic cells and pollen. The pollen coat material flows out from between the columellae of exine to form an adhesive foot at the surface of the papillae (Wheeler et al., 2001). Lipidic constituents in the pollen grains of sunflower belong to two different domains- the external tryphine (pollen coat) and the internal cytoplasmic (internal pollen). The lipidic content of pollen coat is more than that of the internal pollen (Shakya and Bhatla, 2010). Among the total neutral lipids, neutral esters (wax esters) and free fatty acids are the major components of the two pollen fractions. Gas chromatographic profile of free fatty acids has revealed an abundance of saturated and unsaturated free fatty acids in the internal pollen and the pollen coat. As in stigma, the major saturated fatty acids are palmitic (16:0) and stearic (18:0) acids both in pollen coat and internal pollen, thus indicating a functional similarity in the lipidic constituents of stigmatic exudates and pollen (Piffanelli et al., 1997; Shakya and Bhatla, 2010). Lignoceric acid (24:0), which is expressed more in the pollen coat than in the internal pollen fraction, is specifically expressed only in pollen grains, pointing to its role in the involvement of long fatty acids in the signalling mechanism for hydration of pollen on the stigma surface. Among the unsaturated fatty acids, oleic (18:1), linoleic (18:2), linolenic (18:3) and cis–eicosenoic (20:1) acids have been detected in the pollen grains. Linolenic acid (18:3) is the major unsaturated fatty acid in the pollen coat and internal pollen in sunflower, and Brassica napus (Evans et al., 1987; Shakya and Bhatla, 2010). Cis-eicosenoic acid (20:1) is the major component of intact pollen grains in sunflower (Schulz et al., 2000). Pollen coat in sunflower exhibits lipase activity, as has also been reported in Arabidopsis thaliana (Mayfield et al., 2001; Shakya and Bhatla, 2010). Lipid profile of the pollen fractions shows an absence of triacylglycerides in the pollen coat fraction. It is likely that lipases in pollen coat are activated when pollen coat makes a contact with the stigmatic tissue. Pollen coat lipases may be involved in the degradation of the lipids, such as cuticle present in the stigmatic tissue, and they may also participate in various signalling activities (Murphy, 2006). Several esterase isoforms have been detected in the two fractions of pollen (internal pollen and pollen coat). Four and three isoforms have been detected in the pollen coat and

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internal pollen, respectively (Figure 4C; Shakya and Bhatla, 2010). Esterase activity has earlier been localized in pollen intine suggesting its role in cuticle degradation during the entry of pollen tube (Vithanage and Knox, 1979). It has been proposed that esterases from the pollen and stigma cause a species-specific recognition event resulting in the formation of a ‗cutinase complex‘ which digests the cuticle for the successful penetration of pollen tube into the stigma (Knox et al., 1976; Hiscock and Allen, 2008). Only one peroxidase isoforms is present in the intact pollen of sunflower (Shakya, 2008). Three isoforms have been detected in the internal pollen whereas pollen coat does not show peroxidase activity. Pollen peroxidases are known to degrade phenolic compounds (such as chlorogenic acid, caffeic acid and cinnamic acid) present in the stigmatic exudates (Bredemeijer, 1984, 1982). Phenolics present on the stigmatic exudates or stigma surface seem to be involved in the stimulation or inhibition of IAA oxidase activity which influences growth activity in stigma (Shakya, 2008; Bredemeijer and Blaas, 1975).

OTHER BIOCHEMICAL FEATURES OF POLLEN GRAINS AND RECEPTIVE STIGMA Glycoproteins believed to have important role/s in pollen-stigma interaction, are known to be present in the pollen grains (Suraez-Carvera et al., 2005; Kimura et al., 2002; Aelst and Went, 1992; Shakya, 2008). In the pollen grains of sunflower, four glycoproteins have been detected. As in the pollen grain of Elaeis guineensis, an isoforms of 31kDa has been detected in sunflower as well. The other three glycoproteins correspond to the presence of xylanases which help in the hydrolysis of xylan in the cell wall of stigma (Shakya, 2008). Some glycoproteins in stigma are known to be correlated with the expression of S gene proteins, namely SLG (S locus glycoprotein) and SLR-1 (S locus related glycoprotein-1; Luu et al., 1997b). SLG is a polymorphic protein known to have a role in self-incompatibility and it is secreted into the cell wall of the stigmatic papillae (Kandasamy et al., 1989; Umbach et al., 1990; Doughty et al., 1993). The cytosolic fraction of receptive stigma of sunflower contains a single glycoprotein of 31 kDa. The expression of SLR1 gene is reduced at the later stages of pollen-stigma interaction, showing its involvement in pollen-stigma cross-linking (Luu et al., 1997b). These glycoproteins on the papillae surface interact with pollen coat proteins (PCP) and facilitate in the process of adhesion. SLG has been demonstrated to bind to PCP-A1 and about ten PCP-like proteins, pointing towards the involvement of SLG in many other processes in addition to pollen-stigma adhesion (Swanson et al., 2004). Various enzymes, including proteases, are required for the proteolytic digestion of proteinaceous pellicle during the initial stages of pollen-stigma interaction (Luu et al., 1997a; Graff de et al., 2001; Swanson et al., 2004). Protease activity may also result in the damage of proteins and enzymes on the stigma surface, which leads to the activation of pollen cutinase necessary for the degradation of cuticle (Radlowski, 2005). A protease 54kDa protease has been reported in the cytosolic fraction of sunflower at the receptive stage, in contrast with Cynara cardunculus where two proteases have been reported in the storage vacuoles of stigmatic papillae and transmitting tissue of mature stigma (Verissimo et al., 1996; Shakya, 2008). Calcium is an important constituent of in vitro germinating pollen and serves as a chemoattractant for guiding the growth of pollen tube. Membrane-bound calcium appears to be generally

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distributed in the papillae at all the stages of development. Young buds, however, show lesser accumulation of bound calcium in the stigmatic papillae. At the staminate stage, calcium content increases, reaching a maximum at the pistillate stage of stigma development. Bound calcium has been detected in the pellicle and upper region corresponding to the cytoplasmic organelles in the papillae at the pistillate stage of stigma development. The tip region of stigma also shows an increase in bound calcium content. Investigations using 45Ca2+ have revealed calcium uptake by the germinating pollen from the stigma tissue (Bednarska and Butowt, 1995). De-esterified pectins in the apoplast are capable of binding with calcium ions. Upon pollination, bound calcium is liberated due to the enzymatic lysis of de-esterified pectin, leading to an increase in the levels of free calcium (Bednarska, 1989; Lenartowska et al., 2001). In the growing pollen tubes, callose synthesis is also a calcium-dependent process (Bednarska, 1989).

THE PROCESS OF POLLEN ADHESION, HYDRATION AND GERMINATION ON THE RECEPTIVE STIGMA Pollination involves the transfer of viable pollen onto the receptive stigmatic surface. Adhesion is initiated due to non-specific van der Waal forces between the rough surface of stigma bearing papillae and the spikes of pollen grains (Ferrari et al., 1985; Thio et al., 2009). The process of pollen adhesion is rapid, maximal at the receptive stage of stigma and initiates in a similar manner both in self- and cross-pollinated conditions. As has also been reported in Brassica oleracea, the papillae undergo physiological changes with the attainment of stigma maturity thereby affecting their ability to interact with pollen grains (Heizmann et al., 2000; Sharma, 2012). The process of adhesion involves the interaction of proteins present in the pellicle (arabinogalactan proteins) and the pollen wall (Hiscock and Allen, 2008; Losada and Herrero, 2012). As in Brassica sp., the initial stages of adhesion are not dependent on Sterlity locus (S). Therefore, recognition or rejection of compatible or incompatible pollen does not occur at this stage. At a later stage, however, with the involvement of S locus glycoproteins, adhesion between incompatible pollen grains does not increase (Heizmann et al., 2000). The pollen coat and the stigmatic pellicle of the papillae have an important role in the process of adhesion. Upon removal of the pollen coat, a reduction in the degree of adhesion between the internal pollen and stigmatic papillae is evident. Adhesion of few decoated pollen and stigmatic papillae has, however, revealed that pollen coat proteins and carbohydrate-based molecules associated with exine are involved in the process of adhesion (Zinkl et al., 1999; Sage et al., 2009). The importance of pellicle in the process of adhesion is further evident by the treatment of pellicle with acetone which leads to a significant decrease in pollen adhesion suggesting that glycolipids and lipids located at the base of the papillae play a role in the process of adhesion. The lipids of the pollen coat and cuticle and extracellular lipids in the basal region of papillae are likely to form hydrophobic bonds, leading to adhesion between the pollen grains and the stigmatic papillae. After adhering to the stigmatic papillae, the pollen grains hydrate. The exact mechanism by which water, nutrients and other essential molecules are taken up by the pollen grains from the stigmatic papillae, is not yet fully understood (Frion et al., 2012). Various proteins from pollen coat, including glycine-rich proteins, calcium-binding proteins, lipases and cutinases aid in the process of hydration by

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causing the breakdown of lipidic constituents at the pollen-stigma interface (Mayfield and Preuss, 2000; Hiscock et al., 2002b; Updegraff et al., 2009; Shakya and Bhatla, 2010). Pollen grains hydrate leading to an increase in their volume, both in self- and cross-pollinated conditions (Vithanage and Knox, 1977). After making a contact with stigma, the pollen coat material flows out from the columella of the exine to the surface of papillae (Ellemen et al., 1992; Allen et al., 2011). This leads to the formation of ‗attachment foot‘ where an interaction between pollen and stigma-derived biomolecules takes place. Lipids create conditions and govern the movement of water for guidance cue for the development of the pollen tube which emerges from the pore in the middle of colpus area to grow through the attachment foot towards the basal region of the papillae (Ellemen et al., 1992; Hiscocok et al., 2002a). The acceptance or rejection of pollen grains on the stigmatic surface seems to be dependent on the interaction between the pollen and stigma-derived attachment foot (Hiscock, 2000; Sharma, 2012). Some of the incompatible pollen grains are not able to germinate while others pass perpendicularly towards the basal region of the papillae. Some pollen grains grow parallel to the papillate surface of stigma thereby showing some incompatible rejection reactions (Figure 6).

Figure. 6. Transverse section through germinating pollen grain and the associated stigma following cross (A) and self (B) pollination. Magnification: 400X.

In sunflower, cross pollination is favoured due to the presence of self-incompatibility, as also observed in the members of Brassicaceae. Members of both the families (Asteraceae and Brassicaceae) have some similarities in pollen-stigma interaction. The common features include the presence of dry stigma surface (Vithanage and Knox, 1977), three-celled pollen grains (Hiscock and Allen, 2008), release of pollen coat on contact with papillae (Elleman et al., 1992) and arrest of incompatible pollen soon after germination accompanying the deposition of callose on the pollen tube and papillae (Allen et al., 2011; Vithanage and Knox, 1977). Habura (1957) was first to report sporophytic self-incompatibility in sunflower and it was later confirmed by various other plant scientists (Luciano et al., 1965; Asthana, 1973). Recent reports have, however, pointed out flexibility in sporophytic self incompatibility in

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other members of Asteraceae (Hiscock, 2000; Ortiz et al., 2006). As in Senecio squalidus, sunflower shows germination of self-incompatible pollen. Some of the incompatible pollen grains are not able to germinate, while others germinate and grow between the papillae (Vithanage and Knox, 1977). It has been suggested that the degree of self-incompatibility and self-fertility depends on genetic control, environmental factors and the morphology of the inflorescence (Miller and Fick, 1997). Self-incompatibility is compensated in situations when the reproductive opportunity of the pistil is affected. Such cases exhibit pseudo selfcompatibility which involves a delayed acceptance of the otherwise incompatible pollen (Brennan et al., 2011). The modification of self-incompatibility may also be brought about by G locus, which is a second gametophytic ancestral locus, remnant of ancestral gametophytic self incompatibility in Brassicaceae, which permits compatible cross between the individuals with the same S alleles, which are otherwise incompatible (Lewis et al., 1988; Hiscock, 2000; Brennan et al., 2011). Recent molecular analysis has also pointed out that sporophytic selfincompatibility in Asteraceae operates by a mechanism different from the SRK/SCR molecular mechanism operative in Brassicaceae (Hiscock et al., 2003; Tabah et al., 2004; Allen et al., 2011). Further investigations related to seed set and the molecular mechanisms in sunflower are required to understand the physiological nature of self-incompatibility. To sum up, the present work highlights the complex interaction between floral development and environmental factors, which seem to operate through a critical balance of auxin and gibberellic acid distribution at the bud primordium. Subsequently, in order to facilitate an effective pollen and stigma interaction, both the reproductive structure undergo biomolecular changes required for ‗foot‘ formation on the surface of receptive stigma papillae. The knowledge gained so far in this direction highlights the role of reactive oxygen species and associated scavenging enzymes, glycoproteins, calcium and many more biomolecules. Alterations in the availability of these biomolecules correlate with distinct subcellular structural changes both in pollen and stigma. Although a lot is known by now about these structural and biochemical events accompanying floral development in sunflower, further investigations on their physiological, biochemical and genetic aspects will provide further information on the crosstalk mechanisms regulating floret development and pollenstigma interaction.

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Chapter 3

DEVELOPMENT OF FEMALE REPRODUCTIVE STRUCTURES AND APOMIXIS IN SUNFLOWERS Olga N. Voronova* Department of Embryology and Reproductive Biology, Komarov Botanical Institute of RAS, Saint Petersburg, Russia

ABSTRACT The apomixis phenomenon was hardly observed in the genus Helianthus L. under natural conditions. In this study, a series of experiments on interspecific hybridization was carried out on plants of Cytoplasmic Male Sterility lines. A number of anomalies in the development of female reproductive system were uncovered, including an apospory and integumentary embryony. A surprising phenomenon was noted, such as total absence of the embryo sac in some ovules. Lack of the main embryo sac and formation of additional aposporous embryo sacs could be observed in the same ovule at the same time. Investigation of the early stages of ovule development showed that aposporous embryo sacs originated from the same ovule subepidermal cells as a normal embryo sac. Aposporous embryo sac included the same elements, as the main one: the egg cell, the synergids, the central cell with polar nuclei or secondary nucleus, and antipodes. Asynchrony in development of the main and aposporous embryo sacs was noted. Integumentary embryos were observed in unfertilized embryo sac (7-9 day after flowering). These embryos were formed from the group of initial cells in the chalazal region of the integumentary tapetum; their apical poles were turned to the micropyle. The integumentary embryos consisted of 1-2 suspensor cells and 7-8 embryonic cells and by structure corresponded to Asterad-type, as a typical zygotic embryo. Apospory embryo sac and initial cells of integumentary embryo are formed at the fixed stage of ovule development, the first – when female gametophyte forms, the second – when the development of sexual embryo should begin.

Keywords: Helianthus, CMS, apospory, integumentary embryony, interspecific hybridization

*

Department of Embryology and Reproductive Biology, Komarov Botanical Institute, RAS, 2 Professor Popov St., Saint-Petersburg, Russia 197376. tel./ fax: +7(812) 3725441. Email: [email protected].

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LIST OF ABBREVIATIONS ac bc1 bc2 bn cln hc ic in it msc sec

archesporial cell first basal cell second basal cell basal area of nucellus cells of lateral area of nucellus hypostases cells initial cell integument integumentary tapetum megasporocyte subepidermal cells

INTRODUCTION The early embryological studies of cultivated sunflower were carried out by Goldflus (1899) and Nawashin (1900a,b) who described different aspects of formation and development of its reproductive system. Many studies of sunflower reported that the ovule is tenuinucellar, unitegmic, anatropous with integumentary tapetum, Polygonum-type development of embryo sac, and Asterad-type (Senecio-variation) embryo formation (Newcomb, 1973a,b; Toderich, 1988; Yan et al., 1991; Gotelli et al, 2008, 2010). Under natural conditions, only cross-pollination is typical for sunflower and the apomixis phenomenon was hardly observed in genus Helianthus L. (Noyes, 2007). Formation of several aposporous embryo sacs under unfavorable weather conditions, such as low temperature and excessive humidity, is described by Ustinova (1955, 1964, 1970) in sunflower cultivars Zhdanovskiy and Saratovskiy. She suggested that aposporous embryo sacs arise from the integument cells because the nucellus cells are lysed early. Dzyubenko (1959) noted some cases of apospory in ornamental double-flower form of sunflower. She pointed out the presence of a group of cells near the chalazal end of the normal embryo sac that could be initial aposporous embryo sacs. She named these cells the sporogenous cells. Additionally, Toderich (1988) mentioned several examples of the formation of two embryo sacs in the same ovule in different wild sunflowers (H. rigidus, H. petiolaris, H. occidentalis), but she suggested that they originated from two different archesporial cells, which were formed within one ovule. Pleten et al. (2011) reported hemigamy in sunflower lines APS-11. The majority of the earlier studies of sunflower were carried out on its cultivars (Ustinova, 1955, 1964, 1970; Dzyubenko, 1959; Newcomb, 1973a, b; Toderich, 1988; Yan et al., 1991). Cytoplasmic male sterility (CMS) lines are widely used in sunflower breeding because it allows a breeder to avoid the difficult process of manual sterilization of the flower in order to create a genetic variety by hybridization. Unfortunately, the CMS sunflower lines were not studied comprehensively. Previous studies have paid attention mainly to male reproductive structures (Horner, 1977; Simonenko and Karpovich, 1978; Balk and Leaver, 2001) and only in recent years the researchers began to study the female reproductive

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structure of CMS lines using cyto-embryological analysis (Voronova, 2006, 2008a, 2010; Voronova and Gavrilova, 2007; Gotelli et al., 2008). In this study, a series of experiments on interspecific hybridization was carried out on plants of CMS-lines followed by a cyto-embryological research of ovule development. As a result, a number of anomalies in development of female reproductive system were uncovered, including such phenomena as an apospory and integumentary embryony.

METHODS The main material of the study were the sunflower plants of Helianthus annuus cv. Peredovik and lines VIR 114 form A, VIR 116 form A and their fertile analogs VIR 114 form B and VIR 116 form B. Plants were grown in the Kuban experimental station of N.I. Vavilov Research Institute of Plant Industry (VIR) in Krasnodar region (45°12′54.95″N, 40°47′37.12″E). Perdovik plants were cross-pollinated. Whole heads and separated tubular flower of plants of H. annuus cv. Peredovik (standard) were collected and preserved at different stages of flower development, starting from ovule initiation stage (about 1cm heads diameter ) to torpedo embryo stage (over 20 cm heads diameter). Flower heads of CMS plants were protected from foreign pollen by bagging. When the second circle on a head began to flower, they were pollinated with the fresh pollen of wild sunflower species or lines B. The pollen was collected immediately prior to pollination. The following perennial species were used as pollinators: H. ciliaris, H. decapetalis, H. divaricatus, H. gigantheus, H. hirsutus, H. maximiliani, H. mollis, H. nuttalli, H. occidentalis, H. strumosus (all from collections of VIR). Also, the plants of CMS lines form A were pollinated by their fertile analog of the form B. The material for cyto-embryological investigations was preserved beginning from 1 hour and until 9 days after pollination, that is 1, 2, 3, 4, 6, 8, 10, 12, 24, 36, 48 hours after pollination and 3, 4, 5, 6, 7, 8, 9 days after pollination. The whole flower heads or separate tubular flowers taken by pincers from a head were preserved in FAA solution (formalin, acetic acid and 70% ethanol in 7:7:100 ratio). The pieces were then dehydrated in an ethanol series, infiltrated in ethanol-chloroform mixtures and embedded in Histomix®. Permanent preparations for light microscopy were stained with Heidenhain's hematoxylin with alcian blue (for more details see Zhinkina and Voronova, 2000). The method of the total clearedovule by methyl-benzoate was used for accelerated analysis of the material (Crane, 2001; Voronova, 2008b). The sections were observed and photographed using a Zeiss Axioplan 2 Imaging microscope. Cleared ovaries were examined with Nomarscki interference contrast optics.

RESULTS AND DISCUSSION A sunflower ovule is initiated in the base of an ovary, about 7-10 days before flowering. The ovule primordium is formed by periclinal divisions of cell in the subepidermal placental layer (Figure 1a, b, c).

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First stages of ovule development in the sunflower. Figure 1a, b, c – tubular flower at the stage of ovule primordium initiation. Figure 2a, b – the ovule primordium with initial cell in subepidermal layer. Figure 3a, b – the initial cell elongates and starts mitotic division.

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First stages of ovule development in the sunflower (Continued). Figure 4a, b, c – the initial cell divided into archesporial cell and basal cell, integument initiates in epidermal layer of ovule, the ovule starts being curved. Figure 5a, b – the archesporial cell elongates, the integument actively develops, the vascular band initiates in ovule.

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First stages of ovule development in sunflower (Continued). Figure 6a, b, c – tubular flower at the stage of megasporocyte formation, the ovule curves in anatropous position.

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First stages of ovule development in sunflower (Continued).

Figure 7a, b – megasporocyte before meiosis: the first stage of integumentary tapetum formation – two cell in epidermal layer start flatten. Figure 8 – megasporocyte before meiosis, integumentary tapetum is formed around megasporocyte.

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Embryo sac formation and anomalies in ovule development in sunflower. 9 – ovule with chalazal megaspore, three micropilar megaspore degenerated, walls of integumentary tapetum cell are thickened, 10 – mature embryo sac, 11 – mature embryo sac and aposporous embryo sac with egg apparatus, antipodal cells and two polar nuclei, 12 –ovule with a cell complex instead of a embryo sac, 13 – ovary with mature main embryo sac and aposporous embryo sac, 14 – mature embryo sac, integumentary tapetum forms derivates around the embryo sac, 15– ovule without main embryo, 16 – ovule without main embryo sac and with aposporous embryo sac, 17a,b,c – ovule with integumentary embryo in degenerated embryo sac: a,b – neighboring section of one ovule, c - integumentary embryo, 18 – ovule with embryo and endosperm.

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1-4 – CMS line VIR 114 A, 5-8, 18 – cv. Peredovik, 9-17 – CMS line VIR 116 A; 1-13, 17-18 – longitudinal slides were stained with Heidenhain's hematoxylin and alcyan blue, 14-16 – total ovule (phase contrast). ac – archesporial cell, aes – aposporous embryo sac, anc – antipodal cell, an – anther, bc – basal cell, bn – basal area of nucellus, cc – cell complex, ccn – central cell nucleus, ch - chalaza, cln – cell of lateral area of nucellus, dea - degenerating egg apparatus, ec – egg cell, em – embryo, en – endosperm, es – embryo sac, ic - initial cell, ie – integumentary embryo, ii – initial of integument, hc – hypostase cell, in – integument, it – integumentary tapetum, izi – inner zone of integument, mc – micropile, msc – microsporocyte, ms – microspore, ozi – outer zone of integument, ov – ovary, pn – polar nuclei, pt – pistil, sec – subepidermal cell, tc – table-like cell, vb – vascular bands. Scale bars: 1b, 2a, 3a, 4b, 5a, 6b, 7b, 8, 9, 10, 11, 12, 17a,b,c, – 30 mkm, 1a, 4a, 6a – 150 mkm, 13 – 300 mkm.

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Previous studies suggested that usually 1-2 (for wild species up to 3-4) archesporial cells were formed in subepidermal layer of the primordium ovule, which then develops into megasporocyte directly without any mitotic divisions (Ustinova, 1964; Dzyubenko, 1959; Newcomb, 1973a; Toderich, 1988, etc.). In this study, a different order of structures formation were observed. One to two (rarely 3) subepidermal cells (sec) divided mitotically and gave rise to initial (ic) and first basal cells (bc1) located under initial cell in the ovule primordium of cultivated sunflower. That results in couples of cells located around future long axis of an ovule (Figure 2a,b). The ic cell of the central couple divided once again periclinally and formed an archesporial cell (ac) on the top and second basal cell (bc2) on the bottom (Figure 3a,b, 4a,b,c). Both basal cells (bc1+ bc2) became a part of basal area of nucellus (bn). Other pair of cells (analog ic+bc1 cells) located around the archesporial cell formed a lateral area of nucellus (cln) (Figure 4c, 5b, 6c, 8, 9). The archesporial cell represented an apical area of nucellus. Thus, the apical, lateral, and basal areas of nucellus were identified on the early stage of sunflower ovule development. About the division the nucellus onto different areas see Shamrov, 1998, 2008. Therefore, the ovule of a sunflower can be considered not as tenuinucellar, but rather as medionucellar, with sindermalny variation and single-layer subvariation, according to Shamrov's classification (2008). According to Endress‘s classification (2011), it would be an incompletely tenuinucellar ovule, that is without a hypodermal cell layer between the megasporocyte and nucellus apex, but with hypodermal tissue at the nucellus flanks and below megasporocyte. The single integument (in) was developed in epidermal placental layer at the same time as the formation of an archesporial cell (Figure 4b,c, 5a,b). The layer of hypostases cells (hc) developed at the level of initial cells of integument and on top of them, the basal cells (bc1) differentiated. An axial row was formed as ac+bc2+bc1+hc (Figure 4c). The basal cells bc1 increased in size and became strongly vacuolated, with enlarged and poorly stained nucleus. Because two to three such cells were present in an ovule, they formed a peculiar light zone transparent in the center of the ovule. The basal cell located under the archesporial cell was larger than the latter. Archesporial cell (ac) elongated and gave rise to megasporocyte (msc). At the same time the internal epidermal cells of integument located nearby also underwent changes. They started differentiating into integumentary tapetum (it) which was well expressed already at the stage of mononuclear embryo sac and consisted of one layer of dark-stained, 1-2-nuclear, flattened cell of table-like form with thickened walls (Figure 7a,b, 8, 9). Later, the integumentary tapetum became a two to three layers deep and formed derivates around the mature embryo sac (Figure 10, 11, 14), as typical for Asteraceae (Musial et al., 2012, 2013; Bencivenga et al., 2011). During differentiation of archesporial cell to megasporocyte, the cell of basal area of the nucellus underwent one more mitotic division and formed two cells of table-like form (tc1 and tc2). An axial row was formed as msc+tc1+tc2+bc1+hc (Figure 6c). The table-like cells and basal cells together formed the basal area of nucellus. The cells of the basal area of nucellus remained for a long time and formed a pathway from an archesporial cell (or megasporocytes and megaspore) to hypostasis. Formation of rows of table-like cells, the cells of hypostases, in a broad sense, under a megasporocyte is a characteristic for a number of floral plants, in particular for cereals, such as wheat and corn (Batygina, 1974, 1987, Voronova et al., 2003). It was noted that the archesporial cell of the wheat is terminating an

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already existing number of the cells located along a polar axis of nucellus (Batygina, 1974). As one can see, in sunflower the differentiation of archesporial cell begins with the formation of a special cell which will further form a basal area of nucellus. But unlike cereals, where rows of table-like cells remained unchanged up to seed maturation, in sunflower only two table-like cells got formed. Soon these cells got elongated and formed a narrow row together with the underlying cells of basal area of nucellus (Figure 9, 10). Formation of apospory embryo sac took place within the row (Figure 11, 13). Apospory embryo sacs, as a rule, had the improper shape and was formed from the integumentary tapetum behind the main embryo sac, but was not adjoined to it. Initial cells of aposporous embryo sac were positioned near the basal area of nucellus and, probably, originated from the same subepidermal cell (sec) of ovule primordium, like an archesporial cell (Table 1). Thus, the initial cells of aposporous embryo sacs arose from the cells of the nucellus, and not from the integument as proposed by Ustinova (1970). And, probably, they can be related to the cells that Dzyubenko (1959) called sporogenous cells without considering their origin. Cells of lateral area of nucellus (cln = ic+bc1) exist only at early stages of ovule development. Initially located around an archesporial cell, they are forced out downwards during the process of growth (Figure 4c, 5b, 6c, 7a). These cells continued to put pressure on the cells of the lateral area of nucellus and shifted them down towards the chalaza with the development of megasporocyte and then megaspore. The cells of the lateral area of nucellus were compressed, but remained distinct up to the stage of mononuclear embryo sac (Figure 8, 9). The young ovule of sunflower is ortotropuos (Figure 1a,b, 2a, 3a), but one side of the integument got curved because of the more intensive growth (Figure 4a, 5a, 6a,b). The mature ovule of sunflower is anatropous (Figure 13), unitegmic, with integumentary tapetum or endothelium, like in other Asteraceae. Table 1. The sequence of divisions at an early stage of sunflower ovule development

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The ovule showed a zonal differentiation, that is particularly visible in the central part. The cells of the outer zones were elongated with thin cell wall, whereas the cells of the inner zone were disintegrated and had thick cell wall intensively stained with alcian blue (Figure 9, 12, 13, 17a,b, 18). The vascular bundle passed through chalaza and penetrated integument almost reaching the micropyle (Figure 5b, 6c, 7a), like in some other Asteraceae species (Musial et al., 2013). The megasporocyte developed directly from archesporial cell without any mitotical division (Figure 5a,b, 6b,c, 7a,b, 8). The meiotic divisions of megasporocyte produced a linear tetrad of haploid megaspores. Three of them degenerated and Polygonum-type embryo sac was formed from one chalazal megaspore (Figure 9, 10). The embryo sac was completely developed by the time of pollination. The mature embryo sac consisted of the three-cellular egg apparatus, the central cell with big, fused, secondary nucleus, and antipodal cells (Figure 10). Egg apparatus consisted of three pearshaped cells: two synergids and the egg. The egg cell nucleus was rather large, with obvious nucleolus, and positioned in the apical part of the cell. The synergid nuclei were barely distinguishable and positioned in the center of cells. A filiform apparatus with hamiform evagination was found in the basal part of synergids. The bulk of cytoplasm of the central cell was located near the egg apparatus and thin bundles between large vacuoles. The large central cell nucleus was located near the apical end of the egg cell. Antipodes were linear. Antipodal cells were strongly vacuolated, with the large, polyploid nuclei. Usually, there are only two antipodes in the mature embryo sac. Antipodal cell were adjoined to the central cell, usually larger, underlying, and can increase 2-3 times in size. The remains of the third antipode were sometimes observed in the chalazal end of embryo sac. The antipodal complex as a whole took half of the general linear size of the embryo sac. An additional embryo sacs were found in the ovule of the plants pollinated by perennial wild species. They were found at different times after pollination, from two hours to nine days, and in different combination of pollinations (see Table 2). The relative position of additional and main embryo sacs confirmed that the additional embryo sac was aposporous. Besides, the groups of big megaspore-like cells were found around chalazal end of main embryo sacs at earlier stages of development (Voronova, Gavrilova, 2007). These are the cells of the basal area of the nucellus that have originated from the subepidermal cells of the ovule (see Table 1). The main embryo sac in all cases had a normal structure that is characteristic for sunflower. Aposporous embryo sac included the same elements as the main one: the egg cell, the synergids, the central cell with polar nuclei or secondary nucleus, and antipodes. All elements of additional embryo sac were similar to those of the main, but differed in sizes and by the level of differentiation. There were more than three antipodal cells (Figure 11). Aposporous embryo sacs lied below the level of integumentary tapetum. Absence of integumentary tapetum around an additional embryo sac leaded to a variety of shapes and sizes of this embryo sac. Apparently, the integumentary tapetum plays a role of a bearing element and gives definite form to the whole structure of the main embryo sac. The asynchrony in development of the main and additional embryo sac was noted. The additional embryo sacs, as a rule, delayed in development. For example, the main embryo sac could be fully formed while the polar nuclei of additional embryo sac did not fuse yet (Figure 11).

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Table 2. Anomalies in ovule development after pollination of CMS lines by wild perennial sunflower Pollination combination VIR 114 A х H. ciliaris VIR 116 A х H. nuttalli VIR 116 A х H. occidentalis VIR 114 A х H. occidentalis VIR 114 A х H. gigantheus VIR 114 A х H. nuttalli VIR 116 A х H. divaricatus VIR 114 A х H. hirsutus VIR 116 A х H. gigantheus VIR 114 A х VIR 114 B VIR 116 A х VIR 116 B VIR 116 A х H. maximiliani VIR 116 A х H. ciliaris VIR 116 A х H. decapetalis VIR 116 A х H. mollis VIR 116 A х H. strumosus cv. Peredovik (free pollination)

Ovules, in total 35 55 43 85 69 35 35 89 45 119 49 48 46 45 44 40 126

Ovules without any embryo sac, in % 2,9 0 20,9 1,2 0 34,3 14,3 16,9 0 3,4 0 0 0 0 0 0 0

Ovules with apospory embryo sac, in % 51,4 14,5 14,0 11,8 10,0 8,6 5,7 4,5 2,2 1,7 0 0 0 0 0 0 0

A surprising phenomenon, the total absence of embryo sac, was recorded in different ovules of some CMS lines (VIR 114, VIR 116) (Figure 12, 15, 16). Previously, the total absence of the embryo sac was described for tetraploid form of sunflower only (Efremov, 1967; Pustovoit et al., 1976). The authors suggested that the embryo sac underdevelopment was caused by disorders in the preceding meiotic divisions (Pustovoit et al, 1976). Also they noted that in some tetraploid sunflowers the ovule is underdeveloped in general and lacking the embryo sac (Efremov, 1967). In this investigation, all ovule structures were formed normally and phases of their development and relative positioning of elements corresponded to the stages of flower development. Two small and 1-3 large cells with light, strongly vacuolated cytoplasm and small nuclei were found in a cavity surrounding integumentary tapetum, instead of an embryo sac (Figure 12). Possibly, these cells are derivatives of nucellus. Usually, nucellus cells appear squeezed by the developing megasporocyte and degenerate by the time of formation of an embryo sac. It is possible to assume that the transport of nutrients in this zone remains in the absence of embryo sac, and, nucellus cells remain viable for a long time in the absence of effect from outside megasporocyte. Lack of the main embryo sac and formation of additional embryo sacs could be observed at the same time in the same ovule (Figure 16). In case of formation of seeds from similar ovule, the embryo in them could be formed by apomixes only. Integumentary embryos were formed in some ovules of CMS line VIR 116 (Figure 17a,b,c). after parent plants were pollinated by H. occidentalis pollen. Though pollination was carried out, fertilization was not observed even 7-9 days after the pollination. Normally, both, the embryo and endosperm were observed in the ovule at that time. But in some plants of CMS line VIR 116 A, the embryo sac still remained in an 8-shaped form at this stage. It

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was still possible to discern the egg apparatus, the central cell with the secondary nucleus, and the antipodes (Figure 17b). Sometimes the egg cell was significantly increased in size, while synergids were compressed and darkened. The egg nucleus could be larger than the secondary nucleus of the central cell. The cytoplasm of the central cell occupied wall position with a huge vacuole in the center. The secondary nucleus was located near the egg cell. The integumentary tapetum became multilayered and folded, and its cells were transparent with small and poorly visible nuclei. The integumentary tissues near the chalazal pole of the embryo sac were destroyed, the cells were disintegrated and cavities were formed. Integumentary embryos were located in the chalazal region of the integumentary tapetum with their apical poles turned to the micropyle, which is distinct from the sexual embryo (Figure 17a,c). The embryos consisted of 1-2 suspensor cells and 7-8 embryonic cells. Cells were dark-colored and had large nuclei. The lighter nucleoli with dark-colored center were visible in the nucleus (Figure 17a). These embryos corresponded to Asterad-type Senecio – variation (Voronova, 2010). In control pollination of VIR 116 A line with the pollen of the fertile analogue of VIR 116 B and of cv. Peredovik the formed embryos and endosperm at the similar stage of ovule development were observed (Figure 18); later the seeds were fully formed. Unfortunately, the development of fully formed seeds after pollination of the CMS line with pollen of wild sunflower species were not observed. The seeds formed in the head did not contain embryos. It is known that interspecific hybridization in sunflower is possible, but the number of seeds formed is very insignificant, such as 1-10 for a head (Gavrilova and Anisimova, 2003).

CONCLUSION Three types of deviations in the reproductive system of CMS lines were found during this study: absence of an embryo sac; apospory; and integumentary embryony. The last two represent different forms of apomixis. It was noted that apomixes phenomenon is connected with hybridization in different systematic groups (Carman, 1995). CMS lines obtained a sterile cytoplasm from wild sunflower species (Leclerq, 1969) and, therefore, have an interspecific hybridization in their pedigrees (Voronova, Gavrilova, 2007). It is possible, that the reproductive system of CMS lines is less stable than in cultivated species and cultivars. Therefore, the alternative ways of development, such as apospory and integumentary embryony, are rare for the cultivated sunflower but typical for other representatives of the Asteraceae family (Noyes, 2007) and work under stress conditions, for example pollination with an alien pollen, delay or the full absence of the fertilization. Based on the earlier publications (Molchan, 1973; Petrov, 1988; Bencivenga et al, 2011),, it could be assumed that the pollination of cultivated sunflower with the pollen of wild Helianthus species and the absence of normal fertilization can stimulate appearance of different anomalies in the morphogenesis of ovule structures. The disturbance in fertilization can lead to interruption of normal ―cross talk between the sporophyte and megagametophyte‖ (Bencivenga et al, 2011) resulting in the alternative way of development of reproductive system in sunflower lines.

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The interspecific hybridization method was used during the breeding work on sunflower and the pollination process was changed repeatedly. At first, there was a search of self-fertile forms and then, among them, the sterile lines were located. CMS lines were maintained using so called "B lines" or fertile analogs. For production of heterosis industrial hybrids, the process of pollination was changed again and CMS lines were cross-pollinated. The balance existing in reproductive system of stable cross-pollinated cultivars was apparently disrupted in lines and hybrids. Integumentary embryony, as one of apomixes forms, represented indubitable theoretical and practical interest because it implies embryo formation from cells of somatic tissue of ovule. By origin, integumentary embryos represent clones of a parent organism; their formation can result in genetic heterogeneity of seed (Batygina, Vinogradova, 2007). This means that in the progeny of a single plant some of the seeds may have a paternal heredity and part of the maternal. Furthermore, both parents can participate in the endosperm formation. For this reason, possibility of integumentary embryos formation in cultural sunflower perhaps may explain the deviations from expected result of crossings, so-called partial hybridization and lack of splitting in hybrids. These were noted by different authors (Lyashchenko 1940, 1948; Gavrilova et al., 1997; Faure et al., 2002; Gavrilova and Anisimova, 2003). When sunflower was pollinated by wild species under interspecies hybridization, part of progeny tended to resemble a cultivated sunflower, whereas when wild species was pollinated by sunflower, the progeny looked like wild species. It is possible that genes of the endosperm inherited from both parents would be included in the work at the early stages and affected the development of the embryo. It was observed in other plants that the initial cells of integumentary embryos are formed at a certain stage of ovule development, corresponding to the time when the normal embryo should start to develop (Naumova, 1993, 2008). In this study, a similar picture was observed. The alternative (aposporic) embryo sacs were formed if the violation in the development of basic reproductive structures occurred at an earlier stage. The integumentary (somatic) embryos were developed if the violation has occurred at a stage when zygotic embryo must be formed. Apospory embryo sac and integumentary embryo were formed at the determined stage of ovule development, the first during the formation of female gametophyte, and the second during the development of zygotic embryo. Formation of the alternative and additional structures depends on general stage of the ovule development and on the time that passed after pollination.

ACKNOWLEDGMENTS I am grateful to Dr. V. A. Gavrilova (N.I.Vavilov Research Institute of Plant industry) and Dr. V. T. Rozhkova and Dr. T. T. Tolstaja (the Kuban experimental station) for their help in obtaining the experimental material and Dr. T. Lobova for her help in editing the manuscript. This work was performed under the project № 01201255606 and supported by the Russian Foundation for Basic Research (grant № 11-04-01466).

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REFERENCES Balk, J., Leaver, Ch. J., (2001). The PET1-CMS mitochondrial mutation in sunflower is associated with premature programmed cell death and cytochrome c release. Pl. Cell 13, 1803-1818. Batygina, T. B. (1974). Wheat embryology. Leningrad: ―Kolos‖. 206 p. (in Russian). Batygina, T. B. (1987). The grain of cereal. Leningrad: ―Nauka‖. 103 p. (in Russian). Batygina, T. B., Vinogradova, G.Ju. (2007). Phenomenon of polyembryony. Genetic heterogeneity of seeds. Russ. J. Dev. Biol. 38(3), 126–151. Bencivenga, S., Colombo, L., Masiero, S. (2011). Cross talk between the sporophyte and megagametophyte during ovule development. Sex. Plant Reprod. 24, 113-121. Carman, J. G. (1995). Gametophytic angiosperm apomicts and the occurrence of polyspory and polyembryony among their relatives. Apomixis Newsletter 8, 39-53. Crane, C. F. (2001). Classification of apomictic mechanisms. Appendix: methods to clear angiosperm ovules, In: Savidan V., Carman J. G., Dresselhaus T. (Eds.) The flowering of apomixes: from mechanisms to genetic engineering, Mexico, pp. 35-43. Dzyubenko, L. K. (1959). Cytoembryological studies of female generative zone in ovule of the sunflower (Helianthus L.). Ukr. Bot. Zh. 16(3), 8-19. (in Ukraine). Efremov, А. Е. (1967). Morphological and cytoembryological peculiarities of tetraploid sunflower. Genetika 11, 31-36 (in Russian). Endress, P. K. (2011). Angiosperm ovules: diversity, development, evolution. Ann. Bot. 107, 1465-1489. Faure, N., Serieys, H., Cazaux, E., Kaan, F., Berville, A. (2002). Partial hybridization in wide crosses between cultivated sunflower and the perennial Helianthus species H.mollis and H.orgyalis. Ann. Bot. 39, 31-39. Gavrilova, V. A, Anisimova, I. N. (2003). Genetics of cultivated plants. The Sunflower. St.Petersburg, 209 p. (in Russian). Gavrilova, V. A., Tolstaya, T. T., Rozhkova, V. T. (1997). Analysis of interspecific hybrids resulting from crosses between perennial wild Helianthus species and the cultivated sunflower, In: FAO Progress Report 1995–1996, Germany, pp.75–80. Goldflus, M. (1899). Sur la structure et les fonctions de l'assise épithéliale et des antipodes. J. de bot. 13, 87-96. Gotelli, M. M., Galati, B. G., Medan, D. (2008). Embryology of Helianthus annuus (Asteraceae). Ann. Bot. Fennici 45(2), 81-96. Gotelli, M. M., Galati, B. G., & Medan, D. 2010. Structure of the stigma and style in sunflower (Helianthus annuus L.). Biocell 34(3), 133–8. Horner, Y. T., Jr. (1977). A comparative light- and electron-microscopic study of microsporogenesis in male-fertile and cytoplasmic male-sterile sunflower (Helianthus annuus). Amer. J. Bot. 64(6), 745-759. Leclerq, P. (1969). Une sterilite cytoplasmique chez le tournesol. Ann. Amelior. Plant 19, 99– 106. Lyashchenko, I. F. (1940). A case of absence of splitting in sunflower hybrids. Dokl. Akad. Nauk SSSR 27(8), 824–826. (in Russian) Lyashchenko, I. F. (1948). The phenomenon of maternal heredity in the sunflower. Uchenye zapiski Rostov. Gos. Univ. 12(1), 3–26. (in Russian).

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Molchan, I. M. (1973). Genetic and physiological role of pollen in apomictic plants, In: Problems of apomixis in plants and animals, Novosibirsk, pp. 220–228. (in Russian). Musial, K., Koscinska-Pajak, M., Sliwinska E., Joachimiak, A. J. (2012) Developmental events in ovule of the ornamental plant Rudbeckia bicolor Nutt. Flora. 207, 3-9. Musial, K., Plachno, B. J., Swiatek, P., Marciniuk, J. (2013) Anatomy of ovary and ovule in dandelions (Taraxacum, Asteraceae). Protoplasma. 250(3), 715-722. Naumova, T. N. (1993). Apomixis in Angiospers. In: Nucellar and integumentary embryony, CRC Press, Boca Ration, pp. 1-144. Naumova, T. N. (2008). Apomixis and Amphimixis in Flowering Plants, Tsitol. Genet. 42(3), 51-63 (in Russian). Navashin, S. G. (1900a). On fertilization in the Asteraceae and Orchids. Proc. Imperial Acad. Science 13(3), 335-340 (in Russian). Navashin, S. (1900b). Ueber die Befruchtungsvorgange bei einigen Dicotyledonen. Berichte der deutschen botanischen Gesellschaft 18(5), 244-250. Newcomb, W. (1973a).The development of the embryo sac of sunflower Helianthus annuus before fertilization. Can. J. Bot. 51, 863-878. Newcomb, W. (1973b).The development of the embryo sac of sunflower Helianthus annuus after fertilization. Can. J. Bot. 51, 879-890. Noyes, R. D. (2007). Apomixis in the Asteraceae: Diamonds in the Rough. Functional Plant Science and Biotechnology. 1(2), 207-222. Petrov D. F. (1988 ). Apomixis in nature and experiment, Novosibirsk, 211 p. (in Russian). Pleten S., Tarabrina A., Voronova O., Pershin A., Batygina T. (2001) Is there hemigamy in Helianthus annuus? Abstracts Xth International Conference on Plant Embryology (5-8 September, 2001, Nitra, Slovac Republic), p. 50. Pustovoit, GV, Fedorenko TS, Prokopenko AI (1976). Morphological characteristics of female generative organs of tetraploid sunflower. Bulletin VNIIMK 3, 13-16. Shamrov, I. I. (1998). Ovule of flowering plants: structure, functions, origin, Moscow. 350p. (in Russian). Shamrov, I. I. (2008). Ovule classification in flowering plants – new approach and concepts, Bot. Jahrb. Syst. 120(3), 377-407. Simonenko, V. K., Karpovich, E. V. (1978). Cytological demonstration different types of male sterility in sunflower, Nauchno-tehnicheskij bull. Vsesojuz. Gen. Inst. 31, 32-38 (in Russian). Toderich, K. N. (1988). Sunflower (Helianthus annuus, H. rigidus, etc.) embryology. PhD thesis, Leningrad, 256 p. (in Russian). Ustinova, E. N. (1955). The apospory phenomenon in the sunflower. Dokl. Akad. Nauk SSSR 100(6), 1163-1166 (in Russian). Ustinova, E. N. (1964). The variability of female gametophyte in the sunflower (Helianthus annuus L.). Byul. MOIP. Otd. Biol. 69(4), 111-117 (in Russian). Ustinova, E. N. (1970). Apomixis in the Sunflower, In: Apomixis and Breeding, Moscow, pp. 110-116 (in Russian). Voronova, O., Abnormalities in the development of generative organs in some cms sunflower lines, in: Conferinţa Ştiinţifică internaţională „Invăţămăntul superior şi cercetarea – piloni ai societăţii bazate pe cunoaştere” dedicată jubileului de 60 ani ai Universităţii de Stat din Moldova (28 September 2006, Chişinău, Moldova). Chişinău: CEP USM, 2006. P. 335-336.

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Voronova, O. N. (2008a). Development features of a generative system in some CMS-lines of sunflower, Gen. Res. Rast. 6: 76-81. (in Russian) Voronova, O. N. (2008b). Rapid analysis by clarification and its use in embryology. Bot. Zh. 93(10), 1620-1625 (in Russian). Voronova, O. N. (2010). Integumentary embryony in CMS sunflower line. Russ. J. Dev. Biol. 41(6), 394-399]. Voronova, O. N., Gavrilova, V. A. (2007). Apospory in the sunflower. Bot. Zh. 92(10), 15351544. (in Russian). Voronova, O. N., Shamrov, I. I., Batygina, T. B (2003). Ovule morphogenesis in normal and mutant Zea mays. Acta Biol. Cracov. Ser. Bot., 45(1), 155-160. Yan, H., Yang, H.-Y., Jensen, W. A. (1991). Ultrastructure of the developing embryo sac of sunflower (Helianthus annuus) before and after fertilization. Can. J. Bot. 69(1), 191-202. Zhinkina, N. A., Voronova, O. N. (2000). On staining technique of embryological slides. Bot. Zh. 85, 168-171. (in Russian).

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In: Sunflowers Editor: Juan Ignacio Arribas

ISBN: 978-1-63117-347-9 © 2014 Nova Science Publishers, Inc.

Chapter 4

GENETICS AND GENOMICS APPLIED TO SUNFLOWER BREEDING Carla Filippi1,2, Jeremías Zubrzycki1,2, Verónica Lía1,2,3, Ruth A. Heinz1,2,3, Norma B. Paniego1,2 and H. Esteban Hopp*1,3 1

Instituto de Biotecnología, Centro de Investigaciones Veterinarias y Agronómicas, Instituto Nacional de Tecnología Agropecuaria (INTA Castelar), Repetto y Los Reseros, Hurlingham, Provincia de Buenos Aires, Argentina 2 Consejo Nacional de Investigaciones Científicas y Técnicas, Ciudad Autónoma de Buenos Aires, Argentina 3 Facultad de Ciencias Exactas y Naturales, Universidad de Buenos Aires, Ciudad Autónoma de Buenos Aires, Argentina

ABSTRACT Since sunflower domestication by pre-hispanic American cultures at least 3000 BC, the use of empiric and scientifically based genetics led to an amazing genetic diversification of the crop going from sophisticated nutraceutical applications up to ornamental purposes, including the traditional confectionary and oilseed production. Commercial sunflower breeding based on genetics started in the first half of the twentieth century and genomics at its endings, with breeding efforts being directed towards the most economically important traits such as increasing seed and oil yield, improving quality traits and conferring resistance or tolerance to biotic and abiotic stresses. In the last few years, advancements in genotyping and sequencing technologies allowed the development of increasingly dense genetic and physical maps, enabling the development of new breeding strategies based on molecular markers, like QTL mapping, association mapping and genomic selection. The need to increase efficiency and precision has motivated the application of marker assisted selection (MAS) in sunflower breeding programs. This chapter will review the different genomic breeding approaches that are currently used to improve sunflower tolerance to biotic and abiotic stresses, increase oil quality and enhance agronomic yield associated traits in order to reduce the gap between potential and actual sunflower production in the present cultivated sunflower area and *

Corresponding author: Professor H. Esteban Hopp; E-mail: [email protected]

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Carla Filippi, Jeremías Zubrzycki, Verónica Lía et al. under global weather changing conditions that negatively impact on it. An overview of the state of the art on sunflower genomics is presented and the potential of high throughput sequencing and genotyping technologies for crop breeding is discussed.

Keywords: Sunflower, marker assisted selection, QTL mapping, association mapping, cytogenetyc mapping, linkage mapping

INTRODUCTION The Compositae (Asteraceae) is the largest plant family on earth, with over 24000 described species, representing almost 10% of all flowering plant species (Stevens 2010). Compositae species include economically important crops, rare and beautiful wildflowers, invasive weeds and several species harboring common allergens and valuable medical molecules (Dempewolf et al., 2008). They are referred to as Composites because what looks like a single large flower is actually a composite of many, maybe thousands tiny flowers. It includes sunflowers, lettuce, artichokes, dandelions, thistles, daisies, ragweed, goldenrod and chicory (Kane et al., 2011). The genus Helianthus contains about 50 species of annual and perennial herbs (Heiser and Schilling 1981) native to America. It includes diploids, tetraploids and hexaploids, all with the basic chromosome number of 17 (Rieseberg 1991). Asteraceae and its related families Goodeniaceae and Calyceraceae do not have an extended fossil record. In 2010, Barreda et al. described a fossil capitulum and pollen grains from the Eocene that were found in Patagonia, Southern Argentina. This finding evidenced that the family evolved from an ancestor originated putatively in South America about 50 million years ago (Barreda et al., 2010). Sunflower (Helianthus annuus L. var. macrocarpus) was originally domesticated in the east-central part of North America circa 3000-4000 years ago (Crites 1993; Harter et al., 2004; Smith and Yarnell 2009; Bowers et al., 2012) by pre-Columbian civilizations that used it as a source of edible seeds and for other applications (as a source of natural dyes and for ceremonial purposes) (Mandel et al., 2013). Since then, sunflower has been grown with many purposes: as oil crop (its main use), for beauty (ornamental sunflower) and for direct consumption of the seeds (confectionary sunflower). Furthermore, there is an increasing interest in the use of sunflower proteins in human nutrition. The content of protein remaining in cakes and extraction residues after seed oil extraction, accounts for 30 to 50% (Dorrell and Vick 1997). The properties of sunflower proteins are comparable with those of soy and other leguminous proteins (González-Pérez et al., 2005; Zilic et al., 2010). Regarding its main use, sunflower is currently the world‘s fourth largest source of vegetable oil (http://www.fas.usda.gov/). Sunflower oil is considered premium due to its high unsaturated fatty acid composition and low content of linolenic acid. Nowadays, it is cultivated on over 23 million hectares worldwide (http://www.fao.org/), with an annual production of 32 million metric tons, mainly concentrated in the Russian Federation, Ukraine, India, and Argentina (Bowers et al., 2012). Conventional breeding has been successful in raising sunflower yield potential and its stability, as well as in controlling resistance to some fungal diseases, pests and parasitic

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weeds (Sala et al., 2012). The advent and development of molecular markers and genetic maps have facilitated understanding the genetic basis of different agronomic traits. In this chapter, we present and discuss the different genetic and genomic breeding approaches that are currently used to improve sunflower yield and its tolerance to diseases, focusing on the study of pathogen resistance responses and reviewing the state of the art of sunflower genomics.

1. SUNFLOWER BREEDING Commercial sunflower breeding started in most of the producing countries (Eastern and Western Europe, North and South America) between 1920 and 1950 by phenotypic selection, a method of selecting desirable plants from a population on the basis of phenotypic traits. The introduction of heterosis, first described in 1966 (Leclercq 1966), the incorporation of cytoplasmic male sterility after interspecific crossing with H. petiolaris Nutt (Leclercq 1969), and the development of fertility restorer lines by Kinman in 1970 (Miller and Fick 1997), allowed the development of sunflower hybrids. This process is based on a single cytoplasmic male sterility (CMS) source, the PET1. The CMS was associated with the expression of a 16 kDa protein encoded by orfH522 in the PET1 cytoplasm, which is anther-specific reduced in fertility restored hybrids (Moneger et al., 1994). This protein is co-transcribed with the atpA mitochondrial gene in the male sterile lines. The first sunflower hybrids were produced in 1972 and reached 80% of production in five years (Miller and Fick 1997), due to their higher yield and quality potential, high homogeneity, maturing time synchronicity and better adaptation to cultural applications. Sunflower breeding efforts were directed towards the most economically important traits, including: increased seed and oil yield (number of seeds per plant, test weight, 1000-seed weight, low husk content and high oil concentration in the seed), increased harvest index (plant height, head size and shape, angle of the head, leaf area and leaf canopy, early maturation, short stem and uniform height), improved quality traits (oil quality, protein concentration and composition), and resistance to biotic and abiotic stresses (Škorić 1992). Despite the optimism for continued improvement by conventional breeding, new technologies are needed to significantly increase efficiency and precision, and to save time, resources and efforts. Agriculturally important traits, such as yield, quality and some forms of disease resistance are controlled by many genes and known as quantitative traits. The regions within genomes that contain those genes associated with a particular quantitative trait are known as quantitative trait loci (QTL). Owing to genetic linkage, DNA markers can be used to detect the presence of allelic variation for major genes or QTL underlying the traits of interest. The first molecular markers described in sunflower were restriction fragment length polymorphism (RFLP) (Berry et al., 1995, 1996, 1997; Gentzbittel et al., 1995, 1999; Jan et al., 1998), random amplified polymorphic DNA (RAPD) (Rieseberg et al., 1993; Rieseberg 1998) and amplified fragment length polymorphisms (AFLPs) (Peerbolte and Paleman 1998; Flores Berrios et al., 2000; Gedil et al., 2001; Al-Chaarani et al., 2004). However, RFLPs are technically laborious for routine use as molecular markers and while RAPD and AFLP markers have many advantages, they are mostly dominant, abundant but often non-specific

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and not very useful for comparison of a genome-wide synteny of molecular markers for cross referencing genetic linkage maps (Paniego et al., 2007). Nowadays, the most popular molecular markers are microsatellite markers (also called single sequence repeats, SSR) and single nucleotide polymorphisms (SNP). While SSR markers are multiallelic, SNPs are generally assumed as biallelic markers. However, this disadvantage is compensated by the relative abundance of SNPs (Oraguzie et al., 2007) and emerging high throughput profiling technologies. Marker-assisted selection (MAS) is a method whereby a phenotype is selected using molecular markers linked to the trait genetic determinants. The advantages of MAS include: time saving by substitution of field trials with molecular tests; selection carried out at seedling stage; the possibility of combining multiple gene selection; avoid the transfer of undesirable genes by background selection; selection of traits with low heritability; selection of single plants (Collard et al., 2005). One of the most important tools in MAS strategy is having a high-resolution map, composed of polymorphic molecular markers covering all chromosomes, in order to identify markers flanking those QTL that control traits of interest. To avoid losing the selected trait due to recombination events between the marker locus and the QTL, they need to be in strong linkage disequilibrium (at least 0.05). (Du Plessis et al., 2012).

The African bollworm, H. armigera, is an important pest of many crops in many parts of the world (Zalucki et al., 1986; Sharma, 2001). It is also regularly present during the reproductive stage of cultivated sunflower in Africa. Its attractiveness to sunflower is demonstrated by its use as a trap crop in and around organic cotton fields in Tanzania (Cherry et al. 2003). Larvae occur from the budding stage onwards (Von Maltitz, 1993). Levels of infestation vary between localities and seasons, sporadically reaching epidemic proportions. First and second instar larvae are mainly found on sunflower buds, with a feeding preference for involucral bracts. Later instar larvae feed more extensively than younger larvae, consequently doing more damage. One mature larvae feeding inside a bud may completely destroy the immature florets. However, plants infested at the budding stage escape serious feeding damage as the majority of larvae mature during anthesis (Von Maltitz, 1993; Du Plessis, 1997). Larvae of all instars also burrow under and between bracts into the receptacles (Von Maltiz, 1993), and are, therefore often not exposed to insecticides. Insecticides for H. armigera control on sunflower in South Africa are applied aerially and the question often arises whether effective insecticide application is limited by the downward inclination of sunflower heads and by the fact that the heads are often turned in the opposite direction to aerial application. Du Plessis (1997) found aerial application of insecticides for bollworm

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control to be effective during the budding as well as the anthesis and pollination stages when later instars, which are more damaging, occurred on heads. Simulated bollworm damage was done to florets (R-5.1stage) and young achenes in the milky stage (R-6.0) of sunflower in a field trial in South Africa (Du Plessis, 1997). R-5.1 is the reproductive stage at which 10 % of the head area (disk flowers) has completed or is in flower. R-6.0 is the reproductive stage in which anthesis and pollination are completed and the ray flowers are wilting (Schneiter and Miller, 1981). The mean mass of randomly chosen achenes in these heads increased at increased levels of damage (Figure 2). Compensation for lost florets and achenes, therefore, occurred over the entire head. Compared with undamaged heads, the mean mass of achenes was significantly higher at levels of 15, 20 and 30 % damage at both reproductive stages (Figure 2). The total mass of fertile achenes per head did not differ between damaged and undamaged heads, up to a 30 % level of damage (although lower at 5 and 30 % damage levels) (Figure 3) for both reproductive stages evaluated. It showed that sunflower has the ability to compensate for damage (Du Plessis, 1997). Sunflower variety and climatic conditions may provide results which differ from those found by Du Plessis (1997).

Figure 2. Mean mass of 15 achenes surrounding simulated bollworm damage area(s) and chosen at random in sunflower heads at different levels of damage done to florets in the R-5.1 reproductive stage and to young achenes in the milky stage (R-6.0). │= LSDT (P = 0.05). (From: Du Plessis, 1997). (Source: African Crop Science Journal)

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Figure 3. Mean mass of fertile achenes per sunflower head at different levels of simulated bollworm damage (n=24) done to florets in the R-5.1reproductive stage and to young achenes in the milky stage (R-6.0). │= LSDT (P = 0.05). (From: Du Plessis, 1997). (Source: African Crop Science Journal)

Based on the results of this study by Du Plessis (1997), it is proposed that insecticides be applied when at least 20% damage per head occurs, in order to promote timely control measures. Taking into account that the preferential feeding sites of H. armigera are not the achenes, as well as the ability of plants to compensate for damage to florets and achenes, a significant number of larvae can, however, be tolerated without any significant effect on yield. Actual damage is therefore the only criterion which can be used in the determination of economic injury levels for control of African bollworm on sunflower (Du Plessis, 1997). The large volumes of insecticides applied for H. armigera control is therefore not justifiable, since damage levels greater than 20% seldom occur and natural enemies are abundant. Approximately 170 parasitoid species and a large number of predators of H. armigera have been reported from southern and east Africa (Cherry et al., 2003). Furthermore, a nuclear polyhedrosis virus that occurs freely in nature, contribute significantly in controlling H. armigera populations in southern Africa.

CONCLUSION Although many insect species occur on sunflower in Africa, only a few of these obtain pest status. Insecticide application for control of pest species should therefore be done with care since economic injury levels of all, except for H. armigera, have not been determined and natural enemies are plentiful. Further research on the hemipteran pest complex of this crop is needed since the presence of these pests may lead to qualitative crop losses.

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REFERENCES Awmack, C.S. & Leather, S.R. 2002. Host plant quality and fecundity in herbivorous insects. Annu. Rev. Entomol. 47: 817-844. Charlet, L.D., Brewer, G.J. & Franzmann, B.A. 1997. Sunflower insects, pp.183-261 In: Schneiter, A.A. (Ed.). Sunflower technology and production. Madison, Wisconsin, USA. Cherry, A., Cock, M., Van den Berg, M. & Kfir, R. (2003). Biological control of Helicoverpa armigera in Africa. In: Biological control in IPM systems in Africa. Neuenschwander, P., Borgemeister, C. and Langewald, J. (Eds). CABI Publishing in association with the ACPEU Technical Centre for Agricultural and Rural Cooperation (CTA) and the Swiss Agency for Development and Cooperation (SDC). pp 329-346. Cockerell, T.D.A. 1916. Sunflower insects in California and South Africa. Can. Entomol. 48: 76-79. Drinkwater, T.W. 1999. Other soil insect in maize. In: Protection of grain crops: Principles & Practices. McDonald, A.H. & Van Rensburg, J.B.J (Eds.). p 5 – 13. Du Plessis, H. 1997. Feasibility of chemical control of the African bollworm, Helicoverpa armigera (Hübner) (Lepidoptera: Noctuidae) on cultivated sunflower in South Africa. Afr. Crop. Sci. J. 5: 47-53. Du Plessis, H., Byrne, M.J. & Van den Berg, J. 2005. Chemical control of Nysius natalensis Evans (Hemiptera: Orsillidae), a pest of sunflower in South Africa. S. Afr. J. Plant Soil. 22: 94-99. Du Plessis, H., Byrne, M.J. & Van den Berg, J. 2007. Distribution and host plant range of Nysius natalensis Evans (Hemiptera: Orsillidae), in the sunflower production area of South Africa. Afr. Entomol. 15: 310-318. Du Plessis, H., Byrne, M.J. & Van den Berg, J. 2011. The effect of temperature on Nysius natalensis Evans (Hemiptera: Orsillidae) development and survival. Afr. Entomol. 19:709-716. Du Plessis, H., Byrne, M.J. & Van den Berg, J. 2012. The effect of different host plants on the reproduction and longevity of Nysius natalensis. Entomol. Exp. App. 145: 209-214. Du Plessis, H. 1999. Sunflower pest identification and control. In: Protection of grain crops: Principles & Practices. McDonald, A.H. & Van Rensburg, J.B.J (Eds.). p 27 – 32. Du Toit, A.P. & Holm, E. 1992. Diversity, abundance and behaviour of diurnal insects on flowering capitula of commercial sunflower in the Transvaal. S. Afr. J. Plant Soil. 9: 3436. Evans, A.C. 1951. Entomological research in the overseas food corporation (Tanganyika). Proc. Assoc. Appl. Biol. 38: 526-529. Henry, T.J. 1997. Phylogenetic analysis of family groups of the infra-order Pentatomorpha (Hemiptera: Heteroptera), with emphasis on the Lygaeoidae. Ann. Entomol. Soc. Amer. 90: 275-301. Hill, D.S. 1975. Agricultural insect pests of the tropics and their control. Cambridge Univ. Press, London, 516pp. Khaemba, B.M. & Mutinga, D.J. 1982. Insect pests of sunflower (Helianthus annuus L.) in Kenya. Insect Sci. Appl. 3: 281-286. Kruger, M., Van den Berg, J. & Du Plessis, H. 2008. Diversity and seasonal abundance of sorghum panicle-feeding Hemiptera in South Africa. Crop Prot. 27: 444-451.

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Misari, S.M. 1990. Pest complex of sunflower (Helianthus annuus L.) in parts of Nigerian savanna. Savanna 11: 1-11. Nel, A.A. 1998. The effect of a diurnal period of supra-optimal temperature on the seed vigour of sunflower. S. Afr. J. Plant Soil. 15: 19-21. Rajamohan, N., 1976. Pest complex on sunflower – a bibliography. PANS 22: 546-563. Riekert, H.F. & Van den Berg, J. 2003. Evaluation of maize cultivars and rotation crops for resistance to damage by fungus-growing termites. S. Afr. J. Plant Soil 20: 72-75. Sharma, H.C. 2001. Cotton bollworm/legume pod borer, Helicoverpa armigera (Hübner) (Noctuidae: Lepidoptera): biology and management. Crop protection compendium. International Crops Research Institute for the Semi-Arid Tropics. Patancheru, A.P. India. Schneiter, A.A. & Miller, J.F. 1981. Stages of sunflower development. Crop Sci. 21: 901-903. Sweet, M.H. 1964. The biology and ecology of the Rhyparochrominae of New England (Heteroptera: Lygaeidae). Part 1. Ent. Am. 43: 1-124. Sweet, M.H. II. 2000. Seed and chinch bugs (Lygaeoidae). Chapter VI. pp. 143-264. In: Schaefer, C.W. & Panizzi, A.R. (Eds.). Heteroptera of economic importance. CRC Press, New York. Von Maltitz, E.F. 1993. The American bollworm, Heliothis armigera (Hübner) (Lepidoptera: Noctuidae) on sunflower. II. Feeding site preference of the larvae. Phytophylactica 25: 249-252. Zalucki, M.P., Daglish, S., Firempong, S. & Twine, P.H. 1986. The biology and ecology of Helicoverpa armigera (Hübner) and H. puctigera Wallengren (Lepidoptera: Noctuidae) in Australia: what do we do? Aust. J. Zool. 34: 779-814.

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Chapter 11

SOIL AMENDMENTS AND THEIR EFFECTS ON SUNFLOWER GROWTH Fernando López-Valdez1*, Fabián Fernández-Luqueño2, Perla Xóchitl Hernández-Rodríguez1, Minerva Rosas-Morales1 and Silvia Luna-Suárez1 1

Centro de Investigación en Biotecnología Aplicada, Instituto Politécnico Nacional, Tepetitla de Lardizábal, Tlaxcala, México 2 Natural Resources and Energy Group, Cinvestav-Saltillo, Coahuila, México

ABSTRACT An interesting topic in agriculture is the search for forms of fertilisation that have a low impact on soil, plants, humans, and the environment. Wastewater treatment plants almost always separate the organic matter, called wastewater sludge or sewage sludge. This sludge is rich in mineral macronutrients such as nitrogen (ammonium or nitrate), phosphorous (phosphate), potassium, and micronutrients. The sewage sludge might provide the majority of necessary nutriments for growing plants, and also provide many beneficial effects when it is applied to soils, resulting in an improvement in the chemical, physical, and biological characteristics. We know that wastewater sludge and soil microorganisms play an important synergetic role on the released and available nitrogen. Although urea is the most accepted type of fertiliser used worldwide, it does have some inconvenient drawbacks such as pH changes, microflora modifications from the soil, and others; these issues must be considered in order to avoid N losses. An interesting alternative might be the use of Plant Growth-Promoting Rhizobacteria (PGPR, soil bacteria that colonize the roots of plants by inoculation onto seeds or roots and that allow enhanced plant growth) as helpers. Examples such as Pseudomonas, Azotobacter, Burkholderia, Klebsiella, and Bacillus, among other genera, have been reported. In this *

Corresponding author: F. López-Valdez, Centre for Research in Applied Biotechnology (CIBA) - Instituto Politécnico Nacional. Carr. Est. Sta. Inés Tecuexcomac – Tepetitla km. 1.5 s/n, Tepetitla de Lardizábal, Tlaxcala, C.P. 90700, México. Tel.: +52 55 5729 6000 ext. 87805; Fax: +52 248 487 0762. E-mail address: [email protected].

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F. López-Valdez, F. Fernández-Luqueño, P. X. Hernández-Rodríguez et al. chapter, we focus on specific types of fertilisation or soil amendments and their effects on this important crop. In particular, Bacillus subtilis has showed promising results. It is known that these bacteria might improve plant growth by several direct mechanisms as well as indirect mechanisms, such as controlling phytopathogen organisms, or a combination of both. A regular strain of Bacillus subtilis was co-inoculated with urea in sunflower roots, and it was found that the strain temporarily stimulated sunflower cultivar growth. We found that B. subtilis improves the sunflower establishment, particularly its strength and vigour during the early stages, resulting in an increase of the mass and length of roots compared with the control treatment. Application of either organic or mineral fertiliser improved the crop yield under various conditions, but fertilising the crops with sewage sludge might be more environmentally friendly than using mineral fertiliser. Wastewater sludge has a high NH4+ content that is slowly oxidized to NO3while being absorbed by sunflower cultivars. In both cases (organic amendment or bacterial inoculation with urea), the NO3- is not lixiviated or leaked during experiments. Accordingly, the wastewater sludge and B. subtilis might be potential methods of fertilisation for sunflowers or any ornamental cultivars.

Keywords: Bacillus subtilis, fertilisers, sunflower, urea, wastewater sludge

INTRODUCTION Agriculture is an essential discipline that concerns the possible cultivation of domestic plants in order to improve food or biomass productions. Enhancing food security and a sustainable production of foods (agricultural sustainability) are modern global topics. Therefore, fertilisation is a critical topic, particularly aspects such as the reasonable application of fertilisers, reutilization of materials (wastewater sludge, manure, tannery sludge, whey, biochar, inter alia), and application of beneficial microorganisms (PGPRs, mycorrhizal fungi, among other microorganisms) as soil amendments. These could all be strategies that lead to a sustainable agriculture. In this chapter, we explore some of the strategies that we have used to experiment on an important crop, the sunflower.

SUNFLOWER The sunflower (Helianthus annuus L.) is a significant crop and an attractive ornamental plant known throughout the world. This crop is a considerable source of edible oil, protein (Nesterenko et al., 2013; Lin et al., 1974), oil for diesel biosynthesis, phytoextraction, and even animal feed. It shows a well-developed and deeply penetrating root system, allowing a good establishment, which is why it is considered a drought-tolerant plant (Stone et al., 2002). Therefore, the sunflower is easily cultivated, exhibiting growth with minimal or no fertilisation (Ruiz and Maddonni, 2006), and even exhibiting growth under rain-fed conditions. Mexico might be the origin of domestication of this crop. There is evidence of some early remains of H. annuus discovered at the San Andrés site in the Gulf Coast region of Tabasco, Mexico, constituting the earliest record of the domesticated sunflower (Lentz et al., 2001; Wills and Burke, 2006).

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CHEMICAL FERTILISERS It is known that nitrogen (N) is the most important mineral nutrient for non-leguminous plants. This mineral has several chemical forms within the soil, such as ammonium (NH4+-N), nitrite (NO2--N), and nitrate (NO3--N); ammonium and nitrate are both valuable ions for plants. There are several synthetic N fertilisers, such as ammonia, urea, ammonium nitrate, calcium ammonium nitrate, ammonium sulphate, and urea ammonium nitrate, among others. The world's production of ammonia (NH3) exceeded 134.2 x 109 kg y-1 in 2011 (IFA, 2013). With the highest N content (82%), it is feedstock for N fertilisers and other products. However, when ammonia is applied as aqua ammonia (dissolving ammonia in water from 20% to 24% N solution) as fertiliser, special care must be taken: it must not be placed in close proximity to seeds, and it must be handled carefully for safety purposes. Urea is the most common N fertiliser used worldwide, constituting over 71.1 x 109 kg y-1 in 2011 (IFA, 2013). The production of ammonia and urea is increasing annually; in fact, the ratio of world urea production is about 2.1 x 106 kg y-1 (the result was estimated from data from 2002 to 2011). Urea has several advantages over others fertilisers, such as a high N content (46.7%) and solubility (20 °C) at 1,080 g L-1. In addition, it is easier to handle than NH4NO3 or ammonia, easier to store, less corrosive to machinery, and less likely to explode or burn. However, some problems have been reported, including damage to seeds, seedlings or young plants; NO2- toxicity; phytotoxicity of foliar-applied urea; and volatilization of ureaN as NH3 (Bremner, 1995).

WASTEWATER SLUDGE Wastewater sludge or sewage sludge is an unavoidable by-product from wastewater treatment plants. The wastewater sludge is rich in organic matter as well as macro- and micronutrients for plants (N, P, S, and minerals). It can even provide C and energy sources for the growth of soil microorganisms (López-Valdez et al., 2010, 2011b). Also, it has the potential to improve the physical, chemical, and biological properties of the soil. However, sewage sludge can also contain some pathogens—bacteria, virus, protozoa cysts, or helminth ova (Jiménez-Cisneros et al., 2001; Jiménez, 2007)—which are heavy metals and organic pollutants that must be eliminated. According to USEPA, sewage sludge can be classified as Class A or Class B with regard to pathogens (2013). Class A refers to sludge that might be amended to agricultural soils with edible cultivars. In contrast, Class B refers to sewage sludge that might to applied to non-agricultural soils or even soils where ornamental plants will be cultivated. Approximately 37% of the sewage sludge produced in the world was used in agriculture and 12% was used in forestry (Fytili and Zabaniotou, 2008). The remainder of the sludge was not used, commonly resulting in its being confined or burned. Application of wastewater sludge in agriculture is an alternative disposal approach that, relative to the burn-up approach, offers the opportunity to recycle nutrients that enhance plant growth, return organic matter to the soil (López-Valdez et al., 2011b), and avoid CO2-C return to the atmosphere. It has been reported that sludge significantly increases the yield in apple trees (Malus pumila Mill.) (Bozkurt et al., 2010), commun beans (Phaseolus vulgaris L.) (Fernández-Luqueño et al.,

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2010), cotton (Gossypium hirsutum L.) (Samaras et al., 2008), mung beans (Vigna radiata L. cv. Malviya janpriya) (Singh and Agrawal, 2010b), rice (Oryza sativa L. cv. Pusa sugandha) (Singh and Agrawal, 2010a), soybeans (Glycine max (L.) Merrill) (Souza et al., 2009), and sunflowers (Helianthus annuus L.) (Lavado, 2006). The goal of sewage sludge application into the soil is to provide an alternative, to restore degraded or N-depleted soils and simultaneously enhance plant growth (Fernández-Luqueño et al., 2009; López-Valdez et al., 2010, 2011b).

BACTERIA AS HELPERS (PGPR) Another compelling alternative is the application of microorganisms such as bacteria and filamentous fungi (as mycorrhizal fungi). Bacteria are the most abundant microorganisms in soil (up to 6 x 108 cells g-1 of soil and a weight of approximately 10,000 kg ha-1) (Kilian et al., 2000). The number of bacteria found in soil depends on the season, the type of soil, the cultivar used, the moisture content, the oxygen supply in the soil, the tillage and fertilisation of the soil, the penetration of the soil by plant roots, and the depth from which the soil samples were taken (Kilian et al., 2000). Pseudomonas, Arthrobacter, Achromobacter, Clostridium, Micrococcus, Flavobacterium, Azospirillum, Azotobacter, and Bacillus are the most representative genera of bacteria from soil and other environments (López-Valdez et al., 2011a). Bacteria have notorious advantages over fungi such as faster growth, a ubiquitous presence, and the ability to be cultured in vitro. Bacteria might even compete for an ecological niche; bacteria produces metabolites or enzymes that enhance or stimulate the growth of plants through direct or indirect mechanisms. The Bacillus genus shows a wide spectrum of mechanisms that might (a) stimulate plant growth as fungistatic or bactericidal compounds (Singh and Deverall, 1984; Ongena et al., 2005; Forchetti et al., 2007; Sang et al., 2008; Swain and Ray, 2009; Alvindia, 2013), (b) produce phytohormones involved in root or shoot growth (Araújo et al., 2005; Yao et al., 2006; Karadeniz et al., 2006; Forchetti et al., 2007; Swain and Ray, 2009; Ahmed and Hasnain, 2010), (c) induce systemic resistance (Gupta et al., 2000) by volatile organic compounds (Ryu et al., 2003; Ping and Boland, 2004), (d) colonize plant roots (Dijkstra et al., 1987) by remaining very close to the root tip by passive displacement, and (e) produce extracellular enzymes. Another interesting result is the reduction of ethylene production, as described by Penrose and Glick (2003). Also, the Bacillus genus is known to affect the fruit ripening, leaf senescence, flower abscission, germination, cell elongation and proliferation, nodulation, and response to plant pathogen attack. Hence, bacteria that can stimulate or promote plant growth are called PGPR.

SAMPLING SITE AND PROPERTIES OF SOIL We used several soils from different land uses. The first sampling site was located in the former lake of Texcoco (Mexican valley, Mexico; 19°30‘ N, 98°53‘ W) at an altitude of 2,250 m above sea level. This soil is alkaline saline, classified as Typic Ustifluvent with a pH between 8.5 and 11, electrolytic conductivity (EC) between 4 and 170 dS m-1, and water holding capacity (WHC) of 684 g kg-1 soil. The former lake bed is not cultivated, but some

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grasses and small trees can grow. More details are available in López-Valdez et al. (2010). The second soil samples were obtained from Otumba (Estado de México, Mexico; 19°42‘ N, 98°49‘ W), near the former lake Texcoco. The soil, classified as Typic Fragiudepts, was characterized as sandy loam, pH 7.6, EC of 1.1 dS m-1, WHC of 530 g kg-1 soil, and an organic C content of 7.2 g C kg-1 soil. This soil was cultivated with maize, receiving a minimum amount of mineral fertiliser without being irrigated. More details are available in Fernández-Luqueño et al. (2009). The third sampling site is located in Alcholoya, an Acatlán village (Hidalgo, Mexico; 2,120 m above sea level and 20°09‘ N, 98°26‘ W). The soil was classified as Typic Fragiudepts with pH 6.5, EC 0.7 dS m−1, a WHC of 846 g kg−1 soil, an organic C content of 11.1 g kg−1 soil, and a total N content of 1.0 g kg−1 soil. This soil received organic fertiliser (cow excreta) occasionally. More information is available in López-Valdez et al. (2011b). All soils were sampled by augering (0 to 15 cm depth) from three plots equivalent to 0.5 ha. Soils from each plot were pooled and sieved. In total, three soil samples were obtained.

CHARACTERISTICS OF THE WASTEWATER SLUDGE Reciclagua (Sistema Ecológico de Regeneración de Aguas Residuales Ind., S.A. de C.V.) in Lerma (Estado de México, Mexico) treats wastewater from various sources, mainly alimentary industries and households. In the primary treatment, the wastewater is mixed with a flocculant, and the sludge obtained is passed through a belt filter in order to reduce the water content. Although the concentration of heavy metals and toxic organic compounds has been low, (Franco-Hernández et al., 2003) this sludge can be classified as Class B due to its pathogen content. The characteristics of sludge were pH 8.1, CE 7.9 dS m-1, water content 847 g kg−1, organic C content 288 g kg−1, and total N content 41.8 g kg−1. The mineral N was NH4+-N 13 g kg-1, NO2--N 8.3 mg kg-1, and NO3--N 122 mg kg-1 on a dry matter. These characteristics show that sewage sludge is a strong candidate for an organic fertiliser, since the C/N ratio is over 14, meaning that sludge has a high N content. Almost 30% is ammonium, indicating that the mineral N is immediately available for plants; the remaining 70% is organic N (such as proteins, amino acids, and nucleic acids, among others), available for microorganisms that mineralize organic N up to mineral nitrogen. This mineral N is released slowly and remains available for plants.

BACILLUS SUBTILIS We studied a regular strain of Bacillus subtilis that was isolated from the potato rhizosphere (identified by 16S ribosomal RNA, rRNA, gene sequencing). The strain was provided by Dr. Olalde-Portugal (Laboratory of Microbial Ecology, Department of Biotechnology and Biochemistry, Cinvestav, Gto., Mexico). This strain was tested and selected due to its ability to provide positive results in some in vitro tests, thus demonstrating its antagonistic activity against some phytopathogenic fungi (Fusarium oxysporum and Rhizoctonia solani AG1). Also, this strain embodies many characteristics, such as the ability to solubilize phosphate as described by Burr et al. (1984), the 1-aminocyclopropane-1-

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carboxilate deaminase enzymatic activity as reported in Penrose and Glick (2003), the indole3-acetic acid production by the colorimetric method (Azcón et al., 2009), and the root colonization on maize and sunflower in a Petri dish using Phytagel (Sigma Co.) as a medium (Dijkstra et al., 1987). This strain belongs to a collection that is being explored as PGPR. In our opinion, this is a regular or common strain. More information can be found in LópezValdez et al. (2011a).

METHODS The sunflower seeds were provided by the Departamento de Fitotecnia, Universidad Autónoma de Chapingo, Texcoco, Estado de México, Mexico [Department of Plant Science, Autonomous University of Chapingo, Texcoco, Estado de México, Mexico]. The non-treated seeds were received for amendments with sewage sludge, urea, or unamended soil. However, according to López-Valdez et al. (2011a), the seeds were disinfected for inoculation with B. subtilis and were incubated in an agar-agar plate under aseptic conditions in order to determine whether microorganisms grow after one day of incubation. Afterward, these seeds were dressed with a suspension of B. subtilis at 107 CFU mL-1 in 1% carboxymethylcellulose. One hundred and eight sub-samples were prepared as follows: each sub-sample contained 6.5 kg soil in a cylindrical pot with the purpose of obtaining five treatments, three plots, three replicates, and three sampling times (at 37, 60, and 95 days after sowing) during the experiment (López-Valdez et al., 2011b). At the onset of the experiment, 0.5 g urea was added to nine pots from each of the three sampled plots; twelve days after emergence, the nine plantlets were amended with another 0.5 g urea (equivalent to 150 kg N ha-1, the UREA treatment). Nine others pots were amended with 30 g sludge (150 kg N ha-1, the SLUDGE treatment), nine were left unamended (the PLANT treatment), and nine were left unamended and unsown (the CONTROL treatment). The last treatment was amended with B. subtilis (the BACTERIA treatment); the seeds were disinfected, dried, and dressed with the B. subtilis suspension (as described above). In this treatment, the plants were fertilised with 0.5 g urea (75 kg N ha-1). Each pot was sown with three sunflower seeds. Eight days after emergence, two of the three plantlets in each pot were discarded. The tap water was analysed, 19 kg mineral-N ha-1 was additionally added to each treatment during the whole experiment, and no water was leached out from the pots (López-Valdez et al., 2011b). At the onset of the experiment, approximately every two days up to 30 days after sowing, the columns were airtight and the atmosphere was analysed for CO2 and N2O at 0, 3, 15, and 30 minute intervals. On every sample day (37, 60, and 95 days), nine pots were selected at random from each treatment. Soil samples were collected at depths of 0-15 cm and 16-30 cm. The roots were separated from the shoots. The variables measured in the plants were shoot height, root length, fresh shoot weight, fresh root weight, dry shoot weight, dry root weight, seed weight per plant, number of seeds per plant, and total N content; in the soil, the variables measured were pH, EC, NH4+-N, NO2--N, and NO3--N. Also, the gas production was measured as CO2-C and N2O-N. Further details can be found in López-Valdez et al. (2011b). Significant differences between plant and soil characteristics as a result of the various treatments were determined by analysis of variance (ANOVA) and based on the least

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significant difference test, using the general linear model procedure (PROC GLM; SAS Institute Inc., 1989).

UREA NOTES Sunflower Characteristics (Alcholoya Soil) According to our results, the UREA treatment was not significantly different in shoot height and dry root weight compared with the other treatments at one, two, and three months. After two months, the root length and fresh root weight were similar to the BACTERIA and PLANT treatments. Finally, the effect of urea was significantly different in the dry shoot weight compared with the PLANT treatment. Compared with the unfertilised plants, the urea enhanced the dry shoot weight, causing a significant increase of total N content per kg of dried plant (P