Olive propagation manual andrea fabbri, giorgio bartolini, maurizio lambardi, stan kailis csiro 2004 by agrihorti

Olive Propagation Manual

Andrea Fabbri Dipartimento di Biologia Evolutiva e Funzionale UniversitĂ degli Studi di Parma, Italy

Giorgio Bartolini CNR, Istituto per la Valorizzazione del Legno e delle Specie Arboree, Polo Scientifico, Sesto Fiorentino (Firenze), Italy

Maurizio Lambardi CNR, Istituto per la Valorizzazione del Legno e delle Specie Arboree, Polo Scientifico, Sesto Fiorentino (Firenze), Italy

Stanley George Kailis School of Plant Biology, Faculty of Natural and Agricultural Sciences, Crawley, WA Australia

© CSIRO 2004 All rights reserved. Except under the conditions described in the Australian Copyright Act 1968 and subsequent amendments, no part of this publication may be reproduced, stored in a retrieval system or transmitted in any form or by any means, electronic, mechanical, photocopying, recording, duplicating or otherwise, without the prior permission of the copyright owner. Contact CSIRO PUBLISHING for all permission requests. National Library of Australia Cataloguing-in-Publication entry Olive propagation manual. Bibliography. Includes index. ISBN 0 643 06676 4. 1. Olive – Propagation – Handbooks, manuals, etc. I. Fabbri, Andrea, 1948– . II. CSIRO Publishing. 634.6353 Available from Landlinks Press 150 Oxford Street (PO Box 1139) Collingwood VIC 3066 Australia Telephone: Local call: Fax: Email: Web site:

+61 3 9662 7666 1300 788 000 (Australia only) +61 3 9662 7555 publishing.sales@csiro.au www.landlinks.com

Front cover Photos by the authors Figures and Plates: Figures 1.3, 3.7, 3.16, 3.17, 4.1, 4.4, 4.5, 6.3, 6.4, 6.7, 6.8, 6.12, 6.13, 6.18, 6.23. Plates VI, VII, IX, XI, XII, XIII, XVI, XVII, XVIII, XIX by Andrea Fabbri Figures 3.2, 3.4, 3.5, 3.6, 3.15, 3.18, 3.19, 4.8, 6.1, 6.2, 6.5, 6.6, 6.9, 6.10, 6.11, 6.14, 6.15, 6.16, 6.17, 6.19, 6.20, 6.21, 6.22. Plates II, III, IV, V, XIV, XV by Giorgio Bartolini Figures 3.8, 3.12, 3.13, 3.14, 4.2, 5.2, 5.3, 5.4, 5.5, 5.6. Plates I, VIII, X, XX, XXI, XXII, XXIII, XXIV, XXV, XXVI, XXVII, XXVIII, XXIX, XXX, XXXI, XXXII by Maurizio Lambardi Figure 1.2 by Stan Kailis Set in 10.5/13pt Minion Cover and text design by James Kelly Typeset by J&M Typesetting Printed in Australia by Ligare Disclaimer While the authors, publisher and others responsible for this publication have taken all appropriate care to ensure the accuracy of its contents, no liability if accepted for any loss or damage arising from or incurred as a result of any reliance on the information provided in this publication.

Dedicated to Gabriella, Pia Lucia, Marina and Lefki

Authors Andrea Fabbri is Full Professor of Arboriculture and Pomology at the University of Parma, Faculty of Agriculture. A large part of his professional activity has been focused on research and teaching on the olive, in various laboratories and universities in Italy and California. His main olive research subjects have been physiology and anatomy of adventitious rooting in cuttings, flower biology, freezing injury, systematic pomology (biomolecular characterisation), organic cultivation, ecophysiology. His research papers have been published on the most important horticultural scientific journals. He is presently involved in the development of olive cultivation in cold areas of Northern Italy.

Giorgio Bartolini is Head Researcher of the National Research Council (CNR) at the Istituto per la Valorizzazione del Legno e delle Specie Arboree (Trees and Timber Institute) of Florence, Italy. His activity has been focused on research concerning woody plants propagation by cuttings of olive, peach, grapevine, etc.; management of an international data bank (world olive germplasm); individuation of morphological and molecular markers for cultivar characterisation in O. europaea; gene expression induced by low temperature. He participates in joint research projects with research institutes and universities of Italy, California and Spain. He is author or co-author of many research papers published in international journals, books and in proceedings of international congresses and symposia.

viii

Olive Propa ga tion

Maurizio Lambardi is a Researcher of the National Research Council (CNR) at the Istituto per la Valorizzazione del Legno e delle Specie Arboree (Trees and Timber Institute) of Florence, Italy. He is also Lecturer of Woody Plant Biotechnology and Micropropagation at the University of Modena and Reggio Emilia, Faculty of Agriculture. He has wide-ranging expertise on woody plant micropropagation and biotechnology. His main areas of research on the olive have concerned seed germination, tissue culture, genetic transformation, germplasm cryopreservation. He is the author or co-author of over 90 scientific papers and reviews, published in leading international journals, books, proceedings of international congresses and symposia.

Stanley George Kailis is Professorial Fellow at The University of Western Australia in the School of Plant Biology. His antecendents came from the Dodecanese Island, Megisti. His interests focus on quality aspects of the olive. He is particularly interested in the propagation of olive varieties such as Kalamata, Konservolia, Leccino and Manzanilla. Stanley has made presentations on the olive at national and international forums. He has published numerous research papers in national and international journals. He has conducted numerous schools and workshops in Australia on olive growing, olive oil and table olive production, organoleptic evaluation of olive products and olive propagation.

Contact email andrea.fabbi@unipr.it g.bartolini@ivalsa.cnr.it lambardi@ivalsa.cnr.it skailis@agric.uwa.edu.au

Foreword This publication deals with all issues concerning olive propagation. After an historical and thoroughly technical overview of the several available traditional techniques, the text focuses on the more modern, and more extensively employed, nursery procedures. The major recent scientific acquisitions, and the development of technological innovation that is a result of this knowledge, are also illustrated. The authors have interpretated the several subjects in a way that is both synthetical and exhaustive, and in a form that is accessible to all readers, be they students, technicians or farmers. This work represents a stimulus to scientific research in the sector, yet is also of universal interest. Their well-documented work is rich with related issues that invite the reader to go further in the field of plant propagation. Franco Scaramuzzi President â&#x20AC;&#x2DC;Accademia Economico-Agraria dei Georgofiliâ&#x20AC;&#x2122; of Florence, Italy

Contents

Authors

vii

Foreword

Introduction

1.1 1.2 1.3

2 3 7

Fundamentals of plant propagation The importance of propagation for olive cultivation Olive propagation today

Flower and fruit biology in the olive

2.1

Olive cycles 2.1.1 Life or biological cycle 2.1.2 Annual cycle 2.1.3 Fruiting cycle

8 8 11 11

2.2

Stages of olive reproduction 2.2.1 Flower induction 2.2.2 Flower differentiation 2.2.3 Reproductive structures Inflorescence Flowers and flowering 2.2.4 Anthesis and pollination Sterility Fruit set 2.2.5 Fruit growth and seed development

12 12 13 13 13 14 16 18 18 19

Propagation by cutting

3.1

22 22 26

Biology of adventitious root formation in cuttings 3.1.1 Morphology and anatomy 3.1.2 Physiology

xii

Olive Propa ga tion

3.2

Techniques of propagation by cutting 3.2.1 Plant material

27 27

Branches

Shoots

Ovules

Suckers (or pollards)

3.2.2 Shoot collection and cutting preparation

3.2.3 Auxin treatments to promote rooting of cuttings

Using hydro-alcoholic solutions for quick-dip treatments

Using talcum powder formulations

Commercial preparations

Dilute auxin solutions

3.2.4 Techniques to improve the effectiveness of auxin treatments

Soaking the cuttings

Wounding

3.2.5 Root promoting compounds, used in combination

with auxin treatments

Growth regulators

Fungicides

Propagation by grafting

4.1

The purposes of grafting

4.2

Production of olive seedlings

4.2.1 Stone collection and quality of olive seeds

Olive seed dormancy

Sources of stones and quality of seeds

Harvesting time

Extraction, cleaning and storage of stones

4.2.2 Technique of propagation by seed

Pre-sowing treatments

Sowing

Germinating small quantities of valuable seed

Germinating seeds of other Olea species

Transplant and growth of seedlings

xiii

Contents

4.3

Theoretical and practical aspects of grafting 4.3.1 Histology of graft union 4.3.2 Graft incompatibility 4.3.3 Grafting on seedling rootstocks Collection and conservation of scionwood Grafting time Bark grafting technique Care of the grafted plants 4.3.4 Grafting on adult trees Topworking Grafting on suckers Grafting on wild olive trees 4.3.5 Grafted cuttings

55 55 58 58 59 59 59 62 62 62 63 64 73

4.4

Production of clonal rootstocks in the olive

4.5

Grafted plants or self-rooted plants?

5. In vitro propagation of the olive 5.1

Micropropagation 5.1.1 Stage 0: Collection of explants Explants from in field growing stock plants Explants from greenhouse growing stock plants 5.1.2 Stage I: Initiation of cultures Disinfection of explants End of Stage I 5.1.3 Stage II: Shoot proliferation Medium formulation Growth regulators Culture conditions and subculturing Shoot elongation 5.1.4 Stage III: Shoot rooting Rooting on IBA or NAA-containing medium Additional root-promoting methods Root induction by means of dipping method 5.1.5 Stage IV: Acclimatisation of plantlets Aims of acclimatisation How to acclimatise olive microplants 5.1.6 Field performance of micropropagated plants

77 77 81 81 81 82 82 83 83 84 84 87 87 87 87 87 88 88 88 89 89

xiv

Olive Propa ga tion

5.2

Somatic embryogenesis 5.2.1 Somatic embryogenesis from mature tissue explants

90 90

5.3

Synthetic seeds and micrografting

5.4

Considerations on olive micropropagation

5.5

In vitro conservation of olive 5.5.1 Slow growth storage 5.5.2 Cryopreservation

92 92 93

The olive nursery (stock plants, structures, equipment and operations) 96 6.1

Stock plants 6.1.1 Phase change 6.1.2 Training systems for stock plants

97 98 98

6.2

Greenhouses 6.2.1 Glass greenhouses 6.2.2 Plastic film greenhouses 6.2.3 Rigid-panel greenhouses PVC (polyvinylchloride) FRP (fibreglass-reinforced plastic) Polycarbonate, methacrylate, and other materials 6.2.4 Location of the greenhouse 6.2.5 Greenhouse heating 6.2.6 Greenhouse cooling 6.2.7 Computers for ambient control 6.2.8 Beds 6.2.9 Propagation greenhouses Control zone (instrumentation) Rooting zone Hardening zone 6.2.10 Shelter and storage

99 100 101 101 101 102 102 102 103 104 106 106 106 106 107 110 110

6.3

Substrates 6.3.1 Rooting substrates 6.3.2 Substrates for growing plants

111 111 113

6.4

Environmental conditions for rooting 6.4.1 Temperature 6.4.2 Humidity 6.4.3 Light

113 113 114 118

Contents

6.5

Hardening (1st transplant) 6.5.1 Containers and substrates 6.5.2 Transplanting environment

119 120 121

6.6

Plant growing (2nd transplant) 6.6.1 Containers and location of plants 6.6.2 Irrigation and fertilisation

122 123 123

6.7

Plant training

125

6.8

Plant certification 6.8.1 Genetic origin 6.8.2 Good sanitary conditions

126 127 128

Conservation of olive germplasm

129

Appendices Appendix 1 Rooting ability of all olive cultivars of which scientific literature is available

131

Olive germplasm collections throughout the world

133

Appendix 2

1 Introduction

The European olive (Olea europaea L.) is a major source of edible oil and processed edible olives. Today, the olive is cultivated over a total world surface area of almost 10 million hectares, on 60% of which it represents the main crop. The traditional area of olive cultivation is the Mediterranean basin, which includes 95% of the olive orchards of the world, and where more than the 95% of the olive oil and the 75% of the table olives are produced. A rough estimate of the global number of olive trees is over 800 million. The annual yield of olives is estimated at 10 million tonnes, most of which is used for oil production and less than 10% consumed as table olives. Over the last 30 years, the production and the consumption of olive oil have increased together. It is unlikely that this trend will change in the near future, considering the recent introduction or increase of olive cultivation and olive-oil consumption in countries such as Japan, Australia, China and South Africa. Remarkable increases (up to 10 fold) have been observed in several countries, including Australia, where olive-oil consumption has passed the level of 1 kg/person/year. Hence, the volume of olive oil consumed worldwide during the next 10 years is expected to exceed three million tonnes annually. Such volumes of olive oil will require active farming programs and olive trees for both new olive orchards and replacement in existing olive groves. Furthermore, as the olive industry moves from traditional manual methods to mechanised operations, planting stock will need to be developed to meet future challenges. Varietal selection will need to be directed to clones that are early bearing, disease resistant and able to be mechanically harvested, and which will produce quality fruit and oil. Although these developments are the province of plant breeders, the follow-through will fall to the olive propagators and associated nurseries.

Olive Propa ga tion

1.1 Fundamentals of plant propagation Plant propagation consists of the application of specific biological principles and of particular techniques for the multiplication of plants. The plants obtained in the process should be identical, or as similar as possible, to the plants from which they are derived. All living organisms can be described by genotype and phenotype. Genotype is the whole set of genetic characteristics of the organism, which are controlled by a very high number of genes Any living cell of the organism has the availability of the whole set of genes, although only a minor part of these is activated in that given cell, according to its specialisation (i.e. the differentiation process it went through). Phenotype is the sum total of visible or measurable characters of the organism that characterise it as individual (such as, in fruit trees, fruit size, shape and quality, tolerances to stress and disease, phenological stages). A fundamental principle of biology is that the phenotype is the result of an interaction, between genotype and environment. As a result, if we want to obtain plants that are as similar as possible to their parent plants, we must produce individuals with the same genotype, and place them in identical environments. This latter condition is almost impossible to achieve, and at any rate is not a concern for propagators. Propagators should instead be aware of the implications of the genetic characteristics of propagules (any plant parts used for propagation), which are fundamental in defining the qualitative results of the propagation process. In this respect we can divide the propagation systems for horticultural plants in two main groups: reproduction and multiplication. In reproduction, or gamic or sexual propagation, the propagule is the seed (in higher plants), and particularly the embryo, derived from a fertilisation process. The seed carries the genes of two parents, and these can be arranged in a countless number of combinations. Even when it derives from self-pollination, the genotype of a seed is never identical to that of any of the parent plants, as there are several mechanisms that may cause variation in gene presence and position. Therefore the progeny are always to some extent different from any of the parents, and variability also occurs within the seedling population, just as happens with humans. This means that sexual propagation cannot be used for those species in which uniformity of the plantation and true-to-typeness for a series of biological and technical requirements are essential, as is the case of fruit trees. In species like the olive, seed propagation is therefore confined to the production of seedlings, to be used as rootstocks. In multiplication, or agamic or asexual propagation, the propagule is any other part of the tree, and therefore only carries identical somatic cells. This includes shoots, roots, buds and leaves, and even cells from the ovary tissues, which belong to the stock plant genotype. These tissues, being genetically identical to the stock plant, are able to generate individuals with the same genotype, which in turn will be able to be propagated indefinitely in the same way, thus generating a â&#x20AC;&#x2DC;cloneâ&#x20AC;&#x2122;. Olive cultivars are clones, often very ancient clones, derived from countless asexual propagations, made with several techniques since ancient times. The success of propagation, whatever the technique adopted, depends on knowledge of the many aspects controlling the formation of a new individual:

Introduction

• genetic characteristics of species and cultivar which regulate their ability to be propagated • anatomical structure and physiology of the whole plant and of the organs to be propagated • methods most suited to propagate the various cultivars and rootstocks • techniques and structures, and their optimisation to obtain the best technical and economical results

1.2 The importance of propagation for olive cultivation The olive tree, Olea europaea (L.), is one of the most ancient cultivated fruit trees of the Old World, and its importance for the Mediterranean civilisations is witnessed by all classical sources. A remnant of the tropical flora of the mid-Tertiary, the olive is so typical of the Mediterranean that its presence qualifies a climate as Mediterranean, even in other areas of the world. The earliest signs of olive cultivation – the first wave – can be traced back to the 4th millennium BC and, before that, to areas of the eastern Mediterranean coasts and islands, although the ancestors of currently grown olive cultivars are still believed to have been domesticated in the mountainous territory south of the Caucasus, covering today’s eastern Turkey, western Iran, Lebanon, northern Israel, Syria and northern Iraq. From the eastern Mediterranean the olive moved westwards – the second wave – to Greece and the Aegean archipelago, although Crete and Cyprus probably belonged to the oldest olivegrowing centre. In these areas, collectively considered a secondary centre of diversity, the olive grew in importance, and possibly underwent deliberate selection by humans between the 3rd and the 2nd millennia BC. In Crete, in the 16th century BC, there existed in Knossos a huge deposit of clay jars, able to store five times the amount of oil the local population could consume in one year, thus indicating a strong possibility of a developed trade in olive oil. Around the beginning of the first millennium BC a third migration appears to have taken place. Again it was westwards, to Sicily and Tunisia, an area regarded as the olive tertiary centre of diversity. From there, around 600 BC, probably through Etruria (today’s Tuscany),

Figure 1.1 Distribution of the wild olive (Olea europaea oleaster), the progenitor of the cultivated olive. It is still present in most coastal areas of the Mediterranean basin. (Zohary & Spiegel-Roy 1975)

Olive Propa ga tion

the crop is reported by the classical historians to have reached the Romans. Up to this point the olive had moved slowly westwards, first on the ships of Phoenician merchants, and later on those of Greek colonists; these peoples had spread the crop in many other places along the Mediterranean, including Spain, France and northern Africa, with varied results. But the conquest of the whole area by the Roman legions, and its transformation into a vast united empire, made trade and communications far more intense, and the olive benefited from this situation. In addition, when Italy appeared unable to provide the required supply of olive oil, the Romans spread its cultivation in new areas, or favoured it in places where olive groves had already been established but had stagnated. The crop achieved its maximum economic importance in the 2nd to 3rd centuries AD, particularly in northern Africa, but also in Spain, Dalmatia, and French Provence. With the fall of the Roman empire, information about the olive becomes scarce. Its cultivation dropped dramatically, with the reduction in population and the abandonment of large areas that took place during the early middle ages. This was not the case in the territories under Arab rule, where the crop remained important, to the point that its cultivation was forbidden in Sicily in order to protect production in North Africa, probably the main producer at the time. In Europe, olive oil acquired new importance only in the 16thâ&#x20AC;&#x201C;17th centuries, when it became a significant trading commodity for Venetians, who imported it from their Aegean possessions such as Crete and Cyprus. It must be remembered that oil was not only used as food; it also had great importance as a medicine, and for illumination, massaging, soap production and wool processing. Thus, olive plantations slowly began to spread in the Mediterranean areas where they can still be found today, with the exception of most of northern Africa, where they were reintroduced on a large scale much more recently. The arrival of the olive in the western and southern hemisphere is also recent history. Argentina, California, Australia and South Africa â&#x20AC;&#x201C; where the enthusiasm of Mediterranean migrants for the crop had ensured its introduction â&#x20AC;&#x201C; all proved to have suitable environments for commercial olive cultivation. Both the early domestication of the olive and its diffusion in the Mediterranean region have been favoured, or should we more correctly say permitted, by the ability of the species to be propagated with simple techniques. It is certain that the earliest domesticated fruit crops could be transferred from the bush or the forest (where they were browsed by animals and man) via the most rudimentary forms of cultivation, thanks to the possibility of stabilising positive and superior characters that the early gatherers noticed in the wild plants, when transition from hunting/gathering to agriculture was taking place. This suitability to asexual propagation was true for all ancient fruit crops, such as pomegranate, grapevine, date and fig.

Introduction

Figure 1.2. Young olive orchard in Western Australia. New plantations in southern hemisphere countries follow the most modern technical guidelines.

Figure 1.3. Exceptionally grown olive tree in Greenough, Western Australia.

Olive Propa ga tion

The olive, from the origins of its cultivation until the second half of the 19th century, was only propagated agamically, by using either large cuttings, ovules or rooted suckers. The slowness of its diffusion, which was a general feature of fruit crops until recently, made it a normal practice to resort to on-farm propagation, which meant the production of small numbers of trees each year. This also meant the selection and stabilisation of local cultivars, which, given the antiquity of the crop and its spread in the region, account for the high number of genotypes found in the different Mediterranean countries. Caruso (1883) questioned these direct multiplication methods, instead advocating the advantages of indirect multiplication, i.e. grafting on seedlings. In reality, the main real advantage of the grafting technique was the possibility of mass propagating the olive, and therefore dropping the prices of the individual plants. This would produce several further advantages: cultivars with superior characteristics could easily be introduced in new areas of cultivation, planting density could be increased, and rootstocks with positive characteristics could be used. Therefore, although several researchers (Vivenza 1926; Casella 1934; Morettini 1942) in the 20th century demonstrated that direct multiplication was just as good as grafting with respect to olive-tree life and performance, the new technique spread due to the need to provide large numbers of plants for the expanding olive industry. The old systems of propagation survived until recently in many areas including Southern Italy and Andalusia (Spain), but grafting certainly became by far the most important propagation technique. The supremacy of grafting over direct rooting, apart from never-ending disputes over the field performance of the trees (see 4.5), was granted by the simple fact that sufficiently good rooting could only be obtained by cuttings 4 or more years old, and by relatively large ovules and suckers, which made the availability of propagation material quite scarce. It is not surprising then that research into the possibility of using 1- or 2-year-old olive cuttings for direct propagation started as early as 1940. Research centred on cutting characteristics, rooting substances and greenhouse environment. In less than two decades, direct propagation of olive semi-hardwood cuttings became a technical reality. Although the technique would undergo several improvements to increase its efficiency, the fundamental acquisition was available by the mid-1950s. This did not mean that grafting practices with olives were suddenly stopped. On the contrary, in Italy, which is the main producing country for olive nursery trees, cutting propagation slowly conquered a share of the market, which remained around 50% until the 1980s, increasing its share in the following decade to over 70%. More recently, olive tree production by micropropagation techniques is also gaining favour, after decades of research in numerous research stations. This reconquered supremacy of direct multiplication over grafting does not mean that this issue is settled forever; on the contrary, grafting is a technique that will most likely accompany the olive industry into the foreseeable future. The reasons for this are numerous, but some are particularly important. In the first place, not all cultivars are easily (i.e. economically) propagated from cuttings or in vitro, such as many table olive cultivars. Secondly, direct multiplication involves the use of more or less complex structures, which require money and training, and in many situations (such as new areas in developing countries) one or both of them may not be available. Thirdly, although the availability of clonal rootstocks is at present quite limited, research is involved in selecting rootstock genotypes which can improve the industry through effects on tree size, yield efficiency and stress tolerance.

Introduction

The main lesson to draw from this short history of the olive and its propagation is that all available techniques of propagation have had their importance in different historical times, although one technique has at times prevailed over others. On the other hand, the economic success of a commercial olive grove also depends on the choice of planting material and on the characteristics it bears due to the propagation technique, as described in depth in the following chapters. Besides, there are situations in which olive farmers may choose to propagate their own trees, and select the most suitable technique for their needs and conditions. Understanding the basics of olive propagation appears, therefore, to be one of the foundations for the technical training of the modern olive grower.

1.3 Olive propagation today The annual production of olive trees in the main olive-growing countries of the world is around 40 million, with 32 million in the Mediterranean basin and 8 million in the rest of the world (IOOC 2000). From a technical point of view, about 28 million trees are obtained by means of mist propagation, 7 million by grafting and 5 million by traditional techniques (ovule, cuttings from branches, etc.). The misting technique, which spread widely between 1950 and 1960, is the most common propagation method where there are no financial or technical constraints. Under such circumstances, and at any rate wherever the nurseries are medium to large in size, they can afford fairly high investment costs in terms of buildings (greenhouses), instruments (for the management and control of humidity levels, temperature, light, etc.) and skilled personnel. The material used for propagation comes from the nursery itself (stock plants), from neighbouring nurseries and from plants in production, generally from areas not more than 50 to 200 kilometres from the nursery. The result of all this is that, as a rule, the material is indigenous in origin; only where stock plants of exotic cultivars are collected and grown by nurseries or institutions can genotypes of other regions or countries be made available to local producers.

References Caruso, G. 1883. Monografia dell’olivo. Enciclopedia Agraria Italiana, vol. 3. UTET (ed), Torino (Italy), pp. 501–533. Casella, D. 1934. La propagazione dell’olivo nell’Italia meridionale. Proceedings ‘Convegno dell’olivicoltura meridionale’. Bari (Italy). IOOC (International Olive Oil Council) 2000. Catalogo mondiale delle varietà di olivo. Madrid. Morettini, A. 1942. Ricerche sul sistema radicale dell’olivo. Proceedings ‘Convegno di studi olivicoli’. Firenze (Italy), pp. 281–321. Vivenza, A. 1926. L’olivicoltura in Italia: l’Umbria. In VIII Congresso Internazionale di Olivicoltura, Rome, pp. 5–21. Zohary, D. & Spiegel-Roy, P. 1975. Beginnings of Fruit Growing in the Old World. Science, 187 (4174): 319–327.

2 Flower and fruit biology in the olive

Olive propagation may involve several aspects of plant biology. Therefore an introductory level of information on olive flower biology, which ultimately leads to seed formation, is instrumental in achieving a comprehensive knowledge of the object of propagation.

2.1 Olive cycles In the olive, as in all fruit trees, several biological and physiological cycles can be considered, according to the period and the events that are taken into consideration. Here, a life or biological cycle, an annual cycle and a fruiting cycle are synthetically described.

2.1.1 Life or biological cycle The life or biological cycle comprises the whole life-span of the tree. Trees from rooted cuttings and from grafted seedlings will be taken into consideration here (plants from micropropagation can be assimilated to self-rooted trees). The life of a cultivated olive tree is usually divided into four phases or stages. 1. Unproductive stage. During this stage the tree grows at high rates and is characterised by the absence of flowering and fruiting. Lack of production is due only to the absence of a sufficient equilibrium between the canopy and the root system. This stage should not be mistaken for the juvenile stage, which is only typical of seedlings. The very appearance of the first flowers marks its end. 2. Stage of increasing production. Flowering means production, and the tree increases its productive capacity as time passes. Its canopy grows, and with it the number of buds that are susceptible to flower induction. 3. Maturity stage, during which plant size and production have attained a maximum. Productivity can be considered constant, although it may fluctuate greatly from year to year. In this stage, the olive grove produces at its best.

Flo wer and fruit biolog y in the olive

Figure 2.1 Olive phenological stages according to Colbrant and Fabre (in Loussert & Brousse 1978). A, winter stage; B, bud break and elongation; C, inflorescence development; D, flower enlargement (flowers become spherical, on a short pedicel, and bracts diverge); E, corolla differentiation (pedicels elongate, distancing the flowers from the inflorescence axis); F, onset of anthesis, the first flowers open (corollas from green to white); F1, full bloom; G, petal fall (petals darken and separate from the calix); H, fruit set; I, fruit growth (1st stage, the size of a caryopsis); I1, fruit growth (2nd stage, the largest fruits are 8-10 mm in diameter; endocarp sclerification begins); L (not shown), vĂŠraison, the fruit colour turns from green to a dark colour (red and then black).

Olive Propa ga tion

4. Senescence stage. All processes typical of ageing (low vegetative activity, reduction of expansion of the root system, abundant flowering followed by poor fruit set, susceptibility to diseases, etc.) appear and indicate a tendency of the tree to weaken and die. The important stages in cultivated olive trees are the first three, as current practice usually involves uprooting long before senescence begins. The duration of these stages has, on average, changed with time, as olive cultivation technology has evolved. Table 2.1 shows how the durations of the different stages have been evaluated at different times. The dramatic changes in the stages of the olive life cycle are due to a number of technological inputs the industry has received in just half a century, such as soil management technology, fertilisation, pruning, training systems and planting density. Table 2.1 Duration in years of the four stages of cultivated olive trees, according to past and current reports Morettini (1950)

Maillard (1975)

Morettini (1972)

Present day

Unproductive stage

1–12

1–7

1–4

1–3

Production increase stage

12–50

7–35

4–15

3–12

15 onward

12 onward

Maturity stage

50–150

35–150

Senescence stage

150–200

Over 150

Figure 2.2 Yearly biological and cultivation cycles of olive in the Mediterranean, according to Pansiot and Rebour (1960); months indicated by roman numerals refer to northern hemisphere environments. A, rest period; B, period of active vegetative growth; B1, period of slow vegetative growth; C, flower bud differentiation; D, anthesis-fruit set (inflorescence emergence is usually four weeks before anthesis); E, fruit growth; F, pit hardening (endocarp sclerification); G, véraison; H, ripening; I, vernalisation; J, pruning; K, harvest; L, critical period for nitrogen; M, critical period for water.

Flo wer and fruit biolog y in the olive

2.1.2 Annual cycle Productive olive trees go through an annual cycle, which is well described by the succession of phenological stages (Fig. 2.1). Such stages are closely related to a series of physiological events, which in turn determine the timing of cultivation operations (Fig. 2.2). The most important phenological stages are: onset of vegetative activity (bud break), emission of inflorescences, anthesis, fruit set, and vĂŠraison (drupes turning black). All these events occur over a number of days, and both the onset of a given stage and its duration may strongly differ from year to year and among locations, owing to a number of environmental variables.

2.1.3 Fruiting cycle The fruiting cycle begins when the uncommitted bud receives the first stimuli leading to its induction to flower, and ends with the full ripening of the fruits derived from those flowers, and their eventual abscission or harvest. Unlike the annual cycle, then, the fruiting cycle extends over two, sometimes three solar years. This means that, in any given moment, two fruiting cycles are taking place on the same tree, and that they cannot fail to heavily influence each other (Fig. 2.3).

Figure 2.3 Biennial cycle of vegetative and reproductive processes in the olive (Rallo et al. 1994).

Olive Propa ga tion

2.2 Stages of olive reproduction 2.2.1 Flower induction The formation of flower-bearing buds is a process requiring the passage of the meristematic apex of the bud, undifferentiated in its early stages of growth, to a structure carrying flowers. Such a process is commonly divided into two stages: during the first one, the induction phase, the bud undergoes a series of conditionings, both internal and external to the plant, following which, in the meristematic apex, such biochemical modifications occur as ‘to commit’ it to the formation of reproductive structures. This is also termed the ‘irreversibility stage’ to indicate a condition of the bud in which there is

Figure 2.4 Schematic representation of flower bud differentiation in olive (Loussert & Brousse 1978; redrawn from Hartmann 1951). A, undifferentiated bud; B, elongation of the inflorescence axis and appearance of the terminal flower with flower primordia (S); C, well developed sepal (S) and petal (P) primordia; D, sepal and petal primordia at a more advanced stage, and appearance of stamen primordia (E).

Flo wer and fruit biolog y in the olive

only one ontogenic possibility left, i.e. of developing into a fruit bud; otherwise it will not undergo any further development. When the irreversibility stage is overcome, the bud is to be considered ‘induced’ to flower, and the next phase – differentiation – begins. The exact time of flower induction is difficult to determine, mostly because the modifications it involves are of a physiological nature, and therefore non-detectable at the microscopic level. In addition, the duration of this phase does not depend so much on the length of the process in the individual bud, but rather on the fact that it is not simultaneous in all parts of the canopy, and that it therefore takes place over a period of time, which is markedly influenced by environment and genotype. This graduality is present in all cultivars, although to different extents. Scientific evidence seems to indicate, for the olive bud, a two-step process, with a first wave of induction at the end of spring. These buds will undergo a later confirmation of their orientation to form flowers in late autumn, in accordance with the prevailing environmental and nutritional conditions that have occurred in the meantime (Fabbri & Benelli 2000).

2.2.2 Flower differentiation The differentiation stage involves a series of anatomical changes. During this stage the typical tissues of flower structures are formed, reaching completion immediately before anthesis. Flower differentiation takes place in winter (from late February to mid-March in the northern hemisphere) although in some areas it may last longer. The timing and extent of flower differentiation seem to depend on the achievement of specific chilling requirements. This means that low winter temperatures influence not so much floral evocation as the expression of a flowering potential already determined in warmer periods. As a rule, floral differentiation occurs during the 40–60 days before anthesis; therefore the process is completed as the inflorescence emerges and develops. The first signs of differentiation are a broadening of the bud apex towards a more or less flattened conical mass, with the four sepal primordia appearing as minute protuberances on the sides of the apex itself. At the axils of the leaflets enclosing the apex, more meristems (decussate couples) appear and develop, forming a branching that will form the inflorescence, which will comprise a large number of flowers. In the meantime, in each individual flower primordium, the various floral organs (petals, stamens and ovary) are formed centripetally (Fig. 2.4).

2.2.3 Reproductive structures Inflorescence Flower bud inflorescences are borne at leaf axils (a maximum of two per node). Usually flower buds are formed on the shoots developing the year before anthesis. Buds may remain dormant for more than a year and then develop into inflorescences, but in most cases undeveloped buds abscise. The inflorescence axis in the spring grows slowly, and finally emerges from the leaflets that protected it in the winter. Inserted on the main axis, at the axil of small bracts, are the secondary axils, decussate, of decreasing length from base to tip. These can be further branched, and this constitutes an important taxonomical character in cultivar description. Inflorescence colour is green to white or white-yellowish.

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Figure 2.5 Olive inflorescences (courtesy H. Rapoport).

Inflorescences (Fig. 2.5) can be described by their shape and size (10 to 70 mm), flower clustering, pedicel length, flower size, number of flowers (5 to 60). Some cultivars may bear mixed buds, i.e. developing a shoot that bears, in the same yearâ&#x20AC;&#x2122;s growth, inflorescences on the basal nodes; the phenomenon is strongly influenced by the environment and is more common in warm climates. Inflorescence development begins in early spring (e.g. about mid-April in Central Italy), roughly one month after the start of differentiation, usually beginning on the south side of the tree (in the northern hemisphere). It is gradual, and does not follow a regular scheme like anthesis in the inflorescence; it may last 6â&#x20AC;&#x201C;8 weeks, even more in warmer climates, in separate flushes. As a rule, the earlier the emission of inflorescences, the higher the expected production, as fruit set may take place in less dry conditions, but subsequent environmental events may markedly alter the forecast. Flowers are usually borne on 1-year-old shoots, but reports exist of inflorescences developed on 2- and sometimes 3-year-old branches. Flowers and flowering The olive flower is made of four verticils (whorls): calix, corolla, androecium and gynoecium (4 sepals, 4 petals, 2 stamens and 2 carpels; Fig. 2.6). The sepals are light green, short and rounded. The corolla is gamopetalous (tubed) with four white-yellowish lobes. The androecium consists of two opposite stamens inserted on the corolla, and each stamen consists of a filament topped by a large anther that is hemispherical, introverted and longitudinally dehiscent. Anthers contain bicellular pollen grains; the external walls of pollen grains have characteristic structures (Fig. 2.7). The gynoecium is a superior two-loculed ovary, made of two carpels. Each locule contains two anatropous ovules; the style is short and sturdy, and ends with a well-developed, bifid, papillate, plumose stigma. Calix, corolla, stigma and pollen exhibit characteristics that vary with the cultivar, but

Flo wer and fruit biolog y in the olive

Figure 2.6 Typical olive flower (courtesy H. Rapoport).

Figure 2.7 SEM (scanning electron microscope) photograph of olive pollen grains (Roselli 1977).

differences also exist among flowers on the same tree, and therefore such characteristics cannot be used for taxonomical purposes. Two types of flowers are present each season: perfect flowers, containing stamen and pistil, and staminate flowers, containing aborted pistils and functional stamens. The absence of stamens has only been observed in one cultivar, and is therefore very rare. The most important anomalies concern the ovary. â&#x20AC;&#x2DC;Ovary abortionâ&#x20AC;&#x2122; refers to the absence of an ovary, or to a small, imperfect, non-persistent ovary. The perfect flower is evidenced by its

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large pistil, which nearly fills the space within the floral tube. The pistil is green when immature and deep green when open at full bloom. Staminate flower pistils are tiny, barely rising above the floral tube base. The style is small and brown, greenish-white, or white, and the stigma is large and plumose as it is in a functioning pistil. All olive trees display ovary abortion, although at different extents, depending on cultivar, year, environment, inflorescence and type of shoot. Its incidence is variable, from less than 10% to 30–40% of the flowers in the most common oil cultivars, up to 50–60% in table cultivars (Table 2.2). In some Italian cultivars, such as ‘Morchiaio’ in Tuscany, up to 70–90% of ovary abortion is reached. In spite of this, production is usually not depressed as normal harvests require no more than 1–4% of fruit set. One hundred per cent abortion cannot exist in a commercial cultivar, but ‘Swan Hill’, an ornamental cultivar selected by Hartmann in Australia (Hartmann 1967), displays that character, which is advantageous when olive trees are used in urban forestry or as ornamentals. Table 2.2 Percentages of ovary abortion for several olive cultivars. Data were collected from trees located in the Chianti region of Italy (Magherini 1971). Cultivar

1961

1962

1963

1964

Average

Frantoio

8.20

9.40

6.20

6.60

7.40

Moraiolo

29.30

18.00

7.20

12.95

16.08

Pendolino

26.20

22.50

31.40

18.45

23.40

Ascolana tenera

49.70

40.40

56.10

49.65

49.10

Uovo di Piccione

57.60

46.40

56.40

46.45

50.66

Morellona di Grecia

32.60

32.00

45.10

32.95

35.12

Grossa di Spagna

34.80

49.60

46.80

86.60

60.88

Sant’ Agostino

36.80

45.40

45.60

64.85

51.50

Gordales Sevillana

49.10

56.60

59.20

69.15

60.64

Yearly averages

36.03

35.57

39.33

43.07

2.2.4 Anthesis and pollination Full bloom occurs in full spring (e.g. May in warm areas such as California, Southern Italy, Greece and Spain; at higher latitudes and elevations full bloom is delayed into the first weeks of June). Differences can be observed among cultivars (Table 2.3), which are to be taken into account when selecting pollinators. Anthesis normally lasts 2–3 days on individual inflorescences, and 5–6 days on the individual tree, or up to 10–15 days if temperatures are relatively low. A flower is fully opened when both anthers and petals are separated. During the hottest part of the day anther dehiscence takes place, and an abundant amount of pollen is shed. The amount of pollen produced appears to be a varietal characteristic; for example ‘Leccino’ and ‘Frantoio’ produce small amounts of pollen, but larger quantities are produced by ‘Ascolana’, ‘Manzanilla’ and ‘Pendolino’. More important is the pollen’s ability to germinate: this characteristic appears to fluctuate (in vitro) between 12% and 60% (Zito & Spina 1956; Fernandez-Escobar et al. 1983).

Flo wer and fruit biolog y in the olive

Table 2.3 Flowering period of olive cultivars in Umbria, hills of Central Italy (Antognozzi et al. 1975). CULTIVAR Uovo di piccione Nocellara etnea Picholine Santâ&#x20AC;&#x2122; Agostino Bouteillan Carolea Santa Caterina Tanche Bella di Spagna Dolce di Andria Dritta di Moscufo Grossa di Spagna Morchiaio Moresca Nebba Rosciola Santagatese Ascolana tenera Bosana Cucca Fecciaro Gentile di Chieti Gordales Grossanne Itrana Manzanilla Passalunara Raja Sabina Casaliva Cellina Carmelitana Cerignola Corniolo Maurino Mignolo Moraiolo Corsini Morellona di Grecia Raza Sargano Taggiasca Ascolana Semitenera Coratina Dritta di Loreto Frantoio Frantoio Corsini Moraiolo Carboncella Nocellara Messinese Ottobratica Piangente Razzola Caninese Correggiolo Pendolino Rastellina Grappolo Ogliarola Olivago Savino Laurina Leccio del Corno Leccino

June 5

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Pollination is influenced by several factors, the most important being: • temperature – which has the effect of enhancing tube growth, although when it is too high the stigma may get dry. For anther dehiscence the optimum is 30°C with 50% RH (relative humidity). A good value of relative humidity also enhances pollen germination; • rain – which is always negative. Indeed, it may determine pollen grain plasmolysis, dilute stigma secretions, and hinder pollen transport; • wind – which is fundamental for this anemophilous species. When too strong it may transport masses of pollen away from the grove. Although olive pollen can be found as far as 12 kilometres from the originating tree, the effective range is up to 20–30 m. Among nutritional factors, good nitrogen fertilisation has been proven to lengthen the effective pollination period, and therefore to improve fruit set. Sterility Sterility may be due to factors different from those affecting ovary abortion, such as anomalies during meiosis producing (i) imperfect gametophytes (cytological sterility), quite rare in olive, and (ii) incompatibility (factorial sterility). Incompatibility occurs when a perfect pollen grain fails to germinate on the stigma, or germinates, but its tube growth is insufficient for fertilisation; this incompatibility may be between two cultivars (inter-incompatibility) or, as is often the case with the olive, a cultivar is genetically programmed not to be fertilised by its own pollen (self-incompatibility). Most olive cultivars are self-incompatible, or self-sterile, and care must be taken to establish orchards with more than one cultivar. It is therefore important to know which cultivars are best suited to fertilise the one we want to be the principal one. In any case, growing three or four cultivars in the same plot guarantees good set, even if the cultivar we are interested in is considered self-fertile, as is the case for ‘Frantoio’. Fruit set In the absence of sterility barriers, the pollen germinates on the stigma, and develops through the style a tube that reaches the ovary and fertilises the egg (‘double fertilisation’). The viable pistil has two carpels, each containing two ovules, but only one ovule is fertilised and develops. Thus, as a rule, in the fruit only one carpel containing one seed is present; occasionally, there may be none (parthenocarpy) or two. Following fertilisation, the flowers shed petals and stamens, the ovary grows in size, and at this point many pistils shrink and drop. Within a month from full bloom only 7–8% of flowers are still on the tree as fruitlets; of them, only 25% are present by late summer. The average fruit set is therefore around 2%, although yearly fluctuations can be wide. Fruit drop is used by the plant to adapt production to its elaborating surface; other factors may influence fruit drop, such as nutritional and water deficiencies, weather conditions during bloom, sterility, lack of pollinators and pests. Fruitlets compete for survival, starting about 10–15 days after full bloom.

Flo wer and fruit biolog y in the olive

2.2.5 Fruit growth and seed development The olive fruit is a drupe, which means it is made of two main parts: pericarp and seed. The pericarp is made of (i) the skin (exocarp), smooth and with stomata, (ii) the flesh (mesocarp), the tissue containing oil, and (iii) the pit, a lignified shell enclosing the seed. The â&#x20AC;&#x2DC;true seedâ&#x20AC;&#x2122; (Fig. 2.8) consists in a seedcoat and a thick endosperm that both sheathe a large embryo made of flat cotyledons with a short radicle and plumule. As a rule there is only one seed, rarely two. In some Spanish cultivars the occurrence of nucellar polyembryony has been reported. Fruit shape and size, pit size and surface morphology vary greatly among cultivars, and are the most reliable morphological features to distinguish between cultivars.

Figure 2.8 Longitudinal section of an olive stone (seed and endocarp) (Krugman 1974).

Fruit growth is represented by the double sigmoid curve, typical of drupes (Fig. 2.9). Three stages can be separated: 1. Exponential growth, which is characterised by abundant cellular multiplication. The pit achieves a quasi-final size. 2. Growth slows down, or stops for a short period. During this stage the embryo completes its development and the pit hardens (sclerification). 3. Growth resumes due to cell enlargement, and gradually diminishes with time.

Olive Propa ga tion

Figure 2.9 Fruit growth of the cv. Lucques, expressed as a growth index (fruit length x diameter). Plants sprinkler (1) and drip (2) irrigated (Villemur et al. 1976).

This growth model is less evident if growth is represented by weight increases, rather than diameter or volume. The curve can even turn into a single sigmoid if dry weight is taken into account, thus showing that stage two is the one requiring most dry weight accumulation. In the course of fruit formation, and tightly connected to it, the vital ovule develops to form the seed. The embryo makes up most of the seed volume. The seedcoat, derived from the integuments which represented the main ovular tissues, is thin and leathery, and rich in vascular ridges. Between the seedcoat and the embryo is a layer of endosperm, rich in starch (King 1938). The embryo has two quite evident large cotyledons, the embryonic leaves (Fig. 2.10). A short radicle, located at the lowest end of the embryonic axis, will give rise to the root system. Between the cotyledons is a small plumule, from which will develop the future epigeic system, the plant parts that will be exposed to the open atmosphere. The embryo starts growing within three to four weeks after bloom, and reaches the globular stage at week six; after eight weeks the cotyledons appear to be well developed. The embryo is usually completely formed after five months from full bloom. No further morphological or anatomical changes appear to occur in the embryo, although dormancy is imposed on the seed late in the season (see 4.2.1). Seed growth means a gradual embryo enlargement, which at the end occupies most of the space inside the endocarp, at the expense of the endosperm.

Flo wer and fruit biolog y in the olive

Usually pollination and fecundation are essential for fruit set and early seed development. The presence of a vital seed in a growing drupe is not essential, as many fruits have their embryos aborted. As a consequence, many apparently normal fruits have no seed. The fruit can also develop without the presence of a fertilised ovule (parthenocarpy), but in this case the fruit remains distinctly smaller.

References Antognozzi, E., Cartechini, A. & Preziosi, P. 1975. Indagine sulla individuazione dei migliori impollinatori per olive da mensa della cultivar ‘Ascolana Tenera’. Proceedings ‘2nd Sem. Oleic. Int.’. Cordoba, 6–17 October. Fabbri, A. & Benelli, C. 2000. Flower bud induction and differentiation in olive. J. Hort. Sci. Biotech., 75 (2): 131–41. Fernandez-Escobar, R., Gomez-Valledor, G. & Rallo, L. 1983. Influence of pistil extract and temperature on in vitro pollen germination and pollen tube growth of olive cultivars. J. Hortic. Sci., 58 (2): 219–227. Hartmann, H.T. 1951. Time of floral differentiation of the olive in California. Botanical Gazette, 112: 323–327. Hartmann, H.T. 1967. ‘Swan Hill’ a new ornamental fruitless olive for California. California Agriculture, 21: 4–5. King, J.R. 1938. Morphological developmentof the fruit of the olive. Hilgardia, 1: 437–458. Krugman, S.L. 1974. Olea europaea L., Olive. In C.S. Schopmeyer (coord.) Seeds of woody plants in the United States. Agriculture Handbook 450. USDA, Washington, pp. 558–559. Loussert, R. & Brousse, G. 1978. L’olivier. G. P. Maisonneuve et Larose, Paris, pp. 465. Magherini, R. 1971. Osservazioni sull’aborto dell’ovario nell’olivo. L’Agricoltura Italiana, LXXI (5): 291–301. Maillard, F. 1975. In Loussert & Brousse, 1978. Morettini, A. 1950. Olivicoltura (1st edn). REDA, Rome. Figure 2.10 Morettini, A. 1972. Olivicoltura (2nd edn). REDA, Rome. Section of a mature embryo Pansiot, F.P. & Rebour, H. 1960. Amélioration de la culture de l’olivier. (courtesy H. Rapoport). FAO, Rome. Rallo, L., Torreño, P., Vargas, A. & Alvarado, J. 1994. Dormancy and alternate bearing in olive. Acta Hortic. 356: 127–136. Roselli, G. 1977. Osservazioni sulla scultura dell’esina del polline di alcune specie da frutto. 1. Olivo. Riv. Ortoflorofrutt. It., 61(2): 157–163. Villemur, P., Gonzales A. & Delmas, J.M. 1976. A propos de la floraison et de la fructification de quelques varieties d’olivier. L’olivier, 16 (3): 45–47. Zito, F. & Spina, P. 1956. Come germina il polline dell’olivo. Italia Agricola, 93(5): 413–425.

3 Propagation by cutting

Plant propagation by cuttings involves all techniques of asexual propagation in which the whole plant originates from a plant part that is put in the conditions of regenerating a root system and a canopy. This definition excludes grafting, in which the final plant derives from two different parent plants. Micropropagation can be considered a particular type of cutting propagation, but due to its technical peculiarity it is dealt with in chapter 5.

3.1 Biology of adventitious root formation in cuttings 3.1.1 Morphology and anatomy With the exception of the root developed by the embryo during germination, all roots formed in a plant are adventitious, as structures similar to buds do not exist in the root system. Thus, all lateral roots are adventitious, as their induction and differentiation in the root pericycle, and their occurrence along the older roots, depend on external factors. Here, attention will be focused only on roots arising from tissues originally destined for other functions, namely branches and shoots. The process of rhizogenesis, and the factors determining the formation of adventitious roots in aerial organs, are fundamentally the same in all the propagation systems adopted for the olive. As multiplication by semihardwood cuttings is the most common system of obtaining new olive plants by asexual propagation, the rooting process as it takes place in such material (that is, one-year-old or younger shoots) will be taken into consideration (Fig. 3.1). Any difference occurring in other systems will be mentioned when dealing with them in the appropriate sections. It is known that adventitious roots forming in cuttings are of two types: preformed roots and wound-induced roots. The presence of preformed roots has never been demonstrated in the olive, although latent meristems occur in old wood which can turn, under appropriate conditions, into roots. This is the case with ovules and sphaeroblasts

Propa ga tion by cutting

Epidermis Cork Phellogen Phelloderm Cortical parenchyma Fibres Sclereids

Periderm or Cortex

Primary phloem

Metaphloem Phloem parenchyma Secondary phloem

Sieve cells Cambium

Xylem rays

Xylem

Xylem parenchyma Vessel

Pith cells

Pith

Figure 3.1 Section of a 1-year-old shoot (Troncoso et al. 1975).

on branches. The roots obtained by propagating semi-hardwood cuttings therefore originate after the cutting is made by excising it from the stock plant. These are called wound-induced roots, as the first stimulus the rhizogenetic tissue receives is that of the wound, but roots also form through a series of modifications in the cutting environment. The term de novo-formed adventitious roots would therefore be more accurate; but, due to the nature and purpose of this book, the use of the term â&#x20AC;&#x2DC;rootâ&#x20AC;&#x2122;, unless otherwise specified, will indicate such roots. The first events occurring in an olive cutting are merely a response to wounding, and are aimed at isolating the organ from the environment in order to avoid water loss through the cut area, which would lead the excised plant part to

Olive Propa ga tion

desiccation and death. Those outer cells that are suddenly in contact with the atmosphere die, and their remnants, often in the form of a layer of mucilages, are a first shield for the inner cell layers. Vessels, which can cause abundant water loss, are eventually plugged with gums (tyloses). Finally, living cells belonging to several tissues deeper below the cut surface start dividing to produce, within the first two weeks, large lumps of specialised parenchyma cells (callus), which in the end constitute a sort of thick cap on the lower end of the cutting. The completion of the process is indicated by a swelling of the cutting base. Its epidermis gets lighter in colour and shows occasional cracks and callus emergence, while the bottom is completely covered by the callus cap (Fig. 3.2). The top end of the cutting is usually sealed more internally, after a dieback of tissues that goes as deep as the loss of water determines. This can at times determine the loss of the most distal leaves and buds, an event that can negatively affect the survival of the cutting.

Figure 3.2 Bottom end of a cutting prior to root emergence.

The process leading to the formation of new roots in cuttings begins within a few days after excision. Specific differentiated cells abandon their physiological and cytological commitment, and regain the ability to divide (dedifferentiation). A lot of research has been devoted to the location of these early cells, although the task is not an easy one; indeed, a number of tissues and positions in the cuttings (phloem, medullary rays, callus, cortex, xylem, even pith) have been indicated in time. Most authors, though, indicate young, differentiating secondary phloem cells, located in the vicinity of

Propa ga tion by cutting

medullary rays, as those most sensitive to the stimuli leading to dedifferentiation. These cells, now meristematic (i.e. able to divide), start dividing and form, within the cutting, lumps of small cells called root initials. In the beginning these do not differentiate into specialised cells, but rather continue dividing until the size of the root initial is such as to exert a pressure on the outer cell layers, the tissues between the initial and the outside â&#x20AC;&#x201C;usually the outer phloem with its schlerenchyma ring, and the cortex. At this point the structure has assumed a conical shape, with its base towards the central cylinder, and is more properly called a primordium. The primordium keeps growing towards the outside, and in the process its cells differentiate into the various tissues of a root; a strong rootcap makes it possible to push and crush the external cutting tissues, and allows the emergence of the primordium. By the time it emerges, the primordium is well structured (Fig. 3.3), and a connection between it and the central cylinder of the cutting is established. Still, the primordium is not yet functional, that is, it is not yet able to absorb water from the substrate. As can be seen in Fig. 3.1, the olive shoot (and therefore the semi-hardwood cutting) develops a ring of fibre bundles in the primary phloem, which becomes uninterrupted by the differentiation of sclereids in the gaps between one bundle and another. This creates a rigid sclerenchyma ring that may appear to be an insurmountable barrier for the primordium, and indeed this has been suggested as the reason for the failure in rooting of some cultivars (Ciampi & Gellini 1958; 1963). Further research, though, has demonstrated that the barrier can be crushed by the pressure of the growing primordium, or that the primordium can skip it by emerging from the bottom end

Figure 3.3 Root primordium (Ciampi & Gellini 1963).

Olive Propa ga tion

(Fig. 3.4), opening its way out through the softer callus tissue. At any rate, the presence of a thick sclerenchyma ring does not account for the varietal differences in rooting percentages (Troncoso et al. 1975; Fabbri 1980).

Figure 3.4 Young adventitious roots a few days after emergence.

3.1.2 Physiology It has been postulated that a substance called rhizocaline, present in leaves, buds and cotyledons, is able to move to the roots and stimulate rooting (Bouillenne & Went 1933) but, in spite of long years of research, rhizocaline still remains a hypothetical compound. However, different classes of plant growth regulators have been demonstrated to influence, positively or negatively, root initiation, including auxins, cytokinins, gibberellins, ethylene, polyamines, abscisic acid, growth retardants and phenolics. To date, auxins have been shown to have the greatest effect on rooting. Numerous reports have indicated the involvement of auxin in the initiation of adventitious roots, and that division of root initials depends on exogenous and endogenous auxin. Synthetic auxins, such as indole-3-butyric acid (IBA) and naphthaleneacetic acid (NAA), have been shown to be more effective than the naturally occurring indole-3-acetic acid (IAA) for rooting. IBA is actually commonly used in propagating olives by cuttings. The physiological role of cytokinins in root initiation and development is an ambiguous one: kinetin (a common type of cytokinin) has shown either stimulatory or inhibitory effects, depending on its concentration in the tissue. At any rate, in tissue culture the cytokinin levels must be lowered when rooting is desired. Roots are an important site for

Propa ga tion by cutting

gibberellin (GAs) metabolism, but these growth regulators do not appear to have a positive role in adventitious root formation. A similar conclusion has been reached for abscisic acid (ABA), although it seems to play an important role in enhancing root growth. A significant amount of research has been devoted to studying the effects of endogenous ethylene in adventitious root formation (Bartolini & Fabbri 1982; Bartolini et al. 1986a; 1986b). Ethylene certainly affects root initiation and development, although it might play different roles in the two stages. It has been suggested that auxin action takes place by enhancing ethylene synthesis. At present, however, the role of ethylene in the rooting process is not completely understood. Many studies have hypothesised a role for polyamines in the rooting process, and their relationship with auxins and peroxidases. According to Gaspar et al. (1997) IAA and putrescine, an important polyamine, might be required to initiate cell division at the end of the rooting inductive phase. Polyamines induce rooting in the olive (Rugini & Wang 1986; Rugini et al. 1997), possibly at the very early stages of rooting. It has also been suggested that polyamines might be considered precocious markers of rooting. At any rate, as other species are not affected by polyamine treatment, it is highly possible that their effects on the olive are due to the low endogenous polyamine content in olive tissues. Other naturally occurring substances that have been shown to exert an effect on rooting are phenolic compounds, which, according to their chemical structure, may be either stimulatory or inhibitory (Bartolini et al. 1988). In the olive, as in most species, juvenile tissues and organs root better than mature ones, and this peculiarity has been exploited for a series of propagation techniques, particularly to produce clonal rootstocks. Such behaviour has been attributed to larger amounts of stimulators and to a reduced presence of inhibitors, a condition that is gradually modified as tissues age. At any rate, although it is exploited when obtaining plants from rooted suckers, the commercial importance of this feature is negligible for the virtual absence of clonal rootstocks in the olive; on the other hand, propagating cultivars from juvenile or rejuvenated material might delay the onset of fruit bearing for an unacceptable number of years. What results in the end from physiological studies on the olive and other species is that a number of substances are needed to trigger the process, and that such substances are for the most part produced by leaves and buds. The process in the olive is particularly time consuming, requiring relatively high amounts of energy, i.e. carbohydrates, possibly in an appropriate C/N (carbon/nitrogen) ratio. This explains the need, when propagating cuttings made from young shoots, to retain as many leaves as possible, regardless of the technique used.

3.2 Techniques of propagation by cutting 3.2.1 Plant material With varying results, every part of the olive tree has been used for propagation. Shoots and branches are, as a rule, excellent propagation material when the proper techniques are applied. The olive tree is also very efficient at differentiating new meristems at the collar, where the transition from trunk to root system takes place, and on old wood (limbs) where latent unopened buds are maintained. This peculiarity explains the relative

Olive Propa ga tion

ease in propagating the olive from large plant parts, which enabled ancient growers to asexually propagate this tree. Root cuttings are the only organs of the olive not able to produce new plants; indeed, roots do not possess adventitious buds, and are incapable of producing such structures when separated from the parent plant. Branches The use of branches more than 4 or 5 years old – originally by the Phoenicians, Romans and Arabs – is a very ancient method of olive propagation. This method makes use of the possibility of producing shoots (from latent or adventitious buds) and roots from branches. Cuttings (20 to 50 cm long, with a diameter of 5 to 10 mm or more) are obtained from branches during the cold season (autumn–winter). The cuttings are kept until spring in a cool and not excessively damp place, in layers of sand. They are then positioned in an outdoors plot in various ways: • horizontally at the bottom of furrows (Fig. 3.5), covered with a few centimetres of soil. In spring, among the new shoots, the most vigorous are kept for growth and the others are eliminated, unless the cutting is of such a size to support more than one plantlet. After 6 to 8 months the plantlets have reached a height varying between 60 and 80 cm, and at their bottom end, near the insertion on old wood, a small root system should be present. The rooted shoots are then separated from the branch, after which they receive normal nursery care; • vertical or inclined. In this case the new shoots do not always produce roots, which can only be present on the old cutting wood. The new plant is therefore made of a conspicuous piece of the original wood (Fig. 3.6). With this method there can be a serious risk of root rot.

Figure 3.5 Branch cuttings positioned horizontally, before covering with soil.

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Figure 3.6 Vertically positioned rooted branch cuttings.

Branch material is easily obtained from pruning operations, which in this case should be made in winter. As a rule, the cuttings should not be left in the open after collection; they should either be put to root or stratified in sand. The soil should be carefully pressed around the cuttings. In present day routine work, treatments with root-promoting chemicals are useful, and rooting performance is directly proportional to stem diameter. Rooting is also obtained from very old trunks, when uprooted and cut into sections 20–30 cm long; the new shoots and roots are produced from the living tissues that are in the cortex when the piece of wood is buried for a few centimetres (Fig. 3.7). Examples of traditional propagation by branch cutting are: • the ‘Cormoni’ (Apulia and Southern Italy in general) – 40 to 60 cm long branches with their foliage removed; they are planted directly in the definitive planting site in late autumn–early winter, at a depth of 35 to 50 cm, with the terminal 5–6 cm emerging; • the ‘Estacas’ (Spain) – big cuttings, as long as 2 to 3 m, more than 6 cm thick, obtained during pruning. About a third of their length is put into the ground (a hole 1 x 1 x 1 m, enriched with manure) in autumn and the rest is covered by a cone of soil, except the upper 20–30 cm. At the beginning of the following summer, when the apical shoots are 5 to 10 cm long, the mound is removed from around the cutting; • the ‘Garrotes’ (Spain) – cuttings 50 to 100 cm in length, and 3 to 5 cm in diameter. For rooting, they are directly planted out (3 or 4 per hole, at the corners of a triangle or square), or are placed in plastic containers with a volume of 20 to 30 litres of soil.

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Good results are obtained with these methods but they are not sufficient to justify nursery use. On a commercial scale, it is almost impossible to find the large quantities of material necessary for this kind of propagation. However, this technique can be useful for the small grower who needs to replant a small percentage of his orchard each year and is not willing to invest in propagation equipment. Shoots The introduction to the olive industry of the misting technique (Hartmann 1952) for shoot rooting has provided and still provides a valid alternative to the techniques used until then in olive-growing countries, namely grafting, ovules and severalyears-old woody cuttings (as described above). The technique uses shoots developed in the same year, or Figure 3.7 Section of old trunk wood, with a few rooted shoots, 1-year-old shoots which have not before excision. produced fruit (see 3.2.2). Ovules Ovules are characteristic swellings (tissue hyperplasia) which appear as protuberances and are found at the level of the collar (Fig. 3.8). At the ovule site, a build-up of sap due to a slowing down of its circulation occurs. This is most often found at the base of the trunk where the root structure joins the trunk, producing torsions in the vessels and thus slowing sap circulation. This determines hypernutrition of cambium cells, which actively proliferate and produce the extroflection of tissues, particularly parenchyma tissues that constitute most of the ovule. In some cultivars these structures can also be found in the lower trunk parts of 5- to 6-year-old trees. Accumulated starchy substances may cause the formation and emission of adventitious shoots and roots, the type of structure depending on whether or not the ovule is exposed to light. This property has been exploited by growers since ancient times to obtain new cloned plants (Fig. 3.9). The use of ovules for olive propagation is a method that was used mainly in the past and in areas at the fringes of olive cultivation. Francolini (1934) observed that olive trees derived from ovules and grown in poor, shallow and sub-arid soils gave better results than trees from grafting. This method has been more recently used in north-African countries such as Tunisia and Libya (Pansiot & Rebour 1960). The ovules can be removed with special cutting tools, and great care needs to be taken of the exposed tissues, which must be disinfected and protected. Planted at the

Propa ga tion by cutting

Figure 3.8 Ovules on an old olive trunk (* = ovule).

Figure 3.9 Detail of an ovule-propagated olive plant (Morettini 1972).

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beginning of spring, ovules are able to produce new trees as they contain both root primordia and dormant buds. Ovules are generally taken from the enlarged conical base of the trunk, the collar, which is particularly developed in old olive trees (Baldini 1986; Hartmann et al. 1990). Their removal from trunks is not common as it damages the stock plant. Ovules are taken from healthy trees at the beginning of spring or autumn, and they are then kept in slightly damp sand until the time comes to put them under earth in the ovulary, or ovule bed. The weight of an ovule varies, on average, from 100 g to 3 kg. If they are rooted in nurseries their weight is usually 500 to 800 g, but when they are to be planted directly in field their weight depends on the expected rainfall and on the possibility of irrigating. If water is expected to be lacking, very large ovules are used (e.g. up to 5 kg in Sfax, Tunisia). The bark of the ovule must be smooth and light in colour, with slight wrinkling which reveals the presence of latent buds, and the wood must have a healthy appearance. Those that are rotten, damaged or already rooted must be discarded. The ovulary (i.e. the site where the ovule is to be planted) is prepared first by turning over the soil to a depth of 80 cm, then the ovules are placed in furrows (20 cm deep) with the cut surface facing down (Fig. 3.10). Planting distances are 60 cm between rows and 30 cm between ovules along the row. Transplanting is done after 3â&#x20AC;&#x201C;5 years; if the new plant has developed its own roots, it can be separated from the ovule. In this case the ovulary can be used as a continual source of plants, as in the stoolbed technique (Battaglini 1967).

Figure 3.10 Details of ovule propagation in the nursery; distances are in metres (Loussert & Brousse 1978).

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Suckers (or pollards) Suckers develop at the base of the trunk and have their origin in ovules. Generally they are taken from the tree when they have acquired their own ovule masses, and have produced sufficiently developed roots (they cannot therefore be considered cuttings in a strict sense). To help these processes, the base of the tree is covered with a thin layer of soil which encourages the initiation of roots. Shoot girdling can further enhance adventitious root formation. In the spring, the rooted suckers are removed from the stock plant with some old, collar wood, to be grown in the nursery before being planted out. Although this method of multiplication can be used for the replacement of small numbers of trees, it cannot be used at nursery level because it is slow and costly. The use of suckers is part of common practice after a frost, to make up for losses in the most convenient way. Once the death of the upper part of the tree is confirmed, it is cut down and removed to avoid the presence of wood that can easily be attacked by parasites (fungi, insects, etc.), while the stump is left so that suckers can develop freely (the stump usually survives the cold because it is covered by a few centimetres of soil). After about one year, the less favourably positioned suckers are removed so that in the second or third year there is one, or at most two or three suckers, from which to grow a new canopy.

3.2.2 Shoot collection and cutting preparation For the preparation of cuttings, scions can be taken all the year round. In practice this is done in relation to the production cycle of self-rooted plants, which generally coincides

Figure 3.11 Time course of adventitious rooting (% of rooted cuttings) versus monthly shoot growth (mm) in the olive. Values are averages obtained by the authors over several years from cultivars of Central Italy, characterised by intermediate rooting ability; months refer to the northern hemisphere.

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Figure 3.12 Good quality shoot for cutting production.

Figure 3.13 Two cuttings were obtained from the shoot in Fig. 3.12. Figure 3.14 Olive semi-hardwood cuttings ready for planting in the rooting bed.

with two annual peak points of concentration of rooting compounds within tissues, i.e. April and September–October in Central Italy (Fig. 3.11). In temperate climates, at the borderline of the growing area for olives (such as Central Italy), it is possible to get shoots from stock plants that are long enough to make more than one cutting, but not more than once a year. From these shoots (Fig. 3.12), cuttings are obtained by dividing them into 10–15 cm long pieces (Fig. 3.13) of 4–6 mm in diameter, with 4 to 6 nodes. The 4 to 6 leaves at the distal end are retained, while the 2 to 3 basal nodes are left without leaves (Fig. 3.14). The basal cut must be made just under a node, and care must be taken to avoid leaving a stub of the internode, which can be a point of necrosis. Generally, new roots do not grow from internode tissues; they usually develop within node tissues. Once

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the cuttings are prepared, basal treatment with a rooting hormone (see below) must be carried out within 20 to 40 minutes.

3.2.3 Auxin treatments to promote rooting of cuttings Auxins are root-promoting chemical agents. In spite of its belonging to the group of the naturally occurring plant hormones, IAA is not commonly used as a rooting promoter in commercial olive propagation. Indeed, in comparison with IAA, the two synthetic growth regulators below have shown to be stronger root promoters, light- and temperaturestable, and more resistant to microbial decomposition. â&#x20AC;˘ IBA is the best growth regulator for general use because it is not toxic to hardwood and semi-hardwood cuttings over a wide concentration range, and is very effective in the root promotion of a large number of plant species. â&#x20AC;˘ NAA is stronger than IBA in terms of stimulation of olive adventitious rooting. However, because uniform results are difficult to obtain with NAA, its use is mainly restricted to species or cultivars which respond unsatisfactorily to IBA. Some commercial preparations are composed of a mixture of IBA and NAA. As for the majority of woody plants, IBA is therefore the best auxin to promote rooting of olive cuttings. Hydro-alcoholic solutions or talc formulations are most commonly used in practising nurseries, as the nursery workers can easily prepare their own rooting treatment at a low cost by buying the IBA as pure product. Alternatively, commercial powder or gel preparations can be used. Using hydro-alcoholic solutions for quick-dip treatments In this method, the cuttings are treated by quick-dipping (5 seconds) their basal parts (0.5â&#x20AC;&#x201C;1 cm) in an IBA hydro-alcoholic solution (Fig. 3.15), previously prepared from a

Figure 3.15 Quick-dip hydro-alcoholic IBA treatment. Prior to treatment, cuttings are tied in bunches for easier handling.

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concentrated stock solution (see box ‘Preparing a stock solution and a hydro-alcoholic solution of IBA for quick-dip treatments of olive cuttings’). The cuttings are then inserted in the rooting medium under misting (Fig. 3.16). Cuttings are most efficiently dipped as a bundle, rather than one by one. The cuttings should be inserted into the rooting medium immediately after treatment.

Figure 3.16 Insertion of cuttings into a perlite substrate.

PREPARING A STOCK SOLUTION AND A HYDRO-ALCOHOLIC SOLUTION OF IBA FOR QUICK-DIP TREATMENTS OF OLIVE CUTTINGS Stock solution (e.g. 100 mL at 50 000 ppm in absolute ethanol) • Weigh 5 g of IBA (acid form) and place in a small transparent glass jar with a 100-mL mark. • Add absolute ethanol and stir until the IBA is completely dissolved. • Bring to 100 mL final volume with ethanol. • Put in a dark glass bottle and store at 4°C (stable up to one year). Hydro-alcoholic solution for quick-dip treatment (e.g. 100 mL at 4000 ppm IBA in 30% ethanol) • Take 8 mL of stock solution and place in a beaker. • Add 22 mL of absolute ethanol and 70 mL of water (to avoid a partial auxin precipitation, use distilled or deionised water instead of tap water). • Pour a small amount of the solution into a flat container, to a depth of 1 cm. • Use immediately after preparation then discard the used solution; 100 mL is sufficient for a maximum of 600 cuttings.

In rooting treatments, the concentration of auxin is usually expressed in parts per million (ppm). In hydro-alcoholic treatments, it refers to the weight of the auxin compound per volume of the solution (w/v), e.g. 1000 ppm = 0.1 g of auxin in 100 mL of solution. However, to test the effectiveness of different growth regulators (e.g. IBA vs. NAA) in rooting promotion, equimolar concentrations (M) of the compounds, and not ppm, should be compared. With reference to the period of cutting collection, effective

Propa ga tion by cutting

concentrations of IBA in hydro-alcoholic solutions are in the range of 2000 to 4000 ppm: 2000 ppm when natural rooting hormone levels are high (cuttings collected from vigorous vegetative shoots of the stock plant), 4000 ppm when natural rooting hormone levels are low (cuttings collected during the rest period or during bloom). With respect to NAA, treatments with this growth regulator are generally in the range of 1000–2000 ppm. In general, ethanol concentrations in the rooting solution should be not higher than 30%, in order to limit the risks of strong dehydration and injury to the basal cutting tissue. As the quantity of IBA that can be dissolved is related to ethanol concentration, 30% ethanol may not dissolve the large amounts of auxin required to produce concentrated treatment solutions (e.g. over 4000 ppm). Salts of some auxin are available, and they can be simply dissolved in distilled or deionised water. Therefore, when high IBA concentrations are required, the potassium (K+) salt formulation can be used, enabling the preparation of 100% water solutions. However, because of the high cost of the K+-IBA compound, this practice is not common in commercial olive nurseries and talc formulations of auxins are preferred (see below). Using talcum powder formulations In root-promoting talc formulations, auxin is dispersed in the inert talcum powder. Here, concentration of auxin refers to weight per weight (w/w), that is grams of auxin per grams of talc (e.g. 1000 ppm = 0.1 g of auxin in 100 g of talc). Auxin–talcum powder mixtures can be purchased commercially (see p. 39), or prepared by the nursery workers using reagent grade auxin and talcum powder. To get a homogeneous dispersion, the auxin is previously dissolved in a solvent, which is then used to wet the talc (see box ‘Talcum powder formulation’). As with the hydro-alcoholic solution, bundles of cuttings are dipped in the powder with their 1–2 cm basal parts (Fig. 3.17). It may be beneficial to pre-wet cutting bases with water so that the powder adheres better. The cuttings are then lightly tapped to remove excess powder. To avoid brushing off the powder during insertion of cuttings in the rooting medium, a stick can be used to make a 5 mm hole in the medium before each cutting is inserted.

Figure 3.17 IBA treatment of cuttings in talcum powder.

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Talc formulations are advantageous in allowing treatments with high auxin concentrations (10 000 ppm or more), which are very useful when propagation of very difficult-to-root cultivars is pursued (Table 3.1). Another advantage is that talc formulations are easy to use. On the other hand, rooting results are less uniform with talc formulations than with hydro-alcoholic treatments, due to the variability in the amount of powder adhering to the cuttings. TALCUM POWDER FORMULATIONS Preparing 200 g of a 7500 ppm IBA stock formulation • Weigh 1.5 g of IBA (acid form) and put it in a small beaker. • Add diethyl ether and stir until the IBA is completely dissolved (alternatively, absolute ethanol can be used). • Weigh 200 g of pure talc and put it in a large container, so that the powder layer is about 1 cm thick. • Pour the IBA solution into the talc and add more solvent while stirring until all the talcum is wet. • Allow the solvent to evaporate completely (ethanol will require more time to evaporate than diethyl ether). • Stir the formulation until the talc again becomes a free-flowing powder.

Table 3.1 Rooting ability of the most commonly cultivated olive cultivars* High (100–66%)

Medium (66–33%)

Low (33–0%)

Cultivar (country)

Aglandau (France)

Aggezi Shami (Egypt)

Azéradj (Algeria)

Arbequina (Spain)

Azapa (Chile)

Bella di Spagna (Italy)

Ascolana tenera (Italy)

Bardhe i Tirane (Albania)

Bianchera (Italy)

Barnea (Israel)

Bella di Cerignola (Italy)

Biancolilla (Italy)

Bouteillan (France)

Bical Castelo Branco (Portugal)

Büyük Topak Ulak (Turkey)

Coratina (Italy)

Bidh el Hammam (Tunisia)

Carrasquenha (Portugal)

Cailletier (France)

Chemlal (Algeria)

Frantoio (Italy)

Çakir (Turkey)

Chemlali de Sfax (Tunisia)

Gordal de Granada (Spain)

Carrasqueño (Spain)

Domat (Turkey)

Leccino (Italy)

Chalkidiki (Greece)

Empeltre (Spain)

Lechin de Sevilla (Spain)

Chemchali (Tunisia)

Farga (Spain)

Cordovil Castelo Branco (Portugal)

Lucques (France)

Erkence (Turkey)

Gordal Sevillana (Spain)

Manzanilla de Sevilla (Spain)

Galega Vulgar (Portugal)

Leccio del Corno (Italy)

Mission (USA)

Kalamata (Greece)

Lianolia kerkyras (Greece)

Mixan (Albania)

Picholine (France)

Nabali Baladi (Israel)

Moraiolo (Italy)

Picholine marocaine (Morocco)

Nocellara Etnea (Italy)

Nocellara Messinese (Italy)

Picual (Spain)

Ogliarola Messinese (Italy)

Oblica (Croatia)

Sigoise (Algeria)

Oueslati (Tunisia)

Pendolino (Italy)

Taggiasca (Italy)

Salonenque (France)

Verdal (Spain)

Verdale de L’hérault (Spain)

Verdeal Alentejana (Portugal)

*The reported percentages are the average of results obtained by different authors, after IBA treatments (for a more detailed information, see Appendix 1).

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Commercial preparations A number of commercial rooting preparations are available, differing in formulations and concentrations. Essentially, they contain one or more auxins in an inert matrix (talc or, less frequently, gel or paste, often mixed with lanolin to increase adherence), plus some minor additives (such as fungicides and microelements). These products are formulated to be effective for a large number of species. Usually, complete directions and a list of plants tested come with each product. However, there is very poor information on the activity of commercial preparations in inducing adventitious root formation in olive stem cuttings, as the olive is rarely among the tested species. The only guideline that can be suggested is to carefully check the composition of preparations, in order to choose the one in which the active ingredient concentration (auxin) is in the range indicated above. Dilute auxin solutions Low-concentration treatments have been occasionally used in olive propagation by putting the hormonal solution into the misting equipment, e.g. twice a week in the evening, after the last watering, for at least one month. With this technique, proper IBA concentrations are in the range of 50â&#x20AC;&#x201C;200 ppm (Bartolini & Fiorino 1975).

3.2.4 Techniques to improve the effectiveness of auxin treatments Propagation of cuttings from many woody species is positively influenced by the application of some techniques that, in combination with auxin treatments, are able to improve rooting rates, the quality of adventitious roots, or both. Although they have never become routine in commercial olive propagation, worthy of note are the practices of soaking and wounding cuttings before auxin treatments. Soaking the cuttings With immersion of their bases in water (Fig. 3.18), cuttings lose by diffusion a number of natural substances, some of which have an inhibiting effect on the rooting process. At the same time, mechanisms which enable the cutting to root occur more readily. Soaking

Figure 3.18 Trays in which cuttings are left to soak overnight.

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does not alter the annual pattern of rooting response of cuttings, it enhances and stimulates an already existing ability. This effect is shown especially in hard-to-root olive cultivars. The control of water pH during soaking can further improve rooting ability. In the olive, a positive effect on both rooting percentage, and quality of roots has been reported when cuttings were soaked for 24 hours at a pH of 8.5 before treatment with 4000 ppm IBA (Fig. 3.19; Bartolini et al. 1977).

Figure 3.19 Comparison of rooting performance of olive cuttings following soaking in water at pH 8.5 (left) and 7.0 (right).

Wounding Basal wounding is often reported as beneficial to stimulate rooting of difficult-to-root species or cultivars. It consists of making thin longitudinal incisions at the base of cuttings with a sharp knife, before auxin treatment. Wounded cuttings display greater absorption of growth regulators at the base of the cuttings, especially when the quick-dip method is used. Following wounding and auxin treatment, callus and adventitious root production are greater, particularly along the edges of the incisions. With regard to the olive, there is no specific information on the subject. Considering the increase of propagation cost due to this practice, its application is justified in the commercial olive nursery only when rooting of very difficult-to-root cultivars is attempted.

3.2.5 Root promoting compounds, used in combination with auxin treatments Numerous compounds, other than auxins, have been tested over time for their ability to enhance adventitious root formation in olive, mainly in combination with IBA treatments. Although to date none of these substances has entered into the common nursery practice, some promising results are mentioned below.

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Growth regulators Cytokinins: treatments with 75–150 ppm benzyladenine (BA) to the leaves of ‘Leccino’ and ‘Frantoio’ cuttings, previously quick-dipped in a 4000 ppm IBA solution, led to the increase of both rooting, and bud bursting (Bartolini & Del Ministro 1981). Gibberellins: unlike BA, the application of gibberellic acid (GA3, at 125–250 ppm concentrations) to the leaves of ‘Leccino’ cuttings, previously quick-dipped in a 4000 ppm IBA solution, produced a strong inhibition of both rooting, and bud bursting (Bartolini & Del Ministro 1981). Ethylene: this is a gas which, in plants, acts as a hormone in many physiological processes. In rooting, it has been reported to affect the induction of adventitious roots, as well as the elongation of preformed or latent root initials (Mudge 1988). In the olive (cv Maurino), basal treatments of cuttings with 10 mM 1-aminocyclopropane-1-carboxylic acid (ACC), the direct precursor in ethylene biosynthesis, in combination with IBA (4000 ppm), showed a positive effect on rooting only when cuttings were treated 1 to 3 hours after their preparation (Bartolini et al. 1986a.) Polyamines: it has been reported that polyamines can interfere in different ways with adventitious rooting of several species (Hausman et al. 1997). In particular, putrescine exerts a stimulatory effect on rooting, unlike spermidine and spermine, which seem to inhibit the process. As regards the olive, putrescine was shown to promote adventitious rooting of cuttings, in synergy with IBA treatments (Rugini et al. 1997). Therefore, putrescine could be considered for use with cuttings of olive cultivars which respond poorly to auxin treatments alone. Fungicides With many woody species, treatment of cuttings with fungicides, whether incorporated into the auxin-talcum powder or used alone, has been shown to protect newly formed roots from fungal attack, as well as increase survival and overall quality of the rooted cuttings. It should be noted that the use of cuttings affected by fungi and insects should always be avoided to limit the spread of diseases, and also because cuttings suffering biotic stresses have low rooting capacity.

References Baldini, E. 1986. Arboricoltura generale. CLUEB, Bologna (Italy), pp. 396. Bartolini, G. & Del Ministro, M. 1981. Influenze ed interazioni di fitoregolatori diversi sulla radicazione e sull’accrescimento dell’olivo in vivaio. Riv. Ortoflorofrutt. Ital., 6: 451–462. Bartolini, G. & Fabbri, A. 1982. Effetto dell’ACC (Ciclopropano-ammino-1-carbossilato) sulla radicazione di talee di olivo. Riv. Ortoflorofrutt. Ital., 66 (5): 377–384. Bartolini, G. & Fiorino, P. 1975. La multiplication par bouture de l’olivier avec la technique du brouillard: 1, Influence du nombre de feuilles, de bourgeons et des traitements foliaires sur l’émission des racines. Ann. Ist. Sperim. Olivicultura, vol. 3: 1–16. Bartolini, G., Fabbri, A. & Tattini, M. 1986a. The effects of regulators of ethylene synthesis on rooting of Olea europaea L. cuttings. Acta Hortic. 179: 841–846. Bartolini, G., Fabbri, A. & Tattini, M. 1986b. The effect of some exogenous growth regulators on rhizogenesis in Olea europaea L. cuttings. Olea, 17: 19–21. Bartolini, G., Fabbri, A. & Tattini, M. 1988. Phenolic acids and rhizogenesis in cuttings of ‘Frangivento’ olive. Olea, 19: 73–77. Bartolini, G., Fiorino, P. & Bouzar, M. 1977. Ricerche sull’influenza dell’immersione in acqua delle talee. 3, Effetto della bagnatura a pH diversi sulla capacità rizogena in talee di olivo, cv ‘Frantoio’. Riv. Ortoflorofruttic. Ital., 6: 409–417.

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Battaglini, M. 1967. Les méthodes traditionelles de propagation de l’olivier. Proc. ‘Sem. Oléic. Int. ‘. Spoleto (Italy), November, pp. 28. Bouillenne, R. & Went, F.W. 1933. Recherches experimentales sur la neoformation des racines dans les plantules et les boutures des plantes superieures. Annuel Jardin Botanique Buitenzorg, 43: 25–202. Ciampi, C. & Gellini, R. 1958. Studio anatomico sui rapporti tra struttura e capacità di radicazione in talee di olivo. Nuovo Giorn. Bot. Ital., 65: 417–424. Ciampi, C. & Gellini, R. 1963. Insorgenza e sviluppo delle radici avventizie in Olea europaea L. ; importanza della struttura anatomica agli effetti dello sviluppo delle radichette. Giorn. Bot. Ital., 70: 62–74. Fabbri, A. 1980. Influenza di alcuni caratteri anatomici sulla radicazione di talee di olivo cv ‘Frangivento’. Riv. Ortoflorofrutt. Ital., 64, (4): 325–335. Francolini, F. 1934. Contributo allo studio sulla moltiplicazione degli olivi. L’Italia Agricola, 71(1). Gaspar, T., Kevers, C. & Hausman, J.F. 1997. Indissociable chief factors in the inductive phase of adventitious rooting. In A. Altman & Y. Waisel (eds) Biology of Root Formation. Plenum Press, New York, 376 pp. Hartmann, H.T. 1952. Further studies on the propagation of the olive by cuttings. Proc. Amer. Soc. Hort. Sci., 59: 155–160. Hartmann, H.T., Kester, D.E. & Davies, F.T. 1990. Plant Propagation, Principles and Practices (5th edn) Prentice Hall, New Jersey, pp. 647. Hausman, J.F., Kevers, C., Evers, D. & Gaspar, T. 1997. Conversion of putrescine to 1-aminobutyric acid, an essential pathway for rooot formation by poplar shoots in vitro. In A. Altman & Y. Waisel (eds) Biology of Root Formation. Plenum Press, New York, pp. 133–139. Loussert, R. & Brousse, G. 1978. L’olivier. G. P. Maisonneuve et Larose, Paris, pp. 465. Morettini A. 1972. Olivicoltura (2nd edn). REDA, Rome, pp. 515. Mudge, K.W. 1988. Effect of ethylene on rooting. In T.D. Davis, B.E. Haissig & N. Sankhla (eds) Adventitious Root Formation in Cuttings. Dioscorides Press, Portland (Oregon), pp. 150–161. Pansiot, F. & Rebour, H. 1960. Amélioration de la culture de l’olivier. FAO, Rome, pp. 250. Rugini, E. & Wang, X.S. 1986. Effect of polyamines, 5-azacytidine and growth regulators on rooting in vitro of fruit trees, treated and untreated with Agrobacterium rhizogenes. Proc. Int. Congress of Plant Tissue and Cell Culture. Minnesota, p. 374. Rugini, E., Di Francesco, G., Muganu, M., Astolfi, S. & Caricato, G. 1997. The effect of polyamines and hydrogen peroxide on root formation in olive and the role of polyamines as an early marker for rooting ability. In A. Altman & Y. Waisel (eds) Biology of Root Formation. Plenum Press, New York, pp. 65–73. Troncoso, A., Valderrey, L., Prieto, J. & Linan, J. 1975. Algunas observaciones sobre la capacidad de enraizamiento de variedades de Olea europaea L. bajo tecnicas de nebulizacion I. Anales de Edafologia y Agrobiologia, XXXIV, (7–8): 461–471.

4 Propagation by grafting

4.1 The purposes of grafting In the olive, as well as in many fruit species, the purposes of grafting can be multiple. • Cloning (asexually propagating) genotypes (cultivars, selections, elite trees) that cannot be propagated by cutting or other methods, or that can only be propagated with such means at exceedingly high costs. Rooting ability differs markedly among olive cultivars (see Table 3.1 and Appendix 1): e.g. several cultivars for table olives are very hard to root or they do not root at all, while other techniques (trench layering, stoolbed layering, etc.) are little suited to the olive. This makes grafting the only viable technique for such cultivars. • Using the properties of certain rootstocks. Several fruit species have available a series of rootstocks able to hasten growth, anticipate onset of production, control plant vigour and improve fruit quality, but research has not produced anything comparable for the olive. • Using the properties of certain interstocks. However, this possibility has not yet been exploited with the olive. • Changing cultivar on established plants (topworking). Differently from other fruit species, the olive does not suffer from sudden changes in the consumer taste that can make a cultivar obsolete in a few years. But topworking can sometimes be necessary to replace a cultivar, at least in part, e.g. when a pollinator appears necessary in a grove where the planting of a pollinating cultivar has not been made, or the pollinating cultivar proves to be unsuitable to the task (because of a lack of overlapping at flowering times, poor interfertility, etc.). • Repairing damaged parts of trees. Occasionally the roots, the trunk, or large limbs of trees are severely damaged by winter injury, cultivation implements, rodents or

Olive Propa ga tion

diseases. Bridge grafting and inarching (Fig. 4.1) are techniques that can repair such damage and save the tree. • Indexing of virus diseases. These diseases are transmitted by grafting, i.e. an infected scion may infect healthy rootstock. Some cultivars display evident and peculiar symptoms of the infection, and are called ‘indicators’. It is therefore possible to test a genotype for the presence of a given virus by grafting it on a seedling of an indicator plant. As the first virus diseases of the olive have been recently described, it is possible that this technique will also gain importance in the near future for this species. • Research purposes. Grafting is a useful tool for studying a series of physiological aspects of olive biology, but they cannot even be outlined here.

Figure 4.1 Multiple inarching of a damaged olive tree: situation after five years (Wuhan, China).

Propa ga tion by grafting

On the other hand, grafting has some disadvantages: • The ‘seed to orchard’ procedure is far more lengthy and laborious than the ‘cutting to orchard’ one; hence, grafting propagation is far more expensive than cutting propagation; • Seedlings used as rootstocks are a genetically varied population of individuals, and their effects on the grafted cultivar may likewise be varied. This drawback can be overcome to some extent in the nursery by sorting out the seedlings. It can also be remedied after planting by inducing the scions to root and therefore obtaining trees on their own roots. This procedure requires planting by placing the grafting point below the soil level. However, this practice is uncommon in modern olive culture. • In many olive cultivars the germination of healthy and perfect seeds may be heavily delayed, as they must overcome a strong double dormancy (see 4.2.1); in addition to this, the germination percentages of several cultivars are very low. • Grafting is a complex operation which requires long practice. As a consequence, the production of grafted trees is usually restricted to specialised nurseries, where skilled labour is present. Grafting of fruit species can be carried out in many ways (see box ‘Terminology of grafting propagation’), and therefore the number of grafting procedures described in the scientific literature is countless. Not all of them have been adopted internationally, and only the most common procedure used in Mediterranean olive nurseries is described here.

TERMINOLOGY OF GRAFTING PROPAGATION Grafting means to put in contact two portions of tissue or two organs coming from two different trees, in such a way that they will heal and subsequently grow as one composite woody plant.The upper or distal part of this union (the scion) is a short (even just one node) piece of shoot carrying a number of dormant buds. After the healing of the graft union, it will develop into the canopy of the new plant. The plant from which the scion is obtained (the stock plant) should be tested for both its genetic response to the desired cultivar, and its good sanitary condition.The lower or basal part of the graft (the rootstock) will develop into the root system of the new tree. In the olive, it may be a seedling, a rooted cutting, a rooted sucker or ovule, or a micropropagated plant. With the exception of the seedling, propagation techniques give rise to clonal rootstocks, uniform and constant in their characteristics. A particular form of grafting is topworking, in which the rootstock may extend to the whole trunk and to the main scaffold branches (see 4.3.4). Budding is a form of grafting, where the scion is a small portion of tissue carrying one bud. Budding can be undertaken on olives, but it is not the favourite method for propagation.The callus is a mass of parenchyma cells, produced by the meristematic cells of the cambium (the regenerative tissue located between the bark and the wood) or by dedifferentiated cells, usually around wounds. In grafts, callus tissue proliferates to fill all empty spaces between rootstock and scion, determining the healing of the graft point.

Olive Propa ga tion

4.2 Production of olive seedlings Seed propagation in the olive is undertaken mainly for the purpose of producing seedlings to be used as rootstocks for olive trees. The advantage of seed propagation lies mostly in the possibility of producing a very large number of high-quality virus-free rootstocks. Moreover, the seeds are cheap and the production of seedlings can be carried out with little skill and equipment. One important drawback of rootstock production by seed is that the seedlings are heterogeneous in terms of vigour and root development, hence influencing growth characteristics of grafted plants which can differ quite markedly. A proper handling of olive seeds – from fruit collection up to seed germination and seedling development – is fundamental to produce fast-growing homogeneous rootstocks to be used in grafting propagation. The method of grafting scions on seedlings is still largely used in Italy where, in the highly specialised nurseries of Pescia (Pistoia province, Italy), the large numbers of plants required for establishing the new industrial olive plantations have been produced over the last half century. This would not have been possible if only branch cuttings and ovules had been available. The method is also used widely in Argentina, where it has allowed a rapid diffusion of the species. On the other hand, nurseries of other important olive-growing countries such as Spain almost exclusively use stem cuttings for olive propagation.

4.2.1 Stone collection and quality of olive seeds The use of high-quality seed is of prime importance for rootstock production, whether growers collect or produce the seed themselves or obtain it from others. As described above (see 2.2.5), what is commonly called ‘olive seed’ is actually a stone, i.e. the ‘true’ olive seed is enclosed in the fruit stony endocarp (the pit). Olive stones can be considered of good quality when (i) they are genetically true-to-type, and represent the desired olive cultivar or provenance, (ii) they are clean and free from disease and insects, and (iii) the embryos that they contain are viable and capable of high germination. Both wild olive trees (oleasters) and cultivars have been used in the past as sources of stones. A greater percentage of germination and more vigorous seedlings are obtained from the former. On the other hand, seedlings from wild olive stones are often non-cold-resistant, highly heterogeneous in growth, and have few tap roots which are invariably damaged during transplantation, so that rapid recovery is not possible. Moreover, derived seedling rootstocks have very thin bark and very short internodes, making grafting difficult. For these reasons, stones of cultivars are now preferred for propagation in olive nurseries. Olive seeds, both from wild trees and cultivars, are always affected by seed dormancy, for which embryo germinability immediately after fruit ripening is almost nil. Olive seed dormancy In the olive, when the fruit are ready for harvesting, embryo germinability is impeded by a double form of primary dormancy (see box ‘Seed dormancy’): a ‘mechanical dormancy’ due to the stony endocarp of the fruit which does not allow embryo imbibition and expansion (seed-coat dormancy), and a ‘chemical dormancy’ induced by some

Propa ga tion by grafting

germination inhibitors, i.e. naturally occurring chemicals which are presumably located in the endosperm and in its cover (testa). Experimental trials have shown that embryos excised from their seed coats and cultivated in vitro initiate promptly to germinate (Lagarda et al. 1983a; Acebedo et al. 1997), showing that the embryo by itself is not affected by any forms of dormancy. The seeds of many cultivars require up to 4 years for a complete removal of seed dormancy (Voyatzis & Pritsa 1994). Indeed, the seeds from just harvested fruits show nil or very low germinability (see box ‘Germinability’). During their storage, there is a progressive, although slow, natural overcoming of dormancy, so that the germinability increases and reaches a maximum, depending on the cultivar, between the 2nd and 4th year. Then, it gradually decreases in the following years. Further significant, though not generalisable, information on olive seed dormancy and germinability is available, among which (i) a 100% germinability of ‘Arbequina’ is obtained after the seeds are removed from the stony endocarps and maintained in tap water for 30 days (Sotomayor León & Durant Altisent 1994), (ii) the overcoming of dormancy can be promoted by the exposure of seeds to constant temperatures, e.g. 13°C for ‘Picholine’ (Instanbouli & Neville 1977), 15°C for ‘Manzanilla’ seeds (Lagarda et al. 1983b), 10°C (3–4 weeks) followed by 20°C for ‘Chondrolia Chalkidikis’ (Voyatzis 1995). Sources of stones and quality of seeds The best sources of stones for seed propagation are trees of specific cultivars, selected from olive orchards with plants in good conditions of productivity. When possible, it is advisable to select plants of the same cultivar in the inner part of the orchard, in order to reduce cross-pollination as much as possible and, therefore, heterogeneity of seedlings. Once trees producing good quality seeds have been located, it is recommended that the stones are collected every year from the same plants. The choice of cultivars used as sources of stones for seedling rootstock production generally results from the personal

SEED DORMANCY A seed separated from the plant may display primary dormancy, which prevents immediate germination and to various extent delays germination, until given environmental events have occurred. In nature, different kinds of primary dormancy have evolved, two of them concerning olive seeds: • ‘mechanical dormancy’ (or ‘seed coat dormancy’) depends on the presence of seed coats which exclude water and air penetration, and are too strong to allow embryo expansion during germination. Note that in the olive the term ‘seed coats’ refers to both the coats which cover the embryo (testa and endosperm), and the seed enclosing structure (the pit, formed by the stony fruit endocarp). Impermeability and hardiness of seed coats are due to a layer of macrosclereid cells, especially thick-walled on their outer surfaces and coated with a layer of waxy, cuticular substances. Hard or impermeable seed coats, which in nature are softened by the action of microorganisms in the soil, can be mechanically abraded or broken, or softened by means of treatments with acid or basic compounds. • ‘chemical dormancy’, due to chemical compounds which accumulate in embryo coverings (endosperm, inner seed coats) during the development of the seed and act as germination inhibitors, even for a long time after harvesting. Excising the embryo from the seed covering can overcome chemical dormancy.

Olive Propa ga tion

experience of nursery operators, who have acquired, with time, information on seed germinability and superior characteristics of derived seedlings in terms of vigour, quality of root apparatus, diameter reached at grafting time, and tolerance to biotic and abiotic stresses. Examples of cultivars used as stone sources are ‘Canino’, ‘Mignolo’, ‘Maurino’, ‘Moraiolo’, ‘Frangivento’ and ‘Americano’ in Italy; ‘Arbequina’ and ‘Verdal’ in Spain; ‘Allegra’ and ‘Oblonga’ in the USA; and ‘Frantoio’ in Australia. A limited use of stones from wild olive trees is still sporadically made in Italy and Spain. Olive nursery practice has repeatedly shown that different results, in terms of seedling production, are obtained using seeds from small- or large-fruited cultivars. As a rule, there is a strict correlation between fruit and stone size (Table 4.1). Seedlings derived from ‘small’ stones have a greater proportion of taproots than those grown from ‘large’ stones, whose root systems are more reduced and branched. As it is considered that seedlings derived from ‘large’ stones are more precocious and can be grafted 10 to 15 days earlier, it should follow that these seeds have generally better characteristics and their use should be preferable. On the other hand, small stones, although of lower productivity, display a higher and more rapid germination. As a consequence, nursery operators prefer to use small-fruited cultivars (such as those listed in Table 4.1) as donor plants. This orientation is also favoured by the lower costs of small stones. GERMINABILITY Germinability refers to the quickness and the vigour with which embryos start to germinate when the seeds are placed in proper environmental conditions. Several parameters have been proposed to express seed germinability, among which the following are the most used: • the germination percentage (or germination capacity), which indicates the percentage of seeds which are able to germinate and to produce seedlings within a specified length of time. • the germination rate (or average time of germination), which expresses the average number of days required for seed germination. This can be calculated as follows: GR (days) =

N1T1 + N2T2 + … + NnTn total number of seeds germinating

where N values are the numbers of seeds germinating within consecutive intervals of time, and T values indicate the days between the beginning of the test and the end of each subsequent interval of time. Although these two parameters give a good indication of seed quality, growth rate and the morphological appearance of seedlings must also be taken into consideration.

Germination percentages are quite variable, according to cultivar, technique, environment and year. As regards this aspect, research is quite scarce and, in addition, individual experiences (e.g. Scaramuzzi 1957, 1958; Guerrero 1997) are seldom comparable, just like nursery operators’ claims. As a rule, olive seeds are credited with poor germinability, although it varies greatly among cultivars (ranging, with a large approximation, between 5% to 60%). Low germination percentages are mainly due to embryo abortion and mechanical/chemical dormancy of embryos; these latter problems can be overcome by means of techniques such as soaking, scarification and stratification of stones (see 4.2.2).

Propa ga tion by grafting

Table 4.1 Classification of fruit and stones of olive by their size Fruit/stone size

Fruit weight (g)

Fruits per kg of fruits (n°)

Stone weight (g)

Stones per kg of stones (n°)

Small *

>500

<0. 5

>2000

Medium **

2–4

250–500

0. 5–1. 0

1000–2000

Large ***

<250

>1. 0

<1000

* (e.g. ‘Americano’, ‘Arbequina’, ‘Koroneiki’, ‘Mignolo’) ** (e.g. ‘Frantoio’, ‘Leccino’, ‘Mission’, ‘Picual’) Medium *** (e.g. ‘Ascolana tenera’, ‘Gordal sevillana’, ‘Teffahi’)

As mentioned above, the choice of the cultivar from which to obtain seedling rootstocks should take into consideration (other than seed germinability) aspects related to the quality of seedlings, among which are (i) the characteristics of the root system: e.g. cultivars with a root apparatus that grows deeper should be preferred when the rootstocks will be used in sandy soils, and (ii) the speed of growth of the seedlings. It is fundamental that at grafting time – about one year after the transplanting of germinated seeds in the ‘grafting area’ or in pots (see in 4.2.2 ‘ Transplant and growth of seedlings’) – the seedlings have a stem thickness, at about 10 cm from the soil, of at least 5 mm, in order to properly support the grafted scions. Many cultivars reach this diameter over a longer time, and therefore a greater number of seedlings have to be discarded at grafting time. Among the best cultivars for this aspect are ‘Canino’, ‘Mignolo’, ‘Cellina di Nardò’ and ‘Maurino’. Harvesting time In general, the best fruits for seed propagation are those harvested at their full ripening, when the peel and the pulp have the same colour. This condition is reached between middle autumn and early winter (November to January in the northern hemisphere), depending on the cultivar and the climatic characteristics of the growing area. Progress in olive fruit ripening coincides with total carbohydrate accumulation in the seed, and this parameter seems positively correlated with germination ability (Rinaldi et al. 1994). Moreover, an anticipated collection, at the moment of colour change of the fruit, increases the percentage of empty seeds (seeds not containing embryos), as the fruits with empty seeds fall down during ripening. The percentage of empty seeds at ripening is generally in the range of 10–30%. Extraction, cleaning and storage of stones After fruit harvesting, a small quantity of stones can be obtained by hand separation. The olives are opened by cutting the pulp with a knife and the stones are simply scooped out. When large quantities are required, olives are soaked in water for one week or more in order to soften the pulp, after which the stones can be separated following different approaches: • by using special de-stoning machines, which separate the stones from the pulp parts, dirt and other debris • by using a grape shredder, appropriately adapted to be used with olive fruits

Olive Propa ga tion

â&#x20AC;˘ by mixing the olives with an equal quantity of dry oil-cake or sawdust and using the grinder of an ordinary mill Whatever the technique applied to extract the stones from the olives, it is fundamental to keep the stony endocarp intact to avoid damage to the embryos. After extraction, the stones undergo a careful cleaning to remove all flesh residues. A traditional system of cleaning is performed using a concrete mixer (or similar devices), where the stones undergo a mechanical stirring (about 10 min) in a 20% (w/v) sodium hydroxide (NaOH) solution (5 L of solution per 100 kg of stones). Alternatively, calcium hydroxide (CaOH) can be used. The stones are then thoroughly washed to remove any fleshy remnants and disinfected with TMTD 50 (active principle: Thiram at 49% concentration, 300 g per 100 kg of stones). The stones can be dried either naturally in open air if the humidity is low, or artificially with heat or other devices. Cleaned and dried stones (Fig. 4.2) can be stored in cool and ventilated rooms, arranged (i) in 5â&#x20AC;&#x201C;10 cm layers, over grilles that facilitate ventilation, (ii) in jute bales, or (iii) stratified with sand. Alternatively, they can be stored in refrigerators, at about 10Â°C. The stones are generally stored in this way for a minimum of 10 months, during which there is a partial overcoming of seed dormancy. Olive stone storage can be prolonged for up to 3 years with no significant loss of germinability.

Figure 4.2 Olive stones, after extraction from the fruit and cleaning.

Propa ga tion by grafting

4.2.2 Technique of propagation by seed In the olive nursery area of Pescia (Central Italy), mid-summer is by tradition considered the best period to initiate seed propagation, using the stones collected the year before and stored as described above. Before sowing, it is worthwhile carrying out some operations aimed at increasing the germination percentage and at synchronising, as much as possible, seed swelling and embryo development (for a better comprehension of the operation chronology, see box ‘Example of a complete procedure for seedling rootstock production’). Pre-sowing treatments Olive seed germination is hastened by the application of treatments aimed at softening the stony endocarp before sowing. One of the following practices is generally applied: • soaking of stones, performed by placing them in frequent changes of water (e.g. every 1–3 days for a period of 10–20 days). During this process, floating seeds that are empty must be discarded. • chemical scarification, which consists of placing the stones in containers and covering them with 5% w/v sodium hydroxide (NaOH) in water for 24 hours. The mixture should be stirred cautiously at intervals during the treatment to produce uniform results, and the progress of the treatment should be monitored by periodical drawing out of samples to check the thickness of the seed coat. At the end, the sodium hydroxide solution is poured off and the stones are washed for about a week, with the water renewed daily. • mechanical scarification – for large quantities of stones the technique requires suitable equipment (scarifiers); alternatively, the stones can be combined with coarse sand or gravel and tumbled in a concrete mixer. It is essential that

Figure 4.3 Schematic representation of stratified olive stones in moist sand (Baldini 1986).

Olive Propa ga tion

scarification is stopped before the point at which the true seeds start to be injured by the mechanical abrasion. Mechanical scarification is of little application to olive propagation. Regardless of the pre-sowing operation, better germination results are obtained when followed by a 2-month stratification of stones in moist sand (Fig. 4.3), mixed with a fungicide (e.g. TMTD 50, 300 g/hl), in a ventilated and shaded area in the open. The layer of stones and sand can be covered with sackcloth (such as jute sacks) and kept lightly damp. At the end of this period of stratification, the right moment of seed sowing is indicated from the following observations: (i) the pit colour has changed from light to dark brown, (ii) the inner seed coat has changed from dark brown to light green, and (iii) the cotyledons have just started to swell. Just before sowing, a high proportion of empty seeds can be eliminated by discarding any stones floating on the surface of water. Sowing Sowing is undertaken in cold frames (Fig. 4.4), germination benches (Plate I) or simple plastic trays like the ones used for fruits and vegetables (Plate II), under a glasshouse or plastic tunnels. Alternatively, sowing can be done in the open on welldrained nursery soil, in rows which are mulched during the winter months (this system is quite common in Californian olive nurseries). A good olive seedbed should have a loose but fine physical texture, delivering adequate moisture to the seeds and good but not excessive aeration. The subsoil should have good drainage and aeration, and the surface soil should be free of clods and clay, in order to avoid the formation of a crust which can produce ‘knelt seedlings’. The ‘knelt seedling’ has a marked bending of the stem, due to the resistance of the earth’s crust to seedling emergence. Strong stem Figure 4.4 Cold frame for olive seed germination (Pescia, Italy). The lid (framed polythene) is lifted during warm days. bending is long-lasting, and

Propa ga tion by grafting

determines a slackening of post-germination growth; as a consequence, ‘knelt seedlings’ are discarded because they cannot reach adequate stem thickness at grafting time. For the above, olive seedbeds are prepared with three layers as follows: • surface soil: 1 cm layer of river sand • middle: 1–2 cm layer of stones • subsoil: 15–20 cm layer of sand or pumice or a mixture of these. The quantity of stones required is 1.5–3 kg per m2 of seedbed, depending on the size of the stones (see box ‘Calculation of quantity of stones required for sowing’). A precautionary fungicide treatment of the substrate (e.g. with TMTD 50 or Captan, 250 g/hL) is used, to be repeated at least monthly during seed germination in order to avoid or to reduce collar rots due to Pythium and/or Rhizoctonia contamination. Moreover, a wire net around the seedbed should protect the stones from rodent attacks. A good watering of the seedbed makes the soil stick to the seeds. Darkness is not required for olive seed germination. However, in the summer, shading the seedbed during the day can be necessary to avoid excessive temperatures. In autumn, about two months after sowing, seed germination begins and continues through early winter (Plate III). Some nurseries use warm beds and glass panels to stimulate seed germination throughout winter, as well as to protect seedlings against sudden temperature drops. More recently, electric blankets have been placed at the bottom of the seedbed to improve germination. Although the results of this innovation are very interesting, the system is still unusual because of its high cost. At the end of winter it is common to observe a second flux of germination, due to those seeds which had a deeper dormancy; however, many of these late seedlings are choked by the seedlings developed in autumn. Depending on the cultivar and on several other factors (such as seed quality, propagation technique, seed losses due to contamination, rotting, rodents, and knelt seedlings) a good production of seedlings in Pescia nurseries is considered to be in the order of 40–60% of the harvested seeds. For instance, the ‘Mignolo’ is a cultivar with a high proportion of empty seeds; therefore, its final seedling production is generally no more than 50% of the harvested seeds, as about 30% are empty seeds and about 20% are non-germinating seeds or seeds lost during the first stages of germination. CALCULATION OF QUANTITY OF STONES REQUIRED FOR SOWING Determination of stone quantity required for sowing is critical to produce a desired seedling density, which in olive seedbeds should be in the order of 1500–2000 seedlings/m2. Weight of stones (WS) per m2 can be calculated as follows: WS (kg/m2) =

seedling density (n°/m2) NS x GR

where NS is the number of stones per kg of stones, and GR is the germination percentage. GR is expressed as a decimal value. The calculated rate is a minimum and should be adjusted to account for expected losses in the seedbed, as a result of experience in the specific nursery.

Olive Propa ga tion

Germinating small quantities of valuable seed A rapid propagation of a very small quantity of valuable seed (coming, for instance, from cross-pollination, elite trees, etc.) is sometimes necessary. In this case, very high seedling yield can be obtained if olive seeds are removed from the pits by gently using a nutcracker or a small vice. The seeds are then germinated in Petri dishes on filter paper moistened with sterile water. To reduce seed contamination, it is better to treat the seeds with a disinfectant, such as a 2% solution of sodium hypochlorite (NaOCl) for 20 minutes followed by several washes with sterile water, and to operate under the sterile conditions of a laminar-flow hood. The germinability of seeds with a strong chemical dormancy can be stimulated further by treatments with an ethylene-promoter (such as ethephon at 200 mg/L concentration; Lambardi et al. 1994) or cytokinins (such as thidiazuron at 10 ÂľM concentration; Rinaldi & Lambardi 1998). Germinating seeds of other Olea species The seeds of Olea cuspidata are used in some countries (e.g. India and China) for the production of olive rootstocks (Fig. 4.5). Over 70% germination can be obtained if, before sowing, the stones undergo 5â&#x20AC;&#x201C;10 min of chemical scarification with pure sulphuric acid (H2SO4), or are treated for five hours with 3% w/v sodium carbonate (Na2CO3) followed by a 6-hour immersion in 0.5% w/v potassium hydroxide (KOH; Vachkoo et al. 1993). Other experiences show the possibility of easily germinating seeds from Olea cuspidata subsp. asiatica and Olea cuspidata subsp. ferruginea, if they are sown soon after the harvesting of mature fruits and the removal of their endocarps (Legesse 1993).

Figure 4.5 Seedlings of Olea cuspidata from China.

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Transplant and growth of seedlings In the spring after sowing, when the seedlings have 4–6 month growth and are 10–15 cm tall (Plate IV), they have their roots slightly trimmed and are transplanted into wellworked and manured soil, or in pots (2.5 litres; Plate V) filled with a common olive substrate (see 6.3). To obtain uniform plants, seedlings should be selected at this time, with the thin, the knelt and the contaminated ones being rejected. The ‘grafting area’ is a specifically dedicated plot of the nursery where the transplanted seedlings, in pots or in the soil, are grown up to grafting time. When in soil seedlings are arranged in about 1 m wide strips, slightly lower than the earthen alleys (Plate VI) to facilitate surface irrigation and the covering of the plot with mats. Plant density is about 150 units/m2. By the end of the year, the young plants are 30– 50 cm tall, and have the thickness of a pencil at collar level. In the following spring (i.e. 18 months after sowing, one year after transplanting in the ‘grafting area’; see box ‘Example of a complete procedure for seedling rootstock production’) the plants are ready for grafting (Plate VII).

4.3 Theoretical and practical aspects of grafting 4.3.1 Histology of graft union Many studies have been undertaken on graft union formation in herbaceous and woody plants, but so far very little attention has been devoted to the anatomy and physiology of grafting in the olive. At any rate, the process by which a graft ‘takes’ is the same in all species, although minor differences may arise due to the structure of the tissues in contact, which may to a certain extent differ among species (‘takes’ means that the grafted point shows good healing, with evidence of the re-establishment of the vascular connections between the scion and the rootstock). Just as the formation of completely new meristems is necessary for the formation of all adventitious structures, such as buds and roots, the healing of the two structures to form a new individual also requires the formation of a meristematic area, more or less extended between scion and rootstock. The parts of the graft that are originally prepared and placed in close contact do not themselves move about or grow together. Rather, the union is accomplished entirely by cells that develop after the actual grafting operation has been made. The graft union is formed by callus cells that, in the first days, divide quite rapidly. These cells are produced by both structures, although in many instances the scion has been shown to be more active in this stage. Once this cell mass has filled the empty spaces between scion and rootstock, or has at least sealed any possible opening towards the external environment, and put the two structures in contact, within it an array of cells starts differentiating into a vascular cambium, which in turn will produce a vascular system that will be the system of the new individual. In the olive, callus production starts within the first 48 hours (Fontanazza & Rugini 1983), and is abundant by the 7th day; the processes described above take place in the following weeks, and complete vascular continuity is considered achieved by the 60th day under good working and environmental conditions (Fig. 4.6). In short, the major events leading to a successful graft are: adhesion of the rootstock and scion, proliferation of callus cells at the graft interface or callus bridge, and vascular differentiation across the graft interface. The scion will not resume a lasting growth

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Figure 4.6 Healing of a wedge graft, 60 days after grafting (Fontanazza & Rugini 1983). The empty gap is isolated from the outside by callus and new vascular tissue is being formed (arrow).

unless a vascular connection has been established, in order to obtain water and mineral nutrients from the root system of the rootstock. In an analogous way, the rootstock will degenerate or will be forced to develop its own buds if phloem transport is inhibited by lack of connection with the scion, which, by developing, is supposed to send carbohydrates and other metabolites downwards. In addition, the scion must have at least one terminal meristematic region, i.e. a bud, to resume shoot growth, and eventually to supply photosynthates to the root system. Some additional information may be useful in order to better understand and carry out a grafting operation: • For successful grafting the cambial zones of both scion and rootstock should be ‘matched’, i.e. they should be as close as possible. While this is rarely a problem in grafts in which the bark is lifted and ample areas of scion bark are put in contact with similar rootstock parts (such as in feather grafting, which is a type of crown grafting, or in shield budding), difficulties can arise with wedge grafting, as the diameter of the two structures may be different, and lining up of the two cambia could be hard to achieve. In this case it is suggested that the inclination of the scion is varied by a few degrees, to make sure that at least in one point the two generating zones are in contact. • The grafter must make sure to hold the two original graft components firmly together (by wrapping, tying, nailing, etc.); hence, the two parts will not move

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about and dislodge the interlocking parenchyma cells after proliferation has begun, and at the same time the pressure will make empty spaces smaller and more easily filled by callus tissue. It is also suspected, although not definitively proven, that pressure may somehow positively influence the differentiation processes within the callus. EXAMPLE OF A COMPLETE PROCEDURE FOR SEEDLING ROOTSTOCK PRODUCTION The procedure summarised here is applied in an Italian nursery (‘Azienda Pagni’, S. Lucia di Uzzano – Pistoia, Italy) with a long tradition in the production of olive seedling rootstocks. It is a good example of olive seed propagation from fruit harvesting to the grafting of seedlings. 1st year, autumn Olive collection from small-fruited cultivars (such as ‘Canino’, ‘Mignolo’, ‘Americano’)

De-stoning in a grape shredder and stone cleaning by stirring in a concrete mixer with 20% NaOH solution (5 L per 100 kg of stones), after which the stones are thoroughly washed with water to remove any fleshy remnants

Storage of stones in cool, ventilated rooms, stratified over mesh in 5–10 cm layers

2nd year, late spring Stone hydration by soaking for 20 days in water, renewed every 3 days

summer Two-month stratification of stones in the open, in moist sand plus a fungicide (TMTD 50, 300 g/hL); the layer of stones and sand is covered with jute sacks and kept lightly damp

late summer/early autumn Removal of empty seeds by discarding stones floating at the water surface

Sowing in germination benches, under plastic tunnels

autumn 1st flux of seed germination

3rd year, late winter 2nd flux of seed germination

spring Transplanting of seedlings in the ‘grafting area’

4th year, spring Grafting

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• The environmental conditions occurring during and following grafting may be fundamental in regulating the success of grafts. Temperature has a marked effect on the production of callus tissue. The best temperature is considered to be in the range of 25–35°C. Below this callus production slows down; if above it can be faster, but the rate of reserves and water depletion can be too high, and lead to failure. Grafters should therefore make sure to operate in good environmental conditions, by selecting the right season, and using, according to the need, shelters, greenhouses, shading, heating, etc. Topworking may require subsequent protection of the graft. The same type of care also regulates the other fundamental factor, moisture. A graft exposed to very dry air without protection can dry very quickly; hence, the wrapping with various materials of the graft zone, or waxing, or both. Lack of moisture in the soil may also be detrimental, slowing callus production. The adjoining cut tissues of a completed graft union must therefore be kept at a very high humidity level, and the rootstock must be regularly irrigated.

4.3.2 Graft incompatibility There appears to be no incompatibility among Olea europaea cultivars, and the same applies to cultivars grafted on oleasters (Olea europaea L. subsp. oleaster Hoffm. et Link), i.e. seedlings from wild plants growing in now remote areas of the Mediterranean maquis. The olive can be grafted on other species of the genus Olea, and on other genera belonging to the family Oleaceae. Rootstocks can be obtained from trees or shrubs of the genera Phillyrea, Ligustrum, Chionanthus, Syringa, Fontanesia, Forsythia. The olive has been successfully grafted on Olea chrysophylla (Tallarico 1939), Syringa vulgaris (Casella 1932–1934a, 1932–1934b), Forestiera durangensis (Calvino 1930), Fraxinus ornus (Occhialini 1906); no compatibility appears to exist with Phyllirea angustifolia (Milella 1962). Rootstocks of the above-mentioned genera have actually reduced the size of the olive plants, but graft compatibility is limited and within a few years the plants deteriorate (Carocci Buzi 1945). In Asia, in the pre-Himalayan belt, from Pakistan to China, where Olea cuspidata Wall. (= Olea ferruginea Royle) is indigenous, this species is used as a rootstock for the olive. The grafted cultivars have shown a partial incompatibility (swelling of the trunk above the graft), which did not influence plant survival. Fairly old producing plants are common in those countries.

4.3.3 Grafting on seedling rootstocks The olive can be successfully grafted with all grafting and budding techniques. Therefore, in the regions where olive cultivation is a new crop and there is no tradition, and where grafters have developed a particular skill with a given grafting procedure on other fruit species, such skill can be applied to the olive with no great problems, although a minimal testing of techniques to discard visibly bad practices should be carried out with time. Indeed, healthy olive trees, which were originally grafted with various techniques (e.g. whip and tongue grafting, side veneer grafting, saddle grafting, wedge grafting, crown grafting, patch budding, ring budding, chip budding), can be observed in several parts of the world. Nevertheless, only the procedure relating to a type of bark grafting, the ‘innesto a penna’ (‘feather grafting’), is described here, as this is the type of grafting characteristic

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of Pescia (Italy), which has been for a century, and still is today, the world’s most important centre for olive propagation. Pescia nursery operators have tested and perfected this grafting procedure over the years, and the result therefore derives from widespread experimentation. This means that bark grafting is both biologically suitable, and economically viable, in terms of costs, take, etc. Wherever no particular grafting technique has been established in nurseries, our advice is to adopt this technique for olive grafting; where, instead, the skilled nursery hand is already expert in other procedures, it is advisable to use these, although a comparison with the results of bark grafting is recommended. Collection and conservation of scionwood Olive scionwood is obtained from 1-year-old vigorous shoots, collected from mature and healthy stock plants. Stock plants can be located in productive orchards or in in-field collections. However, in order to achieve a maximum uniformity of plants, scionwood should be collected annually from the same plants, and it is better if they are grown in a specific area of the nursery (the ‘plot of stock plants’) where they should be periodically checked for the maintenance in high sanitary and genetic conditions. It is quite common that the scionwood has to be preserved for a period (from few days to some weeks) before it is used for scion preparations, especially when collected outside the nursery. A brief period of preservation can be simply obtained by storing the material (just after collection) in a fresh room of the nursery, in layers (not over one metre) covered with jute sacks. The sacks are kept lightly damp, avoiding excessive humidity in order to reduce the risks of fermentation and of getting mouldy. The day before grafting time, the sacks are removed, as the scionwood should not be moist on the surface at the moment of the grafting operation. When long periods of conservation are necessary (for instance, when large quantities of grafted plants have to be prepared), the material should be stored in a refrigerator at 3–4°C until grafting time. Plastic bags should never be used to store the scionwood. To avoid dehydration, the shoots can be placed in vertical position inside a pot containing 1–2 cm of water, remembering that these basal ends of the shoots must not be used for grafting. Grafting time The graft, since it depends on the bark separating readily from the wood, can be done in spring (April–May in Central Italy, one month earlier in Southern Italy), i.e. after active growth of the seedling rootstocks has started. In general, this optimal condition for grafting olive plants lasts about one month. Grafting olive trees should be avoided during hot windy days. If rootstocks are grown in the greenhouse, it is possible to anticipate the grafting time by about one month. Attempts to graft in different periods of the year (e.g. in late summer or in autumn) have been unsuccessful because either grafts do not heal, or buds do not burst. Bark grafting technique In the olive, bark grafting is generally performed on 18-month-old seedlings in pots or in the field, provided they have attained a thickness of 6–8 mm (all plants below such size are discarded). Only one scion per rootstock is grafted. The whole procedure is illustrated in Figure 4.7.

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Figure 4.7 Bark grafting in olive. 1, Scion (a, side view; b, front view). 2, Vertical incision on the rootstock. 3, Cork lifted before scion insertion. 4, Insertion of scion, and protection with fresh cork (traditional technique). 5, Tying with raffia. 6, Completion of tying. 7, Protection with wax of cut surfaces (Marinucci 1969).

The best scionwood is taken from the middle part of shoots (Plate VIII), rather than from the apical parts (where the wood is still immature) or from the basal parts (too thick). In general, only the portion of scionwood having the right thickness (3–4 mm) should be used to obtain scions. The scion is 6–8 cm long, and is prepared in such a way as to contain a node in its upper third (Plate IX). The two leaves of the node are cut to 1/4–1/3 of their size to limit scion dehydration. One oblique cut is made with the knife along one side at the base of the scion, extending about one-third of the way into the

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scion (Plate X). It was an ancient practice of olive grafters to make a second small, transversal cut, leaving a sort of ‘shoulder’ at the top of the oblique one. The purpose of this shoulder was to minimise the separation of bark and wood after the insertion in the rootstock, especially when using thicker scions. Today, grafters prefer to use thin scions and to avoid the ‘shoulder’ cut. However, the scion should not be cut too thin, or it will be mechanically weak and break off at the point of attachment to the rootstock. To prepare the rootstock to be grafted, the stem is cut 5–10 cm above the soil, i.e. where its diameter is between 6 and 8 mm (pencil size; Plate XI). A vertical knife cut (2 cm long) is then made at the top end of the stub through the bark to the wood (Plate XII). The bark is lifted slightly along both sides of this cut. The scion is then inserted between the bark and the wood of the rootstock, centred directly under the vertical cut of the bark (Plate XIII), and gently pushed downward to penetrate a bit deeper below the bark. The graft zone is then fastened with raffia or rubber bands (see box ‘Main tools and accessories for olive grafting’) in order to produce a firm contact between the scion and the rootstock. After the scion has been fastened by tying, all cut surfaces, including the top of the scion, are thoroughly covered with grafting wax (Plate XIV). In general, best productivity in terms of grafts per hour is achieved with a group of three, i.e. one grafter and two helpers (one devoted to the preparation of plants and materials, one to tying the grafted plant and one to the application of wax; Fig. 4.8). Three experienced workers can make 200–300 grafts per hour (1500–2500 grafts per day).

Figure 4.8 A greenhouse with potted grafted plants, after tying and waxing.

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Care of the grafted plants Both in pots or in the field, it is always better to avoid irrigating the grafted plants for the first 10 days after grafting or, if irrigation is necessary, attention should be paid to keeping the grafting point dry. Callusing takes place within 15–20 days, when petioles naturally abscise from successfully grafted plants. If a correct grafting technique has been applied, the rate of ‘takes’ should be over 75%; rates above 90% are considered normal. If the graft ‘takes’, two new shoots develop on the scion within 30 days. When these have reached 5–10 cm in length, the best shoot is chosen and the other one is removed. In the summer, the young plants are supported with stakes to ensure vertical growth (Plate XV). MAIN TOOLS AND ACCESSORIES FOR OLIVE GRAFTING Knives: The best knives for olive grafting have a folding blade of high-carbon steel. To ensure good quality bark grafting, the knife must be kept always razor sharp. Raffia: This is a fibrous tissue from the upper sides of young leaflets of the palm Raphia vinifera. The use of raffia is an ancient and still common system to tie the graft point in the olive nurseries of Pescia. The raffia must be softened for a few minutes in water before use. As it is stronger and more persistent than rubber bands, it is best to check the graft periodically and cut the raffia soon after the graft has healed to prevent girdling or constriction at the grafting point. Rubber bands: Special rubber bands for grafting have become very common in the last 20 years to fasten grafted olive plants. In comparison with raffia, they have the advantage that they rot and break within a couple of months, i.e. before any damage can occur at the grafting point. However, a periodical inspection of the grafted plants is still recommended. Grafting wax: The application of wax over the grafting point has the aim of avoiding the loss of moisture and the consequent death of the tender cells of the cut surfaces, essential for callus proliferation and graft healing. Moreover, the wax prevents the entrance of rot microorganisms. Beeswax is a natural compound, traditionally used in the olive nurseries of Pescia to cover the graft point, although many other commercial preparations can be equally satisfactory. Beeswax is a hot wax with a good adherence: it is not washed off by rains, it doesn’t melt during hot days, and it retains its pliability during the enlargement of tissues at the grafted point. Before application, the beeswax is softened in water, using a small gas burner that is always kept close to the grafter’s helpers. Care must be taken because beeswax is inflammable.

4.3.4 Grafting on adult trees Topworking The grafting of cultivars on adult plants is used when unproductive cultivars, or those with poor resistance to disease, parasites and adverse climatic factors, or those producing fruit with a low oil content or whose oil is of poor quality, are to be changed to cultivars with better characteristics. The most-used grafting technique is the crown graft, carried out in early to mid-spring on branches with a diameter greater than 5 cm. Three wellspaced primary scaffolds (10–15 cm in diameter) are selected on the mature olive tree and cut back towards the trunk. As the graft may take two years to reach a suitable size, some additional branches are left to provide photosynthetic support (‘nurse branches’).

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Two to three scions are inserted into each limb stump. The scion is prepared in a similar way to that for the bark grafting described above. To protect the graft, grafting wax is used followed by a coat of white latex house paint to keep the temperature down. In response to limb removal, the olive tree is likely to produce large numbers of suckers which must be removed. When the graft â&#x20AC;&#x2DC;takesâ&#x20AC;&#x2122;, the best scion per limb is allowed to grow and the others, together with nurse branches, are removed. Multiple scions can be also topworked on a stump of olive, after the main trunk of the tree has been cut (Fig. 4.9). This practice is performed when a senescent or heavily damaged tree has to be recovered; scionwood may be of the same plant or of a different cultivar.

Figure 4.9 Topworking (bark graft) on an olive stump: a, Scion; b, The stump with the bark lifted; c, Final view of the graft, after scion insertion, tying and waxing.

Grafting on suckers Another form of topworking can be performed on the olive when grave damage (due to cold spells, mechanical damage, severe pathogen attacks, etc.) has involved most of the tree, and it is deemed convenient to reconstruct it by using the existing root system rather

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than replanting new trees. In this case the tree should be cut back to soil level, and a variable number of suckers (as many as possible in the first year) left to grow freely until the canopy is reconstituted and the plant starts producing again; this can take just a few years. If the tree had been propagated by grafting, some suckers (or all of them) may originate from the rootstock, which as a rule was a seedling. Such a canopy is not suitable for production, both because the characteristics of the new tree are going to be different from the desired cultivar (most likely worse), and because the tree will maintain juvenile features for a long period. In this instance it will be necessary to graft the selected suckers with a suitable cultivar, which of course can be different from the pre-existing one. With respect to the technique, it will be a regular topworking procedure, as previously described; the most vigorous and better positioned suckers will be selected for grafting – no more than 3 or 4. The graft should be made as low as possible, to reduce the likelihood of the development of latent buds from the rootstock. Grafting on wild olive trees This technique is still occasionally used in southern olive subzones where the species grows spontaneously as an element in the so-called Mediterranean maquis. Important populations of wild olives used to be found throughout the Mediterranean basin; today they are reduced, and many such areas are protected. These trees used to be converted to producing olive trees by topworking with specific cultivars. This type of activity is a complicated and costly process of adaptation which also calls for soil improvement. Transformation in the Mediterranean maquis requires clearing and the suppression of species which are associated with the wild olive. This, evidently, is in conflict with the established rules for protecting the environment and the integrity of ecological systems, and therefore decisions in each case should not be based on immediate economic results alone. It must be remembered that the wild olive does not have an important taproot, as was commonly believed, and thus does not enjoy special resistance to arid conditions. It is also very sensitive to low temperatures. Further, the natural populations of wild olives are generally grouped in a totally irregular manner and individual trees differ so greatly in vigour and development as to compromise the economic success of the future olive grove. The only advantage, when the wild olive trees are vigorous, is that they rapidly enter the productive phase, but this can be achieved today in young olive plantations when modern methods of pruning and production are employed, as has been shown by recent results of intensive olive production. In the Mediterranean basin, bark grafting of wild olives is carried out in early spring. When livestock can be kept out of the grove during its transformation, grafts are made as low as possible; otherwise, as in certain areas of Sardinia where the arboreal/pastoral system is important, the graft is set much higher up. In Apulia, large spontaneous plants or suckers from wild trees are used. These are lopped in situ at 15–30 cm above the soil and transported to the nursery where, after grafting, they are set, stratified, in a mixture of loose soil and fresh horse manure. By the following autumn these plants have shoots from 80–100 cm long and are ready for final transplanting. Adult plants (8–20 years) are also used, grafted where found, i.e. in the natural woodland, with multiple shoots, after having been lopped at a height of 40–150 cm. The grafting is carried out in spring and

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Plate I Indoor germination benches. Nets are for protection against rodents.

Plate III Detail of seedling development in a tray, six months after sowing.

Plate II Germination in a tray, four months after sowing.

Plate IV Seedlings at transplant time in the spring.

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Plate V Growing seedlings in containers.

Plate VII Close-up of a seedling of the right size for grafting.

Plate VI Sunken plots for seedlings. These are sufficiently developed for grafting.

Plate VIII Medium-sized well-developed shoots (scionwood) from which scions will be prepared.

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Plate IX Steps in the preparation of the scion. From left: scionwood, a one-node scion, leaf trimming, whip cut.

Plate XI Seedling cut back to 5â&#x20AC;&#x201C;10 cm length. Leaves are still present on the stub, but will be excised.

Plate XII A longitudinal slit is made at the top end of the stub. Plate X Detail of the oblique cut on the scion.

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Plate XIII The scion is inserted under the lifted bark of the seedling.

Plate XV Grafted plants at the end of summer. The developed (inclined) shoots need stacking.

Plate XIV The graft is tied with a rubber band, then waxed.

Plate XVI Grafted cuttings: preparation of the two components for the bench grafting.

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Plate XVII Tying after ‘wedge’ or ‘whip tongue grafting’.

Plate XIX Differences in the primary root system of a rooted cutting (left) and of a grafted seedling (right).

Plate XVIII Shoots developing from a grafted cutting.

Plate XX An upgrowing olive shoot from which explants are taken to initiate a micropropagation cycle.

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Plate XXI Uninodal segments of olive which will be used for the initiation of a micropropagation cycle. After the preparation, the explants are immediately treated with disinfectants (see Table 5.2). Plate XXII To reduce the risks of spreading contaminants, shoot cultures are initiated in test tubes.

Plate XXIII Elongated shoots of olive (cv Frantoio) at the moment of subculturing.

Plate XXIV Bi-nodal segmentation of the shoots shown in Plate XXIII, prior to being transferred to fresh OM medium.

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Plate XXV 500 mL glass jars, containing shoot cultures (cv Frantoio) before (left) and after (right) subculturing. Note the different colour of the media, gelled with Gelrite (left) or agar (right).

Plate XXVII Four- to six-node elongated shoots (cv Frantoio) ready for rooting. The callus proliferation at the base of shoots is removed before rooting treatments.

Plate XXVI Gas-permeable boxes (Vitro Vent™; DUCHEFA, Haarlem, The Netherlands) which can be used for both shoot proliferation and storage at 4°C (as shown). Plate XXVIII Rooted shoots of ‘Frantoio’ ready for Stage IV (acclimatisation).

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Plate XXIX Somatic embryogenesis in olive (cv Canino): many secondary somatic embryos at different stages of development, formed at the radicle end of a primary somatic embryo, with no evidence of interposed callus (× 12).

Plate XXXI An isolated well-formed somatic embryo (× 20).

Plate XXX Further embryo development to the cotyledonary stage (× 12).

Plate XXXII Synthetic seed preparation using olive somatic embryos (× 20).

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the plants remain in the woodland until required. On removal of the plant from the woodland it is lopped about 20 cm above the graft and the roots are cut back; it can then be planted in its final site.

4.3.5 Grafted cuttings This technique, quite common for some fruit species (particularly grapes, although it is also possible with apples and pears), is also named ‘bench grafting’ because it is performed at benches by skilled grafters as a large-scale operation. In the olive, it has been proposed in the past (Jacoboni & Fontanazza 1976), as having as its principal object the multiplication of cultivars that do not root readily under mist. Cuttings and scions are taken from 1-year-old shoots, the cuttings from selected plants (generally grown from seed and chosen for their high rooting capacity), the scions from the cultivar to be propagated. The cuttings are 15–18 cm long, prepared leaving two pairs of leaves. The scions (of the same thickness as the cuttings) are prepared with two nodes and one pair of leaves (Plate XVI), and grafted at the bench with the ‘wedge’ or ‘whip and tongue’ technique (Plate XVII). The base of the grafted cuttings is then treated with IBA (2000–4000 ppm), and set in a substrate of peat and perlite, suitably moistened and maintained under plastic cover to preserve humidity. The base of the grafts should be maintained at approximately 28°C. In these conditions of high temperature and humidity, rooting of the cutting and the ‘take’ of the graft occur almost simultaneously within 60–90 days (Plate XVIII). Although the technique has never found any commercial application, it can be a useful method for propagating small numbers of plants when little space is available.

4.4 Production of clonal rootstocks in the olive The use of clonal rootstocks is of fundamental importance for improving productivity of many fruit trees, making it possible to control important characteristics, such as vigour, precocity of production, resistance to parasites, diseases, and adverse environmental and soil conditions. Unlike the case for most fruit species, information on olive rootstocks is thus far scattered and fragmentary, not only because in many olive producing countries the use of grafting is limited but also because, even where it is generally practised, the rootstocks used are produced from seed, either of cultivars or of wild types. Although there has been no systematic research on the best combinations of scion/rootstock which would allow any control of the characteristics of the olive tree, in recent years some clonal rootstocks have been proposed as a result of studies carried out mainly in California, Italy and Spain (Bartolini et al. 1998). For example: • ‘Oblonga’ (California), employed because of its resistance to Verticillium dahliae • ‘FS–17’ (Italy), which induces a reduction of vigour in the grafted cultivar • ‘Lechin de Sevilla’ (Spain), tolerant to dry and calcareous soils and inducing cold resistance in the grafted cultivars (‘Gordal sevillana’, ‘Manzanilla de Sevilla’, ‘Morona’) • ‘Verdial de Badajoz’ (Spain), tolerant to dry and calcareous soils (grafted with ‘Manzanilla de Sevilla’)

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• ‘Hojiblanca’ (Spain), tolerant to dry and calcareous soils and inducing cold resistance in the grafted cultivar (‘Aloreña’) It has to be noted that the above information is generally agreed on in the various countries, or declared by breeders; however, definitive experimental data are not yet available.

4.5 Grafted plants or self-rooted plants? One of the first questions the olive grower has to face is which type of planting material should be selected: grafted or self-rooted plants. It must be remembered that not all cultivars can be obtained self-rooted; in this case, as for many table cultivars, the choice of the grafted tree has no alternative. When both types of material are available at comparable prices, then the problem arises as to which is best. The grafted versus selfrooted trees issue is an old one. A discrete amount of research was undertaken on this topic in the 1960s and 1970s, both in Italy and California. Generally speaking, no significant or generalisable differences were noticed in specific trials with respect to productivity (Guerriero et al. 1972, 1974). Thus, in Central Italy (Tuscany) grafted trees proved to be more productive; in the south (Sicily) the advantage was for the self-rooted trees. But a closer examination of the results shows different behaviours in different experimental fields, and, above all, different performances depending on the cultivar being evaluated. Most growers are interested in better vigour in the first years after plantation; even in this case differences exist among cultivars which cannot be generalised: ‘Moraiolo’ rootlings appear weaker than grafted plants, at least in the first years of culture; ‘Frangivento’ seems to have an opposite behaviour, while ‘Leccino’ shows no differences. Self-rooted ‘Frantoio’ was weaker if compared to plants grafted on ‘Frantoio’ seedlings, but more vigorous if compared to ‘Frantoio’ grafted on ‘Lea’ seedlings. Other cultivars (‘Manzanillo’, ‘Mission’, ‘Gordal’) have quite differentiated behaviours in the grafted/self-rooted comparison (Hartmann 1958). Substantial differences occur with respect to the development of the root system (Scaramuzzi 1963). Rooted cuttings develop their roots initially at a unique level in the soil, usually depending on the depth of planting. Usually, the root apparatus is quite superficial, with a wide geotropic angle. Seedlings on the other hand – irrespective of the fact, of course, that they are or not grafted – develop their roots from an axis of a discrete length, usually around 20 cm, and having a narrower geotropic angle tend to grow deeper into the soil (Plate XIX). This means that in arid, sandy, windy areas, self-rooted plants need in the beginning to be staked and irrigated regularly in the dry periods, and require greater care than usual. Practical experience does not always confirm this point of view: in certain arid zones (Southern Tunisia) the olive has long been propagated by ovules without any adverse effect on resistance to drought or longevity. Grafted trees, on the other hand, have a few drawbacks: the main one is that whenever it is necessary to reconstitute a damaged canopy (e.g. by frost, diseases, or senescence) all suckers coming from the stock, which are not suitable for production, need to be eliminated. Grafted trees should be planted with the grafting point below the soil level; however, it must be

Propa ga tion by grafting

remembered that with time, soil levels subside due to erosion, and the rootstock may appear and produce its own suckers. In addition to the above, and related to the container technology that is dominating in almost all nurseries, it should be remembered that all types of plants undergo transplants in containers of different sizes at least twice. This operation means breakage (often voluntary cuts, called transplant pruning) of longer roots, and regeneration of a root system, more or less compact, of a cylindric shape, and uniformly distributed. This means that, morphologically speaking, 2-year-old root systems coming from the two types of propagation are not easily distinguished, and consequently an analogous behaviour is to be expected in the soil. Finally, a known truth among plant physiologists is that roots, in their development, are far more influenced by the edaphic environment than by their genetic characteristics; this means that the alleged innate characteristic of better survival in the soil of the taproot, were it true, can hold true only in the very first period of root development. Then, in sandy soils, roots go deep in search of water, in clay soils they stay closer to the surface; that is, they respond to environmental stimuli in the same way. The opposite has never been demonstrated. This is not the final word on the issue. More research is needed on a wide scale to ascertain with precision the effect of grafting, and of the different genotypes used as stocks. Of course clonal rootstocks able to control plant life are sought, and all this needs to be checked in quite different environmental conditions.

References Acebedo, M.M., Lavee, S., Linan, J. & Troncoso, A. 1997. In vitro germination of embryos for speeding up development in olive breeding programmes. Sci. Hortic. 69 (3–4): 207–215. Baldini, E. 1986. Arboricoltura generale. CLUEB, Bologna (Italy), pp. 396. Bartolini, G., Prevost, G., Messeri, C. & Carignani, G. 1998. Olive germplasm: cultivars and world-wide collections. FAO, Rome, pp. 462. Calvino, M. 1930. Nuovo porta innesto dell’Olivo, Forestiera durangensis. Il Coltivatore. Carocci Buzi, C. 1945. Sulla nanificazione dell’Olivo. Ann. Ist. Sper. per l’Oliv. e l’Oleificio, Imperia. Casella, G. 1932–1934a. Primo contributo su alcuni porta innesti dell’Olivo. Ann. Staz. Sper. Frutt. Agrum., Acireale, n. s, vol. 1, n. 5. Casella, G. 1932–1934b. L’innesto dell’Olivo sul Lillà (Syringa vulgaris). Ann. Staz. Sper. Frutt. Agrum., Acireale, n. s, vol. 1, n. 5. Fontanazza, G. & Rugini, E. 1983. Graft union histology in olive tree propagation by cutting-grafts. Riv. Ortoflorofrutt. It. 67(1): 15–21. Guerrero, A. 1997. Nueva olivicoltura (4th edn). Ed. Mundi-Prensa, Madrid, pp. 282. Guerriero, R., Scaramuzzi, F., Crescimanno, F.G. & Sottile, I. 1972. Ricerche comparative tra olivi innestati ed autoradicati. Osservazioni nei primi anni di impianto. Tecnica Agraria, 4. Guerriero, R., Scaramuzzi, F., Crescimanno, F.G. & Sottile, I. 1974. Confronto tra diverse combinazioni di innesto dell’olivo su piante autoradicate. Tecnica Agraria, 4. Hartmann, H.T. 1958. Rootstock effects in the olive. Proc. Am. Soc. Hort. Sci., 72: 242–251. Instanbouli & Neville, 1977. Distinction entre germination physiologique (ou activation) et germination morphologique chez l’oliver (Olea europaea L.). C. R. Acad. Sc. Paris, t. 284: 2235–2237. Jacoboni, N. & Fontanazza, G. 1976. Un nuovo tipo di propagazione dell’olivo: l’innesto-talea in cassone riscaldato alla base. Italia Agricola, 133: 104–112. Lagarda, A., Martin, G.C. & Polito, V.S. 1983a. Anatomical and morphological development of ‘Manzanillo’ olive seed in relation to germination. J. Am. Soc. Hort. Sci. 108(5): 741–743. Lagarda, A., Martin, G.C. & Kester, D.E. 1983b. Influence of environment, seed tissue and seed maturity on ‘Manzanillo’ olive seed germination. HortSci. 18(6): 868–869.

Olive Propa ga tion

Lambardi, M., Rinaldi, L.M.R., Menabeni, D. & Cimato, A. 1994. Ethylene effect on in vitro olive seed germination (Olea europaea L.). Acta Hortic. 356: 54–57. Legesse, N. 1993. Investigations on the germination behaviour of wild olive seeds and the nursery establishment of the germinants. Sinet: an Ethiopian Journal of Science 16 (2): 71–81. Marinucci M. 1969 – Innesto. In Enciclopedia Agraria Italiana, Vol. VI. R. E. D. A., Roma. Milella, A. 1962. Ricerche sull’affinità di innesto fra la Phillyrea angustifolia e l’Olea europaea L. var. sativa. Studi sassaresi. Occhialini,O. 1906. L’inesto dell’Olivo sull’Orniello. L’Italia Agricola, n. 22. Rinaldi, L.M.R. & Lambardi, M. 1998. In vitro germinability and ethylene biosynthesis in cytokinintreated olive seeds (Olea europaea L.). Adv. Hort. Sci. 12 (2): 59–62. Rinaldi, L.M.R., Menabeni, D., Lambardi, M. & Cimato, A. 1994. Changes in carbohydrates in olive seeds (Olea europaea L.) during fruit maturation and their correlation with germination. Acta Hortic. 356: 58- 61. Scaramuzzi, F. 1957. Ricerche sul potere germinativo di semi di diversa età in Olea europaea L. L’Agricoltura Italiana, 56: 21–43. Scaramuzzi, F. 1958. Influenza dell’epoca di raccolta dei frutti sulla germinabilità dei semi di olivo. L’Agricoltura Italiana, 58: 387–402. Scaramuzzi, F. 1963. Osservazioni comparative sull’apparato radicale degli olivi in vivaio. L’Italia Agricola, n. 12. Sotomayor León, E.M. & Durant Altisent, J.M. 1994. Breaking of dormancy in olive seeds (Olea europaea L.). Acta Hortic. 356: 137–142. Tallarico, G. 1939. L’innesto dell’olivastro di Etiopia. L’Italia Agricola, n. 5. Vachkoo, A.M., Mughal M. S., Gupta O. P., Sharma K. C. 1993. Effects of some chemical and mechanical treatments on seed germination of olive (Olea cuspidata Wall.). Adv. Hortic. For. 3: 105–108. Voyatzis, D.G. 1995. Dormancy and germination of olive embryos as affected by temperature. Physiol. Plant. 95 (3): 444–448 Voyatzis, D.G. & Pritsa, T. 1994. The onset and disappearance of relative dormancy of olive embryos as affected by age. Acta Hortic. 356: 148–151.

5 In vitro propagation of the olive

5.1 Micropropagation Micropropagation is a powerful in vitro technique which allows propagation of true-totype pathogen-free genotypes, under controlled climatic conditions (temperature, photoperiod, light intensity) and in asepsis. Over the last two decades, many advances have been made towards the optimisation of the various steps involved in tree micropropagation, and several cultivars and rootstocks of fruit species (e.g. apple, peach, plum, pear, kiwifruit) are today commercially multiplied by means of this technology. In comparison with traditional cutting and grafting propagation, the features of micropropagation are: • mass propagation of specific cultivars or clones can be obtained, providing yearround nursery production • pathogen-free high-quality plants are produced, with a high standardisation of their morphological and agronomical characteristics • clonal propagation of difficult-to-root cultivars can be improved • techniques are available (i.e. meristem culture and micrografting) to obtain virusfree plants from infected donor plants • the distribution of commercial material in international trade is facilitated, allowing also movement of germplasm materials across quarantine barriers Micropropagation is a multi-step procedure, starting with the excision and the introduction in vitro of explants from a donor plant, and ending when micropropagated microplants are successfully transferred and established in vivo (Fig. 5.1). Murashige (1974) proposed dividing micropropagation protocols into three stages. As the technique evolved, other stages were indicated, so that at present five stages are considered critical to successful micropropagation:

Olive Propa ga tion

Figure 5.1 Schematic representation of a complete micropropagation cycle (Baldini 1986). Roman numerals refer to the different stages of the procedure (see text).

• • • • •

Stage 0 – Collection of explants Stage I – Initiation of cultures Stage II – Shoot proliferation Stage III – Shoot rooting Stage IV – Acclimatisation of plants

Stages I, II and III are carried out under in vitro conditions, and require a laboratory equipped for tissue culture (for a comprehensive description of the facilities and equipment of a tissue culture laboratory, see George 1993). No matter what the size of the laboratory, it should include three basic components:

In vitro propa ga tion of the olive

1. the ‘preparation area’ (Fig. 5.2), where media and glassware are prepared and sterilised. 2. the ‘transfer area’, where explants are introduced in vitro and periodically subcultured in tissue culture media. Transfers are done in a laminar airflow hood (Fig. 5.3), i.e. a working bench where the operator is exposed to a horizontal flow of sterile air (due to a passage through a filtering panel, mounted in the rear of the hood, outward on a positive pressure gradient). 3. the ‘growing area’, consisting of climatic chambers (Fig. 5.4) or large growing rooms where temperature, light intensity and photoperiod are controlled. At the start of the 1990s, few olive cultivars could be efficiently established and propagated in vitro, particularly when explants from adult trees were used. Micropropagators faced several problems in the development of effective protocols, among which were: (i) the difficulty of getting sterile cultures when explants (shoot tips, nodal segments) were collected from in-field or greenhouse plants, (ii) the heavy oxidation of tissues following explant preparation, and (iii) the laboriousness of establishing shoot cultures with some cultivars. Since the mid-1990s, impressive progress has been made towards the solution of these problems, so that an optimisation of the various steps involved in olive micropropagation has been gradually achieved. Table 5.1 lists the cultivars for which a complete procedure of micropropagation from mature trees has been published in scientific reports, and gives concise information on related protocols.

Figure 5.2 The preparation area of a micropropagation laboratory.

Olive Propa ga tion

Figure 5.3 A laminar airflow hood (Intern. PBI, Milan, Italy). Here the shoot cultures are periodically subcultured by an operator working under sterile conditions (see also text).

Figure 5.4 A climatic chamber (ProClimatic, Imola, Italy) where the shoot cultures are grown under controlled temperature and light conditions.

In vitro propa ga tion of the olive

5.1.1 Stage 0: Collection of explants The collection of shoots from stock plants, the primary source of explants (see below) to be introduced in vitro, is the first step of olive micropropagation. The choice of stock plants as a source of explants cannot be undertaken indiscriminately; as for cutting and grafting propagation, micropropagation takes great advantage of the use of elite stock plants, selected for their superior phenotypic characteristics, maintained in good sanitary conditions, and periodically controlled for genetic stability. Stock plants can be grown under field conditions or in a greenhouse (see also 6.1). Explants from in-field growing stock plants The initiation of micropropagation from adult field plants is difficult and time consuming, mainly because of the high contamination and strong oxidation of tissues following their introduction in vitro. In order to minimise oxidation, the best results are obtained when explants are collected from vegetative structures with strong characteristics of juvenility, such as: • vigorous upgrowing shoots (Plate XX) • suckers • ovules An example of use of ovules is proposed by Cañas et al. (1992): here, shoots are induced to sprout from ovules cultured in semi-aseptic conditions, after which uninodal leafy explants are excised and cultured in vitro. Following axillary bud stimulation, conventional shoot proliferation and rooting can be obtained. It is important to remember that when using this type of explant, a strong condition of juvenility can persist even after the in-field plantation of micropropagated plants, for which the onset of fruiting can be delayed for one or more years. Explants from greenhouse growing stock plants To yield more hygienic explants, potted stock plants, grown under greenhouse conditions, are the best material to initiate in vitro propagation of the olive. An ideal stock plant repository, as a source of explants, can be established in the greenhouse by grafting scions from selected cultivars onto potted seedlings. When old (or even senescent) trees have to be reproduced, stock plants can be derived from two or more cycles of repeated graftage (‘serial graftage’; Hartmann et al. 1990). The technique produces a strong effect of rejuvenation, and the stock plants which are obtained are a source of explants very reactive to the initiation of in vitro propagation. Plastic greenhouses or glasshouses are most suitable for stock plant growing. In both cases it is important that the plants do not receive water by overhead irrigation. Devices that provide water to the plants directly in the pot or by capillarity, such as trickle irrigation and hydroponics, are suitable. It is advisable to carefully control stock plant sanitary conditions. For this purpose, culture indexing tests for bacterial and/or fungal contaminations can be made (Cassels & O’Herlihy 2003). Stock plants with strong microbial contamination must be removed, as well as plants that are suspected of virus infection.

Olive Propa ga tion

5.1.2 Stage I: Initiation of cultures The purpose of Stage I is to initiate an axenic culture of the olive. Working with woody plants, the developmental stage of an explant is always of relevant importance. Indeed, the physiological condition and the size of the explant may markedly influence the success of a micropropagation protocol. In the olive, best explants to initiate in vitro culture are tender apical twigs and uninodal segments (1â&#x20AC;&#x201C;2 cm in length, Plate XXI), excised from vigorous shoots, soon after spring sprouting. Occasionally, bi- or tri-nodal segments are used. Disinfection of explants Tissue disinfection before its introduction in vitro is a fundamental step in olive micropropagation. Although old procedures also included passages in ethanol solutions, the present tendency is to avoid their use because they cause tissue dehydration. Depending on the level of contamination of the stock material, treatments with sodium hypochlorite (NaClO; Fig. 5.5) at different concentrations, alone or in combination with mercuric perchlorate (HgClO2), can be applied. In Table 5.2, three different procedures are reported. In case the level of contamination of in-field stock material is unknown, the procedure indicated for greenhouse-grown plants can be used at first. However, if decontamination is not achieved, stronger disinfection procedures can be applied afterwards.

Figure 5.5 Disinfection of explants in a sodium hypochlorite solution.

Before the application of whichever disinfection procedure, it is also advisable to: â&#x20AC;˘ induce a slight water stress on potted stock plants by diminishing the amount of watering a few days before explant collection, in order to reduce microbial contamination

In vitro propa ga tion of the olive

• handle the material as soon as possible, after excision from the stock plants • rinse the explants in running tap water for at least 30 minutes, before initiation of the sterilisation procedure The olive is a woody species with a naturally high content of phenolics and, as a consequence, explant tissues are easily killed or necrosed from the strong oxidative reaction following their excision and in vitro introduction (see box ‘Oxidation of explants and medium’). To limit oxidation, the culture medium can be supplemented with antioxidants, such as ascorbic acid (10–20 mg/L) and citric acid (100–200 mg/L), or the explants can be pre-treated (30–60 min) in a solution of sterile water plus antioxidants before being plated on the culture medium. However, these procedures are ineffective when high quantities of phenols are released. Moreover, antioxidants such as ascorbic acid are only suited to short period interventions because they very quickly become strong oxidants themselves. Hence, when strong oxidation occurs, the best intervention consists of frequent subculturing of explants (up to one every 24 hours) in fresh medium until browning of the medium is no longer observed. OXIDATION OF EXPLANTS AND MEDIUM When plant tissues are exposed to a stress condition (such as the injuries produced by explant excision), metabolism of phenolic compounds is stimulated. The phenolic components are then released in the medium from broken cells and their neighbouring cells. In general, phenolics are labile products that are very easily oxidised, giving rise to quinones and polymerised material. Such oxidation products are very phytotoxic and strong oxidants themselves, inducing an irreversible oxidation process. A strong browning of the medium, particularly in the area surrounding the explant, is a clear symptom of phenolic oxidation and, as a consequence, of explant degeneration.

End of Stage I Following sterilisation, explants are cultured in test tubes (one per tube) for 4–6 weeks (Plate XXII), in order to avoid the diffusion of contaminants that have escaped disinfection. After this period, the explants evolve into microshoots of 2–3 cm, which can be moved to Stage II.

5.1.3 Stage II: Shoot proliferation In the majority of woody species, shoot proliferation is generally pursued through the development and proliferation of both axillary and adventitious buds, as a result of the stimulation produced by exogenous cytokinins. The former are the buds naturally present at the axil of leaves, the latter are de-novo formed buds, mainly originating at the basal ends of shoots. Olive shoots are characterised by a strong apical dominance, which influences their natural growth habit and which is not suppressed in vitro by cytokinin treatments. Hence, the proliferation of shoots through the stimulation of axillary and adventitious buds is of minor importance, and shoot ‘multiplication’ is achieved at each subculture mainly by means of uni- or binodal segmentation of elongated shoots (Plates XXIII and XXIV).

Olive Propa ga tion

Medium formulation In general, a micropropagation medium consists of inorganic salts (macro and microelements), vitamins, carbohydrates, and a gelling agent. Specific growth regulators (exogenous hormones) are then added at different stages of the micropropagation OLIVE MEDIUM FORMULATION procedure. In the olive, a specific medium (as All the compounds are reported as mg/L. regards inorganic salts, vitamins and amino Macroelements acids) was formulated on the basis of the • KNO3 1100 analysis of the main mineral elements of • NH4NO3 412 shoot apices from field plants, during their • Ca(NO3)2 ⋅ 4H2O 600 • CaCl2 ⋅ 2H2O 440 rapid growth (Rugini 1984). The medium, • KCl 500 known as OM (see box ‘Olive medium • MgSO4 ⋅ 7H2O 1500 formulation’), marked an important step • KH2PO4 340 forward in the improvement of olive axillary Microelements bud stimulation and shoot multiplication. • FeSO4 ⋅ 7H2O 27.8 The effectiveness of OM formulation for • Na2EDTA 37.5 • MnSO4 ⋅ 4H2O 22.3 a wide pool of olive cultivars is demonstrated • H3BO3 12.4 in Table 5.1, as two-thirds of the procedures • ZnSO4 ⋅ 7H2O 14.3 were developed using the medium in its • Na2MoO4 ⋅ 2H2O 0.25 original formulation or with little • CuSO4 ⋅ 5H2O 0.25 modification. As regards the carbon source, • CoCl2 ⋅ 6H2O 0.025 • KI 0.83 it has been shown that, in comparison with Vitamins sucrose, mannitol (one of the major • Myo-inositol 100 carbohydrates of olive metabolism) increases • Thiamine. HCl 0.5 olive shoot proliferation, improves the • Pyridoxine. HCl 0.5 general quality and uniformity of shoot • Nicotinic acid 5.0 cultures and reduces basal callus formation • Biotin 0.05 • Folic acid 0.5 (Leva et al. 1994). Mannitol should be Amino acids supplemented at 30–36 g/L concentrations. • Glycine 2 Finally, various gelling agents can be used to • Glutamine 2194 produce a semi-solid medium, such as agar (at 0.5–0.8% concentrations) or agar substitutes (e.g. Gelrite, Phytagel™) at 0.2–0.3% concentrations. The pH of the medium is adjusted to 5.7–5.8 before autoclavage (also see box ‘Medium preparation’). Growth regulators Among the various cytokinins, zeatin is the one producing maximum stimulation of shoot proliferation for the olive. This growth regulator is applied in a wide range of concentrations, alone or in combination with other growth regulators. As can be seen in Table 5.1, its concentration ranges, when used alone, from 0.5 mg/L (cv ‘Maurino’, Bartolini et al. 1990) up to 10 mg/L (cv ‘Arbequina’, Otero & Docampo 1998). However, for the majority of olive cultivars, the best proliferative concentration is in the range of 1 to 4 mg/L. It is commonly stated that the other cytokinins cannot replace zeatin in promoting olive bud stimulation. In general, a lower proliferative efficiency is reported for both

Table 5.1 Summary of best culture conditions in micropropagation of olive cultivars. Only procedures that were initiated with explants from mature trees are reported (m, modified medium; NR, not reported). From Lambardi and Rugini 2002. Cultivar

Original explant Medium

Shoot proliferation hormones

ratio

Medium

Shoot rooting hormones

Reference

Micrografted shoots

DKW (a)

BA, 4.4 µM + IBA, 0.05 µM

7–8

/2 DKW

IBA, 0.5 µM

Revilla et al. 1996

Arbequina

Uninodal explants

zeatin, 10 mg/L

IBA, 3 mg/L or NAA, 1 mg/L

Otero & Docampo 1998

Arbequina, Empeltre, Picual

Uninodal explants

BA, 1 µM + TDZ, 1 µM

2.5–2.8

Compost

IBA, 15 µM + IAA, 10 µM

Garcia-Fèrriz et al. 2002

Carolea

Uninodal explants

zeatin, 4 mg/L

NAA, 2 mg/L or IBA, 2. 5 mg/L + Putrescine, 160 mg/L

Briccoli Bati et al. 1994

Dolce Agogia

Uninodal explants from suckers

/2 MS

zeatin, 10 mg/L + IBA, 0. 5 mg/L + GA3, 0.5 mg/L

/2 Knop / Heller (b)

NAA, 4 mg/L

Rugini & Fontanazza 1981

Frantoio

Uninodal explants

zeatin, 4 mg/L or 2iP, 4 mg/L

9–10

Various

NAA, 1 mg/L

Rugini 1984

FS-17

Uninodal explants

mOM

zeatin, 2 mg/L or 2iP, 4 mg/L

7.2

/2 MS

NAA, 2 mg/L

100

Mencuccini 1998

Kalamon

Uninodal explants (c)

mOM

zeatin, 10 mg/L + crude extract (d)

mOM

NAA, 2 mg/L + crude extract (d)

Rama & Pontikis 1990

Kalamon

Uninodal explants

WPM (e)

BA, 1 mg/L + IBA, 1 mg/L + GA3, 0. 1 mg/L

9.6

WPM

IBA, 2 mg/L

Dimassi-Theriou 1994

Maurino

Tri-nodal explants

mMS

zeatin, 0.5 mg/L or TIBA, 0.5 mg/L

mMS

K-IBA, 500 mg/L (‘dip method’)

Bartolini et al. 1990

Memecik, Domat

Nodal explants

mOM

BA, 1 mg/L

Meski

Uninodal explants

mOM

zeatin, 1 mg/L

mOM

IBA, 1 g/L (‘dip method’)

100

Chaari-Rkhis et al. 2002

Moraiolo

Shoot apices

zeatin, 4 mg/L or 2iP, 4 mg/L

9–10

Various

NAA, 1 mg/L

Rugini 1984

Nocellara Etnea

Uninodal explants

mOM

zeatin, 4 mg/L

5.6

mOM

NAA, 1. 5 mg/L

100

Briccoli Bati & Lombardo 1995

Tunisian cultivars (f)

Uni- or binodal explants

mMS

zeatin, 2 mg/L + kinetin, 1–2 mg/L (or BA, 2 mg/L)

1–8

NAA, 2 mg/L

100

Chaari-Rkhis et al. 1999

b c

From Driver & Kuniyuki (1984) From Heller (1953) Uninodal explants from shoots developed from excised ovules

d e f

/2 OM

/2 mMS

Seyhan & Özzambak 1994

Crude extract was obtained by fresh tissue maceration and methanol extraction of olive ovules From McCown & Lloyd (1981) Cvs ‘Sig de Sfax’, ‘Chemlali de Sfax’, ‘Chetoui’ and ‘Meski’

/2 OM

In vitro propa ga tion of the olive

Arbequina

Olive Propa ga tion

6-γ,γ-dimethylallylaminopurine (2iP), and 6-benzyladenine (BA), the latter being the most used cytokinin in tissue culture. BA, in particular, is not very effective in the olive, as it produces short thin shoots and, occasionally, abundant callus proliferation. Over time, researchers have proposed various combinations of growth regulators – containing, besides the above-mentioned, kinetin, thidiazuron (TDZ, a cytokinin-like substance), gibberellic acid (GA3), and IBA – to promote the stimulation of axillary buds; however, as they are always restricted to one or few specific cultivars, they cannot be considered for general application. Nevertheless, in consideration of the high cost of zeatin, efforts are currently being made to find cytokinin combinations that can replace, or at least reduce, zeatin use, with no marked reduction of shoot proliferation. Towards this end, the combined use of zeatin with 2iP and TDZ seems promising. MEDIUM PREPARATION To prepare the culture medium, add deionised high-quality water (one-half of the final volume) to a large beaker. While stirring, add required amounts of each ingredient from stock solutions by pipette or a graduated cylinder. Alternatively, an appropriate amount of the commercial OM preparation (DUCHEFA, Haarlem, The Netherlands), containing microelements, macroelements and vitamins, can be used. Then add appropriate amounts of mannitol and agar. Growth regulators may be added at this time, or after autoclavage by filter-sterilisation under the laminar airflow hood when the medium has cooled to 40–50°C (see box ‘Addition of zeatin to the culture medium’). The solution is made up to the prescribed volume by adding water, after which the pH is adjusted to 5.7–5.8 by means of a pH meter and drop-by-drop additions of 1M HCl or NaOH. The solution is then heated (in a microwave oven or on a hot plate) to melt the agar. After stirring, the hot solution is dispensed into the culture jars, which are then closed with a plastic film and sterilised in an autoclave for 20 min at 121°C. After autoclavage, the jars are cooled to below 70°C before being taken out off the autoclave. After gelification, the medium is ready for use when it reaches room temperature.

ADDITION OF ZEATIN TO THE CULTURE MEDIUM To prepare 100 mL stock solution of zeatin at 200 mg/L concentration, dissolve 20 mg of zeatin with few drops of 1M sodium hydroxide (NaOH) inside a 100 cc beaker. Stir while adding distilled water. Transfer into a 100 cc graduated flask and bring to the final 100-mL volume. To have a medium with 1 mg/L concentration of zeatin, add 5 mL of the stock solution to 1 litre of culture medium (1 mg/L = 4.56 µM). Stock solutions can be stored at 4°C for up to one month. All cytokinins are unstable during autoclaving. Zeatin, in particular, can lose part of its efficacy during sterilisation at 121°C. Therefore, it is advisable to filter-sterilise an appropriate aliquot of stock solution directly in the culture jars after autoclavage. However, if addition before autoclaving is necessary, a good approximation is obtained by increasing the desired concentration by 50%. Similar procedures can be followed to prepare stock solutions of the other cytokinins, with the exception of TDZ which has to be dissolved with the addition of a few drops of dimethylsulfoxide (DMSO.)

In vitro propa ga tion of the olive

Culture conditions and subculturing In commercial laboratories, glass jars of 500–750 cm3 of volume (diameter 9–10 cm), closed with glass lids and wrapped in plastic film, are used for olive shoot proliferation (Plate XXV). The jars contain 15–20 shoots on average, which are cultured on 100–150 mL of semi-solid medium. Alternatively, polypropylene containers can be used, such as Magenta™ (SIGMA Chemicals, vol. 350 cm3) and Vitro Vent™ (DUCHEFA, Haarlem, The Netherlands, vol. 900 cm3) boxes (Plate XXVI). The latter have closures which allow continuous gas-exchange between the inner volume of the container and the outside environment, markedly reducing condensation and accumulation of volatile compounds, such as ethylene and carbon dioxide. These characteristics improve the quality of olive cultures. The jars are maintained in climatic chambers, where optimal growth conditions for olive shoot cultures are: 23–25°C of temperature, 16h of photoperiod, 40–60 µmol m-2 s-1 of photosynthetically active radiation provided by cool-white fluorescent lamps. Depending on the growth characteristics of specific cultivars, shoot cultures are transferred to fresh medium every 4–7 weeks to maintain them in good proliferative activity. Shoot elongation Following proliferation, shoots of some olive cultivars are too short to be used directly for rooting. In this case, an elongation phase (i.e. an ‘intermediate’ stage between stages II and III) is very useful. Olive shoot elongation is stimulated by a passage in fresh medium containing filter-sterilised GA3 (20–40 mg/L). After 3–4 weeks, when shoots have elongated to 4–5 nodes, they are ready for rooting (Plate XXVII).

5.1.4 Stage III: Shoot rooting Obviously, selection for uniformity and removal of abnormal, aberrant or diseased shoots should be made prior to stage III. Rooting on IBA or NAA-containing medium Great advances have been made in the rooting of micropropagated olive shoots over the last decade, so that even cultivars ‘recalcitrant’ to cutting propagation can now be satisfactorily rooted in vitro. Medium formulation and auxin treatment are both important factors to promote olive shoot rooting. Table 5.1 shows that with only two exceptions (‘Dolce Agogia’, ‘FS–17’), rooting was performed on the same medium of shoot proliferation, generally by reducing to 1⁄2 the strength of the original macroelement formulation. As regards root-promoters, both IBA and 1-naphthaleneacetic acid (NAA) have been indicated as most effective for the stimulation of rooting. IBA is supplemented at concentrations of 2–3 mg/L (9.8–14.7 µM). As NAA produces a stronger stimulation than IBA, it is most often applied at 1 mg/L concentration (5.37 µM). In comparison with cytokinins, IBA and NAA are more stable when heated at 121°C. Therefore, they can be added to the rooting medium before autoclavage with no need to increase the concentration. Additional root-promoting methods Over time, several additional rooting methods have been proposed to improve micropropagation of difficult-to-root olive cultivars, in combination with auxin

Olive Propa ga tion

treatments. Experimentation has proved the effectiveness of two of them, i.e. the basal etiolation of shoots and the addition of polyamines to rooting medium. It is well known that many woody plants root best if placed in the dark during the auxin treatment period. However, in the olive a similar stimulatory effect, together with a better maintenance of shoot quality, is obtained with the etiolation of only the basal portion of shoots (Rugini et al. 1993). Basal etiolation of shoots is easily achieved by painting the outside of the jars black, and by covering the agarised rooting medium with black sterile polycarbonate granules. Olive rooting is also promoted by the addition of 1 mM putrescine to an auxin-containing rooting medium. This treatment promotes earlier rooting and increases both the rooting percentage and the number of roots per shoot (Rugini et al. 1997). Root induction by means of dipping method As an alternative to the traditional use of rooting media, shoots may be dipped into a liquid rooting solution and inserted directly into an auxin-free medium. Obviously, the auxin concentration is markedly higher in the dipping method, in comparison with the common system of using a rooting medium. Rugini and Fedeli (1990), for instance, reported high rooting percentages by dipping entire microcuttings, or only their basal parts, in an IBA solution (100–200 mg/L) for 10–20 sec. Because of its laboriousness, it is advisable to try the dipping method when gelled rooting media do not produce acceptable and consistent results.

5.1.5 Stage IV: Acclimatisation of plantlets After 3–5 weeks from the beginning of the rooting stage, rooted microplants (Plate XXVIII) are ready for in vivo acclimatisation. Aims of acclimatisation Acclimatisation is a critical point in micropropagation, due to the drastic change of climatic conditions (humidity, light intensity, asepsis) which characterises the passage from the in vitro to the in vivo environment. This problem is accentuated by the particular histology of leaves from in vitro culture, which makes them even more prone to desiccation, as well as poorly functional in the acquisition of autotrophic conditions. In the olive, for instance, micropropagated plants of ‘Nocellara del Belice’ and ‘Nocellara Etnea’ have leaves with a notable reduction of both thickness and central vascular bundle diameter when compared with in-field grown plants. The epidermal layer shows a thin cuticle, the stomatal density is low, and the palisade tissue is composed of only one layer, instead of the normal 3 to 4 layers. These differences are more accentuated in the ‘Nocellara del Belice’ cultivar (Cozza et al. 1997). Moreover, the in vitro-formed root apparatus has histo-anatomical characteristics which makes it not very functional when moved to compost substrates. Hence, the acclimatisation stage accomplishes the functions of (i) reducing as much as possible the stress resulting from the transfer of plantlets to in vivo conditions, (ii) facilitating the production of a new functional root apparatus, and (iii) allowing a gradual shift of plantlets from mixotrophic conditions (i.e. the typical in vitro condition where energy for growth is almost entirely obtained from sucrose, and only in a very limited part from photosynthesis) to autotrophic (i.e. entirely photosynthetic) conditions.

In vitro propa ga tion of the olive

How to acclimatise olive microplants High humidity is essential for successful acclimatisation. Once de-flasked, olive microplants should be treated carefully and moved to a protected environment as quickly as possible, because they can desiccate in a hot dry atmosphere. Moreover, hygienic precautions should be carefully maintained during the entire procedure of de-flasking. Hence, olive acclimatisation can be pursued through the following steps. 1. Microplants are removed from the flasks under the sterile air of the laminar airflow hood. It is recommended that when the microplants are rooted in agar, the gel should be gently washed away from the roots in lukewarm water, in order to avoid plantlet contamination due to sucrose and other organics trapped in the residual agar; additionally, the rooted part of the microplants can be dipped or soaked in a fungicidal or bactericidal solution. 2. Plantlets are then inserted into plastic or paper containers (pots, cells or trays), containing appropriate free-draining compost substrates. All containers should be new, or if previously used, scrupulously cleaned. Different mixtures of peat, vermiculite, perlite, soil and sand can be used to prepare a good compost for olive plantlet acclimatisation. For instance, a mixture of peat, soil and perlite (1:1:1) can be satisfactory. The material from which compost is prepared should have been sanitised or pasteurised to eliminate bacterial or fungal infections. 3. The plantlets are then transferred to a tunnel covered with transparent plastic film. As soon as they have been set out, the plantlets can be drenched or sprayed with a fungicide, so that both plantlets and compost are treated. The plantlets are then periodically watered to maintain an appropriate level of air humidity (RU = 85â&#x20AC;&#x201C;90%). However, the compost substrate should not be too wet during acclimatisation, as under such a condition algal growth is promoted and plants are liable to fungal and bacterial attacks. Plastic tunnels are the cheapest system for the acclimatisation of olive plantlets, but they can become overheated during the hot season and may need to be shaded (e.g. with a 50% shade cloth). Temperature and humidity should be monitored frequently. Alternatively, the plantlets can be acclimatised in a glasshouse under mist or fog conditions, misting requiring more careful control during initial acclimatisation to avoid over-watering of the substrate. 4. For the final hardening of rooted plantlets, relative humidity should be gradually reduced by opening the tunnel for increasing periods of time, or by reducing the frequency of fogging/misting. Following acclimatisation, a one-year period in the nursery is required before their final transfer to the orchard.

5.1.6 Field performance of micropropagated plants The first olive orchards established in Italy with micropropagated plants have recently started to bear fruit, from which a high-quality extra-virgin oil has been obtained. To date, few studies have dealt with the observation of the genetic and agronomic characteristics of micropropagated olive trees after in vivo acclimatation and in-field plantation. However, working with molecular markers, there is consistency in evidencing morphological and genetic true-to-typeness of in vitro propagated trees with regard to

Olive Propa ga tion

the donor genotype. Moreover, researchers agree that after in-field plantation, micropropagated olive trees do not exhibit juvenile traits and have a short unproductive period, as they start to bear fruit in the 2nd/3rd growing season. Over two years of juvenility was observed only in plants micropropagated from sucker explants or from ‘Canino’ somatic embryos (Rugini, pers. com.)

5.2 Somatic embryogenesis From the 1980s onward, great efforts have been made to develop procedures of olive regeneration other than axillary bud stimulation, such as somatic embryogenesis (see box). An important reason for these studies was to explore non-conventional methods of genetic improvement, i.e. transformation by Agrobacterium, somaclonal variation, protoplast manipulation or microprojectile bombardment (Lambardi et al. 1999a; Rugini et al., 2000). In this context, it is fundamental to apply bioengineering techniques to in vitro culture systems characterised by high morphogenetic potential in order to get plant regeneration from manipulated cells. Moreover, the olive can take advantage of efficient regeneration techniques (such as somatic embryogenesis) in other ways, i.e. (i) by overcoming the problem of non-rootability of some cultivars, which requires laborious grafting procedures, and (ii) with the production of synthetic seeds (see 5.3). Although somatic embryogenesis from very young material (i.e. from zygotic embryos and seedling explants) has been repeatedly reported, this approach is of little interest in terms of both propagation and transformation of olive cultivars. Therefore, we report here only of somatic embryogenesis initiated from mature tissue explants, that is when the explants come from adult plants (in-field, or previously reproduced by micropropagation). SOMATIC EMBRYOGENESIS The term refers to the development of a bipolar structure (= a complete embryo) from somatic cells, i.e. from the cells of an explant cultivated in vitro (‘direct somatic embryogenesis’), or from de-differentiated cells (= callus cells) originated from the original explant (‘indirect somatic embryogenesis’). This latter system is the most common when working with woody plants, and it involves the culturing of explants with strong growth regulators (such as 2,4dichlorophenoxyacetic acid or 2,4-D) to induce the production and proliferation of ‘embryogenic lines’, i.e. callus lines which are competent for the formation of somatic embryos. A certain amount of genetic variability (‘somaclonal variation’) may be inherent in this system of plant reproduction. Consequently, careful field monitoring for variability is required when somatic embryogenesis is tested for large-scale plant production.

5.2.1 Somatic embryogenesis from mature tissue explants A cyclical system of somatic embryogenesis from mature olive tissue was described by Rugini and Caricato (1995), and subsequently indicated as a ‘double regeneration system’. The original explants were petioles excised from micropropagated shoot cultures, previously established from adult trees of the cvs ‘Canino’ and ‘Moraiolo.’ The sequential steps required for the regeneration of plantlets by somatic embryogenesis were the following:

In vitro propa ga tion of the olive

1. A small quantity of embryogenic callus was induced from the petioles cultured in the dark on a gelled half-strength MS medium, supplemented with 30 µM TDZ, 0.54 µM NAA and 4% sucrose. 2. When the calli were subcultured on OM medium, supplemented with 0.5 µM 2iP, 0.44 µM BA, 0.25 µM IBA, 0.42 mM cefotaxime, 1 g/L casein hydrolysate and 3% sucrose, proembryonic masses were produced. 3. Primary somatic embryos differentiated from these masses when placed on filter paper soaked in liquid OM medium. 4. When primary somatic embryos were then subcultured on a gelled hormone-free OM medium containing 0.1% activated charcoal, a profusion of secondary somatic embryos were induced directly from the epidermal layer of the primary embryos, mainly from their basal parts (Plate XXIX), with no callus production (Lambardi et al. 1999b). Once established, this cyclical system of secondary somatic embryogenesis can be maintained for years by monthly subculturing (Plate XXX). 5. Perfect somatic embryos at the cotyledonary stage (Plate XXXI) were then isolated and germinated after their transfer into shaken liquid OM medium. In the embryogenic masses, together with a majority of perfect somatic embryos, several other forms could be recognised, such as fused embryos, claviform structures, embryos with fused cotyledons (Benelli et al. 2001a). These abnormal structures never showed any further development.

5.3 Synthetic seeds and micrografting Important advances have been made recently on synthetic seed technology. The synthetic seed (or ‘synseed’) is composed of an explant (in general, a bud or a somatic embryo) encapsulated in a drop of alginate. After its solidification, the synthetic seed can be handled, stored or germinated in a way similar to a normal seed, producing a shoot (when a bud is used) or a somatic seedling. Micheli et al. (1998) proposed a synthetic seed technique aiming towards the improvement of conventional micropropagation and the application to germplasm preservation. After encapsulation in an alginate matrix containing a nutritive medium, apical and nodal buds from micropropagated shoot cultures (cv Moraiolo) maintained up to 49% viability and regrew satisfactorily after storage at 4°C for 45 days. However, as rooting was unsatisfactory and restricted only to microcuttings formed from apical bud beads, the technique seems still a long way from being applicable in practice. Recently, olive somatic embryos (cv Canino) have also been used for synthetic seed preparation (Plate XXXII). The feasibility of olive in vitro propagation by micrografting has been reported. Revilla et al. (1996) induced rejuvenation of mature olive trees (cv Arbequina) after cycles of shoot micrografting on in vitro-grown seedling rootstocks. Troncoso et al. (1999) cleftmicrografted uninodal explants (from in vitro-grown ‘Cañivano’ seedlings) on in vitro ‘Arbequina’ seedlings, prepared with a cut just under the basal pair of leaves. After 60 days, 85% grafting survival was achieved, which then dropped to 67% after microplant hardening. These results suggest that synthetic seeds and micrografting should be further explored as additional methods of olive propagation.

Olive Propa ga tion

5.4 Considerations on olive micropropagation Although the first attempts to propagate the olive in vitro occurred over 20 years ago, the commercial exploitation of this technology, mainly in Italy and Spain, is still in its early stages. There are two main reasons for this delay, in comparison with other fruit species: first, many important olive cultivars were very difficult to introduce and establish in vitro; second, the stimulation of axillary buds and the subsequent shoot elongation required the presence in the medium of high concentrations of zeatin, a costly cytokinin which contributes considerably to raising the final cost of micropropagated plants. Since the mid-1990s, the number of cultivars that could be successfully propagated in vitro has risen considerably, and only a minor number is today still considered recalcitrant (among them important ones, such as ‘Leccino’, ‘Picholine’, ‘Kalamata’). Moreover, numerous medium constituents (e.g. specific macro and microelements, mannitol, cytokinins other than zeatin, auxins, polyamines) have been successfully tested to improve the various steps of micropropagation, with the goal of economising protocols. As a consequence, olive micropropagation is now competitive and, for some cultivars, can be proposed as more economical than traditional cutting propagation. At the same time, significant progress has also been made in the development of other regeneration systems, such as synthetic seeds and somatic embryogenesis. In this latter system, knowledge of important aspects, such as embryogenic potential of tissues, effect of exogenous growth regulators, histological origin and development of somatic embryos, has been acquired, opening the door to a more ends-directed exploitation of the technique for olive propagation and genetic improvement.

5.5 In vitro conservation of the olive Tissue culture offers important applications to the medium- and long-term preservation of plant germplasm. The possibility of in vitro storage of olives has only recently been explored, with promising results. Recent advances in ‘slow growth storage’ and cryopreservation are concisely summarised here.

5.5.1 Slow growth storage Slow growth storage refers to the maintenance in vitro of shoot cultures at low temperatures (in general 4–5°C), i.e. in conditions of reduced cell metabolism. Lambardi et al. (2002a) showed that, when an 8h photoperiod (25 µmol m-2 s-1 photosynthetically active radiation) was applied, shoot cultures of ‘Leccino’ and ‘Frantoio’ could be stored for no longer than 4 months at 4°C in gas-permeable containers. In contrast, when shoots were stored in the same conditions but in the dark, after 8 months 80% (‘Frantoio’) and 90% (‘Leccino’) of the shoots regrew satisfactorily and produced new nodes similar to those of unstored shoots. Gardi et al. (2001) stored shoot cultures of ‘Ascolana Tenera’, ‘Frantoio’ and ‘Moraiolo’ at 6°C and in the dark. ‘Frantoio’ was the one that performed best, as 100% survival and regrowth could be obtained after 5 months of conservation, while both the other cultivars maintained this maximum regrowth potential only up to 2 months.

In vitro propa ga tion of the olive

5.5.2 Cryopreservation Promising results have recently been obtained with the application of cryogenic techniques to the long-term conservation of olive germplasm (Lambardi et al. 2002b). Martinez et al. (1999) obtained 30% survival of olive shoot tips (cv Arbequina) following the removal of up to 30% of their moisture content, direct immersion into liquid nitrogen (–196°C), and rewarming at room temperature. Lambardi et al. (2002a) applied a procedure of vitrification and one-step freezing in liquid nitrogen (Fig. 5.6) to shoot tips excised from in vitro-grown shoot cultures of the cv Frantoio. Following re-warming at 40°C and planting in a regrowth medium, 15% survival rate was achieved, but only from shoot tips which had been obtained from apical buds. The surviving shoot tips remained green and started regrowth 4 weeks after planting. In contrast, the application of encapsulation-dehydration procedures was not effective in the protection of either ‘Frantoio’ (Benelli et al. 2001b), or ‘Arbequina’ (Martinez et al. 1999) explants during ultra-rapid freezing. As an alternative to shoot tips, embryogenic cultures of olive proved to be a highly suitable material for cryopreservation using the vitrification approach (Lambardi et al. 2002a, 2002b).

Figure 5.6 A 35-litre container, used for germplasm preservation in liquid nitrogen (–196°C).

Olive Propa ga tion

Table 5.2 Disinfection procedures using sodium hypochlorite (NaClO) and/or mercuric chloride (HgCl2) of apical twigs or nodal segments from different stock plants for the initiation of olive micropropagation. Concentrations of NaClO refer to a commercial product having 6% of active chlorine. Stock plants

Disinfection procedure (*)

Reference

Greenhouse-grown plants

1. Treat with 11% NaClO + 0.035% HgCl2 (3–5 min) under vacuum 2. Rinse 30–40 min in sterile water

Rugini & Fedeli 1990

In-field grown young trees

1. Treat with 0.2% HgCl2 (20 min) under vacuum 2. Rinse several times in sterile water

Briccoli Bati & Lombardo 1995

In-field grown old trees

1. Treat with 0.1% HgCl2 (5 min) 2. Rinse once in sterile water 3. Treat with 17.5% NaClO (15 min) 4. Rinse three times in sterile water

Martino et al. 1999

few drops of a surfactant (e.g. Tween 80) are always added to the disinfectant solutions.

References Baldini, E. 1986. Arboricoltura generale. CLUEB, Bologna (Italy), pp. 396. Bartolini, G., Leva, A. R. & Benelli, A. 1990. Advances in in vitro culture of the olive: propagation of cv Maurino. Acta Hortic. 286: 41–44. Benelli, C., Fabbri, A., Grassi, S., Lambardi, M. & Rugini, E. 2001a. Histology of somatic embryogenesis in mature tissue of olive (Olea europaea L.). J. Hortic. Sci. Biotech. 76: 112–119. Benelli, C., De Carlo, A., Lambardi, M. & Lynch, P.T. 2001b. Vitrification of shoot tips, nodal segments and embryogenic tissue of olive (Olea europaea L.) for germplasm cryopreservation. Acta Hortic. 560: 137–140. Briccoli Bati, C. & Lombardo, N. 1995. Propagazione in vitro della cv Nocellara etnea. Proceedings ‘L’Olivicoltura Mediterranea: stato e prospettive della coltura e della ricerca’. Cosenza, Italy, January 26–28, pp. 249–257. Briccoli Bati, C., Librandi, A. & Sirianni, T. 1994. In vitro propagation of two olive cultivars. Abstracts ‘VIIIth International Congress of Plant Tissue and cell Culture’. Firenze, June 12–17, p. 72. Cañas, L. A., Avila, J., Vicente, M. & Benbadis, A. 1992. Micropropagation of Olive (Olea europaea L.). In Y.P.S. Bajaj (ed.) Hightech and Micropropagation I. Biotechnology in Agriculture and Forestry, Vol. 17. Springer, Berlin Heidelberg New York, pp. 493–505. Cassels, A.C. & O’Herlihy, E.A. 2003. Micropropagation of woody trees and fruits: pathogen elimination and contamination management. In S.M. Jain & K. Ishii (eds) Micropropagation of Woody Trees and Fruits. Kluwer Ac. Pub., Dordrecht, pp. 103–128. Chaari-Rkhis, A., Trigui, A. & Drira, N. 1999. Micropropagation of Tunisian cultivars olive trees: preliminary results. Acta Hortic. 474: 79–81. Chaari-Rkhis, A., Maalej, M., Chelli Chaabouni, A. & Drira, N. 2002. ‘Meski’ olive variety propagated by tissue culture. Acta Hortic. 586 (vol. 2): 871–874. Cozza, R., Turco, D., Briccoli Bati, C. & Bitonti, M.B. 1997. Influence of growth medium on mineral composition and leaf histology in micropropagated microplants of Olea europaea. Plant Cell Tiss. Org. Cult. 51: 215–233. Dimassi-Theriou, K. 1994. In vitro propagation of cv. Kalamon olives (Olea europaea sativa L.). Adv. Hort. Sci. 8: 185–189. Driver, J.A. & Kuniyuki, A.H. 1984. In vitro propagation of Paradox walnut rootstock. HortScience 19: 507–509. Garcia-Fèrriz, L., Ghorbel, R., Ybarra, M., Mari, A., Belay, A. & Trujillo, I. 2002. Micropropagation from adult olive trees. Acta Hortic. 586 (vol. 2): 879–882. Gardi, T., Micheli, M., Piccioni, E., Sisani, G. & Standardi, A. 2001. Cold storage of micropropagated shoots of some fruit species. Italus Hortus 8 (4): 32–40 (with English abstract). George, E.F. 1993. Plant Propagation by Tissue Culture. Part 1, The Technology. Exegetics Lim., Edington (UK), pp. 560.

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Hartmann, H.T., Kester, D.E. & Davies, F.T. 1990. Principles of propagation by seed. In Plant Propagation, Principles and Practices (5th edn). Prentice Hall, New Jersey, pp. 104–136. Heller, R. 1953. Studies on the mineral nutrition of in vitro plant tissue cultures. Ann. Sci. Nat. Bot. Biol. Vég. 11th Ser. 14: 1–223. Lambardi, M. & Rugini, E. 2002. Micropropagation of olive (Olea europaea L.). In S. M. Jain and K. Ishii (eds) Micropropagation of Woody Trees and Fruits. Kluwer Ac. Pub., Dordrecht, pp. 621–646. Lambardi, M., Amorosi, S., Caricato, G., Benelli, C., Branca, C. & Rugini, E. 1999a. Microprojectile-DNA delivery in somatic embryos of olive (Olea europaea L.). Acta Hortic. 474: 505–509. Lambardi, M., Caccavale, A., Rugini, E. & Caricato, G. 1999b. Histological observation on somatic embryos of olive (Olea europaea L.). Acta Hortic. 474: 67–70. Lambardi, M., Benelli, C., De Carlo, A., Fabbri, A., Grassi, S. & Lynch, P.T. 2002a. Medium- and longterm in vitro conservation of olive germplasm (Olea europaea L.). Acta Hortic. 586 (vol. 1): 109–112. Lambardi, M., Lynch, P.T., Benelli, C., Mehra, A. & Siddika, A. 2002b. Towards the cryopreservation of olive germplasm. Adv. Hort. Sci. 16(3–4): 165–174. Leva, A.R., Petruccelli, R. & Bartolini, G. 1994. Mannitol ‘in vitro’ culture of Olea europaea L. (cv. Maurino). Acta Hortic. 356: 43–46. Martinez, D., Arroyo-Garcia, R. & Revilla, A. M. 1999. Cryoconservation of in vitro grown shoot-tips of Olea europaea L. var. Arbequina. CryoLetters 20: 29–36. Martino, L., Cuozzo, L. & Brunori, A. 1999. Establishment of meristem tip culture from field-grown olive (Olea europaea L.) cv. Moraiolo. Agr. Med. 129: 193–198. McCown, B.H. & Lloyd, G. 1981. Woody plant medium (WPM). A mineral nutrient formulation for microculture of woody plant species. HortScience 16: 89. Mencuccini, M. 1998. Micropropagazione della cv FS–17. Olivo & Olio 6: 35–39. Micheli, M., Mencuccini, M. & Standardi, A. 1998. Encapsulation of in vitro proliferated buds of olive. Adv. Hort. Sci. 12: 163–168. Murashige, T. 1974. Plant propagation through tissue cultures. Ann. Rev. Plant Phys. 25: 135–166. Otero, M. L. & Docampo, D.M. 1998. Micropropagation of olive (Olea europaea L.) cv. Arbequina from juvenile cuttings. Phyton 63: 133–140. Rama, P. & Pontikis, C.A. 1990. In vitro propagation of olive (Olea europaea sativa L.) ‘Kalamon.’ J. Hort. Sci. 65: 347–353. Revilla, M.A., Pacheco, J., Casares, A. & Rodriguez, R. 1996. In vitro reinvigoration of mature olive trees (Olea europaea L.) through micrografting. In vitro Cell. Dev. Biol. -Plant 32: 257–261. Rugini, E. 1984. In vitro propagation of some olive (Olea europaea sativa L.) cultivars with different root-ability, and medium development using analytical data from developing shoots and embryos. Sci. Hortic. 24: 123–134. Rugini, E. & Caricato, G. 1995. Somatic embryogenesis and plant recovery from mature tissues of olive cultivars (Olea europaea L.) ‘Canino’ and ‘Moraiolo’. Plant Cell Rep. 14: 257–260. Rugini, E. & Fedeli, E. 1990. Olive (Olea europaea L.) as an oilseed crop. In Y. P. S. Bajaj (ed) Legumes and oilseed crops I. Biotechnology in Agriculture and Forestry, Vol. 10. Springer, Berlin Heidelberg New York, pp. 593–641. Rugini, E. & Fontanazza, G. 1981. In vitro propagation of ‘Dolce Agogia’ olive. HortScience 16: 492–493. Rugini, E., Biasi, R. & Muleo, R. 2000. Olive (Olea europaea var. sativa) transformation. In S.M. Jain & S.C. Minocha (eds) Molecular Biology of Woody Plants. Vol. 2. Kluwer Ac. Pub., Dordrecht, pp. 245–279. Rugini, E., Jacoboni, A. & Luppino, M. 1993. Role of basal shoot darkening and exogenous putrescine treatments on in vitro rooting and on endogenous polyamine changes in difficult-to-root woody species. Sci. Hortic. 53: 63–72. Rugini, E., Di Francesco, G., Muganu ,M., Astolfi, S. & Caricato, G. 1997. The effects of polyamines and hydrogen peroxide on root formation in olive and the role of polyamines as early marker for rooting ability. In A. Altman and Y. Waisel (eds) Biology of Root Formation and Development. Plenum Press, New York, pp. 65–73. Seyhan, S. & Özzanbak, E. 1994. Shoot multiplication of some olive (Olea europaea L.) cultivars. Acta Hortic. 356: 35–38. Troncoso, A., Liñan, J., Cantos, M., Acebedo, M.M. & Rapoport, H.F. 1999. Feasibility and anatomical development of an in vitro olive cleft-graft. J. Hort. Sci. Biotech. 74: 584–587.

6 The olive nursery (stock plants, structures, equipment and operations)

In order to carry out all activities concerning the production of self-rooted and grafted plants in the plant nursery, several structures are necessary such as: greenhouses with different purposes (propagation, plant growing, etc.), warehouses to store various materials (substrates, containers, chemicals, etc.), garage and repair shop, etc. Moreover, the nursery should have a specific plot in which to grow and manage stock plants (i.e. genetically and sanitarily controlled stock plants, from which to collect scions and cuttings). These structures should have size and characteristics proportional to the average plant number to be produced and to the foreseeable duration of the nursery itself. Deciding the proper size of a nursery to be created from scratch is not easy, as many and sometimes peculiar factors have to be taken into account. Here, basic technical information is given for establishing a small- to medium-sized olive nursery. For larger enterprises, a thorough evaluation of the site, the economical and market conditions, and available technical options should be made by expert professionals (for a comprehensive description of facilities and equipments of the nursery, see Nelson 1985). Whether the nursery is to operate in a warm, temperate or cold climate means that there will be differences in plant growth rhythms, and at times there will even be different techniques used. All this implies, among other things, differences in capital investment. The basic data needed to determine the size of an olive nursery, in a Mediterranean type environment, are as follows: 1. Greenhouse: one m2 of rooting bed may host 500â&#x20AC;&#x201C;700 cuttings, and can be used at least twice a year; the average rooting percentage can be, considering the cultivars propagated by cutting, in the 40â&#x20AC;&#x201C;80% range. To produce 10 000 plants per year, about 20 m2 of rooting bed is required, together with about 100 m2 of hardening greenhouse, and 1000â&#x20AC;&#x201C;2000 m2 of lath-house or non-shaded area, to allow plant growth before planting or marketing.

The olive nurser y (stock plants, structures, equipment and opera tions)

2. Water: to serve a medium-sized nursery a small reservoir to store water is important, whatever the origin, or alternatively a source supplying at least 30 litres/minute. 3. Stock plants: vegetative growth is greatly influenced by environment and management practices; on average, at the 3rd–4th year from planting the stock plant is able to produce about 30–60 shoots suitable for cuttings preparation, both in spring and autumn; as each shoot is usually able to yield two cuttings, the overall production is about 200 cuttings/plant/year. In the following years the ability of the tree to produce shoots of a good quality increases to 300–500 cuttings/tree/year. Taking rooting percentages into account, the production of 10 000 plants/year requires not less than 40 stock plants. If the stock plants are managed with different techniques, such as container growing with fertigation, or hydroponics, the number of plants needed for the same production of wood rises to 60–100, due to the smaller tree size (see 6.1). If normal bearing plants are chosen as the source of wood for cuttings, their canopies being far larger, the amount of available material is higher if compared to ad hoc stock plants. Suitable wood is collected during annual pruning operations, which ordinarily take place at the end of winter, and lasts until the beginning of inflorescence emergence (in the Mediterranean basin from January through April). However, further collection of wood should not be made during the year, to safeguard amount and regularity of production, since the best shoot for cutting production is also the wood that is likely to bear fruits the next season. 4. Substrates: perlite is used for propagation, and it requires no particular preparation. However, the substrates to be used for transplants of rooted cuttings, or for successive transplants in the nursery must be prepared beforehand. To this end, a platform or a shed should be planned in the vicinity of the propagation, hardening and growing greenhouses, in which to mix (by hand, with concrete mixers, or with other suitable devices) and store a few cubic metres of the individual components and of the mixtures needed for the various transplants. A few thousand pots should be stored in the same place, of at least two sizes: 400–600 mL for the first transplant, 2–3 litres for the second. 5. Sundry materials: stakes for the plants that reach the second transplant (bamboo or plastic sticks), tying machines and tying materials, or tubelets that affix plants to supports, tags of different colours, secateurs for cutting preparation, wax, mastic or similar products, tying materials and knives for grafting.

6.1 Stock plants Stock plants have various uses: • as a source of material for propagation (subject to periodic health and genetic checks) • as a means of conserving genetic resources • for genetic improvement

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As they are the source of material for propagation (cuttings, buds, microcuttings, seed), controls must be carried out to ensure that they maintain good health and genetic trueto-typeness. Nurseries producing olive trees, as with those producing other fruit trees, need to use ‘controlled’ material for propagation which corresponds to the genetic and morphological characteristics set out in the certification procedure (see 6.8). The methods used in the growing of stock plants vary according to the ‘product’ to be supplied for propagation: seed, shoots for grafting and cuttings, buds and microcuttings for micropropagation.

6.1.1 Phase change Morphological and physiological traits expressed in different parts of the same plant may follow a particular pattern depending on the developmental age at which the cells are laid down (‘phase change’). The shift from vegetative to reproductive is known as ‘maturation’. The condition of a seedling before maturation is also called ‘juvenility’, and it involves a number of morphological, anatomical and physiological differences from the adult tree. The most visible characteristic of young seedlings is their incapacity to produce flower buds. Other morphological characteristics in the olive are thorniness, smaller leaf size, shorter internodes, darker leaf colour, etc. But the most important feature of juvenile material is the ability to regenerate, and the ease with which it produces adventitious roots. Material for cutting propagation should never be taken from seedlings, as the resulting trees would be of unknown characteristics, and all positive features fixed by cloning would be lost; besides, those trees would take several years, possibly more than a decade, to produce the first flowers, as this is the duration of the juvenile stage in the olive. Still, the reversion to a less adult condition in adult trees, and therefore the acquisition of a partial juvenile condition, may occur in mature tissue on certain occasions. As this phenomenon, called rejuvenation, may involve an improvement in the rooting ability of cuttings, studies have been devoted to the issue, and methods set up. These are: • vigorous pruning to stimulate strong and rapid growth. The planting of trees so as to form a hedge, as is done with many fruit trees, is not suited to olive trees because the development of adventitious shoots on branches older than 3–4 years may be insufficient or erratic in many cultivars. • the use of growth regulators. Little has been reported on the use of this class of compounds with olive stock plants. Gibberellins seem to act as positive growth stimulants in plants and to enhance rooting in cuttings (Fontanazza & Rugini 1977).

6.1.2 Training systems for stock plants Not much has been reported on this aspect. Experience acquired to date is neither abundant nor definitive. It would seem that it is best to place the plants at minimum distances of about 2.0 x 2.5 m (Fig. 6.1), to make mechanised operations possible. The lowest branch height is established at 80–90 cm from the first year. From the second year, when propagation material is being taken, shoots are excised while leaving the two basal

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buds (nodes) as spurs, and leaving untouched all the shoots that have not grown sufficiently. Thus future development is aided by avoiding, as far as possible, an imbalance between development below and above ground. If there is excessive vegetative growth it is advisable to carry out normal cutbacks. Trees pruned this way acquire their definitive shape after 4 to 5 years, at which point they will each produce over 200 cuttings annually (see above).

Figure 6.1 Stock olive plant plot in an experimental olive farm in India (Kullu, Himachal Pradesh).

6.2 Greenhouses The main site for propagation is the greenhouse. Greenhouses are structures, covered with transparent materials to allow natural light to penetrate, in which plants grow. These structures are usually artificially heated, and are quite different from cold and warm frames. They also differ from other types of structures because they are relatively high, to permit comfortable work and, in some cases, the use of agricultural machinery. The greenhouses can be fixed or mobile. The former cannot be disassembled or moved. The latter are made in such a way as to allow their removal at different times of the year to different plots, or their complete disassembling. These mobile greenhouses can overcome the problems of specific replant disease, and of infestations of several soil pathogens; obviously, such problems do not arise with container cultivation. Greenhouse covering material should: â&#x20AC;˘ reduce the passage of solar radiation as little as possible, with special reference to the visible part of the light spectrum; this aspect is not important for the rooting zone.

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• be sufficiently transparent to short infrared radiation, and not allow the passage of long infrared rays, reflected by the soil and by the plants. Maximising the above allows the storage of a fair part of the sun’s energy as heat, resulting in what is commonly termed the ‘greenhouse effect’. The first material used to cover the greenhouses was glass. In recent decades several plastic materials, both as films or as rigid panels, have been introduced.

6.2.1 Glass greenhouses Until 1950, glass was the only transparent material used to cover greenhouses (Fig. 6.2); it is now one of the most expensive covers. The use of glass allows the building of any greenhouse structure, any automation, and the adaptation to cold and snowy climates. The structure of glass greenhouses can be wood (usually not wider than 6–8 meters) or metal (painted or zinc-plated steel, aluminium, usually 12–15 metres wide, and sometimes wider). Today many firms build greenhouses from prefabricated parts.

Figure 6.2 Series of traditional glass greenhouses (Davis, University of California, USA).

The properties of glass depend on its nature and composition, its homogeneity, its colour and thickness, and on the profile of its surfaces, which can be smooth on both sides, or with one smooth side only, the other being left rough. In this latter case the smooth side of the pane should be the external one, to cause minimal reflection losses, which are about 14% when the sun is low on the horizon. The smooth glass can be of a ‘simple’ thickness (1.6–1.9 mm), or ‘double strength’ (2.7–3.2 mm). Raw glass includes the ‘gardener’, ‘printed’, ‘hammered’, and ‘lined’ types. It is also possible to find more resistant types of glass – ‘tempered’, ‘armed’ (with a metal mesh inside) – able to satisfy all security needs for greenhouse operation.

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6.2.2 Plastic film greenhouses Many types of flexible plastic films (mylar, polyvinyl chloride, polyethylene, polycarbonate) have been used as greenhouse covers. Polyethylene is today’s most used compound; it is cheaper than glass, has a low specific weight, and can be adapted to any greenhouse shape. It is impermeable to water and to water vapour, but is permeable to gases, particularly carbon dioxide and oxygen. Polyvinylchloride transparency is similar to that of glass, and remains so even with ageing of the material, allows a better thermal performance, and is more permeable to water vapour than polyethylene. Due to the presence of ultraviolet rays (UV), the exposure of plastic films to direct light limits their longevity to 1–5 years, according to plastic type and thickness. Plastic films make possible the creation of double walls, where the empty space is filled with forced air. This technique permits a reasonably effective thermal insulation of the greenhouse, and overcomes small sudden temperature drops.

6.2.3 Rigid-panel greenhouses PVC (polyvinylchloride) PVC panels are much less important today than in the past. Initially, they had a low cost and a ‘life’ of five or more years, although they could be profitably used for little more than two years. Light transmission rapidly decreases with time, and more so when they are lined. They are available in many colours, although the colourless ones are the most used. Not being completely rigid, PVC panels can adapt to any greenhouse type (Fig. 6.3).

Figure 6.3 Hardening greenhouses, built with corrugated PVC.

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FRP (fiberglass-reinforced plastic) This type of material for greenhouse covering was also used much more in the past. As with PVC, corrugated panels were used for their great resistance, while flat panes have had limited use . They show a remarkable resistance to factors that affect their ‘life’. Light transmission through the transparent panels is similar to that of glass, while heat loss is slightly inferior. The cost is very close to glass covers. Polycarbonate, methacrylate, and other materials These materials have been widely used in the last few years, and are marketed as panes, undulated sheets, or with air spaces. The latter type, of different sizes and thickness, is particularly suited to maintain the thermal status of the greenhouse, with resulting energy savings. It has a noteworthy mechanical resistance and long life, of at least ten years. The low weight of these materials allows cost reduction for the supporting structures of the greenhouse, while at the same time the safety conditions for those who operate inside the structure are improved, compared to the glass equivalent.

6.2.4 Location of the greenhouse Many factors need to be considered in the location of the greenhouse. Important among these are quantity and quality of plants, dimensions of the market, degree of automation wanted, and availability of skilled labour. The following aspects should be carefully considered. • Space for ancillary structures: the place in which the greenhouse is located should also accommodate areas with general services for the greenhouse, such as office buildings, warehouses, shelter for machinery, and storage of substrates, pots and fertilisers. If the greenhouse is to be used for the production of potted plants, or for propagation by cuttings, there must be a site nearby (building, greenhouse, or other) where the potting operations can be carried out, in a more or less mechanised way. • Topography: the greenhouse site should allow for the construction of large greenhouses, if necessary made of modules built over a given period of time, with all connections ready for the installation of automatic devices. The greenhouse should be located on well-drained soil, as should the rest of the nursery. If prevailing winds are present, natural windbreaks should be planted, if necessary in multiple rows. In areas where snowfalls are likely, trees should not be planted near the greenhouses. • Climate: climate is an extremely important factor in the choice of the greenhouse construction site. For propagation purposes, a climate with moderate thermal variations is necessary, in order to avoid or to reduce to a minimum the use of very expensive technology for cooling or heating. A climate in which environmental humidity is never too low is to be preferred; otherwise the problem can be overcome by adopting the ‘fog’ system in the greenhouse section in which propagation is performed, or by using this system together with the ‘mist’ on the propagation benches. Particularly windy locations are to be avoided, because construction would become too expensive. Excessive levels of solar radiation are

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easily manageable with the use of shade cloth; if they are naturally too low, such a location should not be used as plants will tend to grow too tall and thin. Labour: workers should be easily available; sites too distant from towns and facilities can pose problems. Equipment breakdowns are frequent, and can severely reduce production: interruptions of mist for two hours or more, due to power failures or other reasons, can compromise the rooting success of cuttings. The list of ‘accidents’ that can take place in a nursery is very long. Hence, the advice is to double up on all those devices controlling ‘delicate’ points of the nursery, in order to overcome temporary crises; in this case, the choice can be for less sophisticated or expensive solutions. Accessibility: the greenhouse must be built on a site easily accessible by a road that is connected to the main network. All vehicles must have easy access to the greenhouse. Water: water is a limiting factor, both qualitatively and quantitatively. In the nursery site the supply of water, necessary to the various sectors such as greenhouses, shaded areas and growth sectors, must always be easily accessible. A consumption of 20 L/m2/day in the growth sector, supplied 2–3 times per week, should be expected; the nursery’s water reserves will therefore need to be adequate for the needs of the greenhouses and growth areas (50–80 m3 as reserves, and a constant supply of about 30 L/min). Water quality is also an important factor. Using excessively saline water – i.e. high in sodium chloride (brackish waters) – requires the use of expensive desalting systems. Also the presence of high amounts of other salts (mainly carbonates and sulphates) requires the use of decalcifiers or other systems to reduce their concentration. Orientation: the choice between north–south and east–west orientation for the greenhouse is important, with reference to its structure, which can determine more or less pronounced shading, especially light transmission occurring in the winter (in the summertime the problem does not exist).

6.2.5 Greenhouse heating In greenhouse management, one of the most difficult problems is the control and maintenance of the appropriate temperature. The temperature requirements for rooting fluctuate widely in the course of the biological cycle of the olive plant, and the natural ambient temperature changes continually. In propagation greenhouses there is also the need to be able to modulate the temperature of the rooting bed at the cutting base with respect to the surrounding environment. The latter is kept at a much lower temperature to avoid bud break before root emergence. Heating can be produced with systems using hot water or air. Water and air heating is obtained with: • • • •

oil or methane burners, or by burning different types of wood solar panel systems or heat pumps electricity geothermal steam

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The use of solar panels and heat pumps is still far from being economically advantageous because of high establishment costs, although they may be considered as a support for traditional systems such as methane and oil. Whatever the heat source, the best policy is that of investing sufficient financial resources in the beginning to obtain constantly low heating costs. Water heat distribution is made with several systems, the most widespread being radiators and, if necessary, forced water circulation. Heated air systems are under either forced (Fig. 6.4) or free circulation; humidifiers check and adjust the relative humidity of the air if it drops below a minimum level. Direct use of electricity as a heat source is limited because of costs, and limited to emergency systems or localised heating. In this case interesting perspectives have been opened by the marketing of low voltage (24 V) ‘electric Figure 6.4 Fan positioned between a rooting area and an blankets’, adjustable to the bottom of intermediate room of the greenhouse. any bed, and electric heating ‘cables’ (24 V), to be inserted below the soil level or inside the floor to raise the temperature at the base of pots. Both systems are long lasting and very resistant. Finally, electric generators should be considered whenever the nursery investments are high (elevated productions), the weather conditions are frequently adverse, and prolonged (over 1 hour) power failures are common.

6.2.6 Greenhouse cooling To reduce the high temperatures that often occur when sun radiation is continuous (greenhouse effect), especially in the hot season, the most common solution is shading the greenhouse with white paint or with shading nets. Such systems are not sufficient in hot climates, or whenever the outdoor temperature is high (i.e. above 30°C) and constant. Alternative solutions are: • ventilation: this can be natural, and obtained by opening lateral windows, both on the roof and along the walls. Simple automatic systems can control their gradual opening (Fig. 6.5). Ventilation can also be forced using electric fans placed on the roof and along the walls.

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Figure 6.5 Natural ventilation obtained by opening all windows and doors.

Figure 6.6 A cooling system based on water evaporation.

â&#x20AC;˘ a cooling system: this is a system of forced ventilation, based on air cooling facilitated by water evaporation (evaporating water absorbs the heat in the air inside the greenhouse). The system consists of a pad of porous material placed in a

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metal mesh structure and kept moist by water dripping from above (Fig. 6.6). In the facing wall, fans are inserted that expel the air from inside the greenhouse; in this way, a light air draft is created which passes through the wall containing the pad and is thus highly moistened. The airflow is then able to ‘absorb’ excess temperature. • air conditioning: ambient cooling is made by refrigerated air conditioners, of a size and number in proportion to the ambient volume and to the temperature differential. Where heat pumps are used, these can be also used for cooling.

6.2.7 Computers for ambient control In order to optimise all parameters within the greenhouse, it is possible to control window opening, bench and ambient heating, and air humidity by means of a computer which, where necessary, can manage more than one greenhouse at the same time. The same system can also manage outdoor operations, such as irrigation and fertigation. Installation of these systems is not profitable for small nurseries. For medium-sized nurseries the running economics are tied to continuous production (including species other than olives) or to high yearly production, over 200 000 plants.

6.2.8 Beds Beds, or benches, are structures whose surface serves several needs, such as propagation, support for pots and trays, and cultivation. It is worth remembering that all operations carried out on beds can also be made on the ground, with an obvious economical saving in construction costs. However, the greenhouse should be planned in such a way as to make working conditions as comfortable as possible, allowing the workers to operate as much as possible in an erect position. The beds can be fixed in brickwork or to a metal frame, or be in a mobile structure (zinc-plated steel or aluminium). More details on propagation beds are provided in the following paragraphs.

6.2.9 Propagation greenhouses Plant propagation, among the many activities taking place in the greenhouses, is the most complex and demanding one, from both a technical and an economic point of view. Usually the supporting structure of the greenhouse is zinc-plated steel, aluminium, or, more seldom, wood. The cover is glass panes or plastic materials. The greenhouse is usually divided into three zones, according to their most common use, i.e. the control zone, the rooting zone and the hardening zone. Control zone (instrumentation) The control zone takes up a few square metres, and contains the greenhouse environmental controls: window opening and closure, water regulation for mist and irrigation, temperature control (both in the beds and ambient), and water treatment. When the water contains over 500 mg/L of total salts a decalcification, filtration and demineralisation treatment must be operated, to bring the value down to 100–150 mg/L, necessary to avoid damage to the greenhouse equipment and to avoid the formation of a calcareous layer on the leaf surfaces, which would limit leaf functionality. In medium to

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large nurseries, equipped with advanced technology, the greenhouse control area should be located in a nearby building. Rooting zone This is the most important zone of the greenhouse. Here, the optimum conditions for the rooting of cuttings are created. In this zone are the ‘rooting beds’ (Fig. 6.7); if they are fixed, corridors 40–60 cm wide should be left, and larger passages of 60–80 cm are to be allocated if the number of beds is large and therefore the passage of cumbersome materials is to be expected. If movable beds are used, the surface for rooting cuttings is greater, and as a rule only space for one corridor should be left and allocated when planning the greenhouse. This is because when one bed is filled it can be pushed aside towards the wall with work passing on the second, which in turn will be pushed against the first when done, and so on, thus obtaining an uninterrupted block of beds. Compared to the brickwork and concrete of fixed beds, the use of movable beds involves the utilisation of more expensive materials. Aluminium, suited to very humid environments (zinc-plated steel being less expensive), always has a shorter life, and requires more careful maintenance. Rooting beds are of various widths, but usually never above 120–140 cm (i.e. two arms’ length) to allow comfortable operation. They should have a raised edge (about 10–12 cm to contain the substrate); if an edge is absent, rooting is obtained in containers of different type and size (usually plastic trays at least 10 cm deep, with a holed or net bottom, fixed or removable) which are placed on the bed.

Figure 6.7 Concrete rooting bed. Bed and nozzles can be covered with a polythene film.

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Whatever the system chosen for managing the cuttings in the greenhouse, either directly in the beds or in containers laid on them, the greatest care should be taken to avoid any waterlogging. The mist technique can be used throughout the year, but, when the outdoor temperature is below 15–20°C, it is less efficient and the addition of bottom heating of the cuttings becomes necessary in order to keep the lower ends of cuttings at a constant temperature range (18–25°C). Such heating systems, when placed at the bottom of the bed, are made of: • water radiators (plastic or copper pipes; Fig. 6.8) • low voltage electric resistances (24–48 V) • heating sheets, made of electric resistances, embedded in high thermal resistance plastic materials, which can be also used to sterilise the upper layers of the substrate

Figure 6.8 Copper pipes for bottom heating.

When the heating systems are placed below the beds they can consist of: • forced hot air • tubes with fins (hot water) Quite often, in order to improve the uniformity of temperature at the bottom of cuttings, and to favour a more rapid disposal of the rooting medium’s water, and a better air exchange, 5–8 cm of draining material (such as gravel or crushed stone) is interposed between heating surface and substrate (Fig. 6.9). Finally, the rooting medium can be laid on the draining material (Fig. 6.10), often

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Figure 6.9 Sheets of metal mesh being laid over gravel in a rooting bed.

Figure 6.10 The final layer of rooting substrate (perlite) completes the rooting bed.

Figure 6.11 Trays with perlite used as rooting bed.

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above a net to make its eventual removal easier, or in containers (boxes or trays of different shapes) with holes in the bottom. When the rooting and hardening zones have an enclosed heating system in the floor, containers (Fig. 6.11) can be placed on the floor (Fig. 6.12). In this case, beds become superfluous. A light pressure on the substrate will make it easier to insert the cuttings and keep them upright.

Figure 6.12 When trays are used, the use of a built-in heating system may improve rooting of cuttings.

Hardening zone This part of the greenhouse usually occupies the largest part of the surface area (Fig. 6.13). The rooted cuttings (rootlings) are placed here soon after potting, and stay in this sector for an acclimatisation period of variable length, from a few weeks to some months (1st transplant, see 6.5). The development of the root systems may be enhanced, especially in the cold season, by keeping the pots on heated surfaces (which can also be the greenhouse floor), while air heating can be kept to a minimum. Irrigation is performed with nozzles, allowing greater delivery capacity than those for mist; the same equipment can be used for fertigation.

6.2.10 Shelter and storage A medium-sized nursery (100 000 plants per year) needs a sheltered area of about 100 m2 for tools and equipment, including a workshop for minor repairs. The area available for storage of the various ingredients used to make up the potting substrate should be as large as possible, near the greenhouses and accessible to all transport vehicles. There should be a shelter for the equipment used to mix the ingredients of the potting compost and space for the storage of the latter (several cubic metres).

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Figure 6.13 Hardening greenhouse with potted plants.

A few square metres can be sufficient for the storage of the necessary pots. These should be sheltered, especially from the sun. Arrangements for the storage of pesticides must respect the safety regulations of the country concerned. Since these are generally poisonous or hazardous it is always wise to store them separately in a well-aired place which is not accessible to all, and with clearly visible warning signs describing the risks involved and regulations for use.

6.3 Substrates 6.3.1 Rooting substrates The rooting substrate should allow the cuttings to maintain an upright position, and permit the creation around the cuttingsâ&#x20AC;&#x2122; base of optimal conditions for adventitious root formation; that is, the right levels of moisture, aeration and temperature. During the time spent in the rooting medium the newly formed roots are not able to absorb nutrients, and therefore these substrates are not required to contain available mineral nutrients. To achieve these results, several materials have been used, alone or in mixtures (Fig. 6.14), the most common being perlite, peat, vermiculite and sand. Perlite is by far the most-used substrate. It is a grey-white siliceous material of volcanic origin, mined from lava flows. The crude ore is crushed and screened, then heated in furnaces to about 760Â°C, at which temperature the small amount of moisture in the particles changes to steam, expanding the particles to small, sponge-like kernels that are very light, weighing only 80 to 100 kg per cubic metre. Perlite is placed in trays of

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Figure 6.14 Examples of different substrates: 1, organic compost; 2, peat moss; 3, pumice; 4, perlite; 5, mixture of the four substrates.

various sizes where rooting by cuttings is obtained, or directly in the rooting bed, in layers 10–12 cm thick. This substrate is by far the most efficient at maintaining the same characteristics over time and in all rooting situations. It is marketed in two types of different grain size, 1–3 mm and 4–6 mm, the largest being the preferred type in olive nurseries. The physical and chemical features of the two types are identical: both hold three to four times their weight of water (more than enough for olive rooting needs), and pH is neutral; temperature is kept constant at the bottom of cuttings, with poor exchange with the surface. In given circumstances, water may have to be applied heavily to the substrate and the cuttings, e.g. when the temperature gets too high; in such instances water accumulates in the lower substrate layer, and careful and complete drainage should be ensured to avoid harmful waterlogging. In such situations it is recommended that perlite is laid on a metal net, with 2–3 mm mesh. The net size should be increased to 3–4 mm if it is made of plastic material. With a smaller mesh size, surface tension could keep the water from dripping through the mesh. The use of mixtures such as perlite added with peat, coconut fibre or vermiculite (in volume:volume ratios of 2:1 to 4:1) has given good results, at times better than perlite alone. However, such substrates are not always easily reproducible as ratios and composition (peat, for instance, can vary quite a lot in characteristics), and are certainly more expensive, because of the cost of materials and, above all, the labour required for preparation. Peat consists of the remains of aquatic vegetation which have been preserved under water in a partially decomposed state. The composition of different peat deposits varies widely, depending on the vegetation from which it originated, state of decomposition, mineral content, and degree of acidity. Peat moss is the least decomposed type of peat, and the most used in horticulture. It varies in colour from light tan to dark brown. It has a high moisture-holding capacity (15 times its dry weight), a high acidity (pH of 3.2 to

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4.5), and contains a small amount of nitrogen (about 1%) but no phosphorus or potassium. In olive propagation, peat is mostly used in the growing stage, and in mixtures to modulate its high acidity. Vermiculite is a micaceous mineral that expands markedly when heated; chemically, it is a hydrated magnesium-aluminium-iron silicate. When it constitutes the substrate, or forms part of a mixture, watering must be carefully checked, as water can be absorbed in excessive quantities that are harmful for the olive. Sand is not a good substrate when used alone, as it has a very poor water retention capacity, it is very heavy, and its particles may cause injuries to the cuttings when they are inserted in the substrate. It should preferably be fumigated or steam pasteurised before use, or at least washed, as it may contain weed seed and various harmful pathogens. It is used in mixtures when other materials are not available, or when costs have to be kept to a minimum. To conclude, whatever the materials used to prepare the substrate, it should: • • • • • • • • •

not interfere with the pH of the soil solution, which should stay neutral not allow waterlogging not let heat to rise to the upper layer of the substrate be pathogen-free allow gas exchange allow insertion of cuttings without damaging tissues allow cuttings to stand erect for the duration of propagation be cheap have low weight

6.3.2 Substrates for growing plants The production of this type of substrates (always mixtures) requires a platform or a shed near the greenhouse in which to mix (with concrete mixers or more dedicated machines) and store a few cubic metres of the various components and the substrates. At this stage, the substrate must meet almost all the criteria mentioned for the rooting substrates; but the root system should also gradually adapt to real soil conditions, and therefore nutrients appear in small amounts, and the physical features of the media get increasingly more similar to those of actual soil. This said, as a rule the choice should fall on what is most easily and cheaply available in the area in which the nursery is located.

6.4 Environmental conditions for rooting 6.4.1 Temperature Control of the temperature at the base of the cuttings and around the leaves is very important. Though the optimal temperature at the base of cuttings is 24 to 26°C, maintenance of such temperatures may sometimes be too expensive in the coldest months in terms of the running costs of a greenhouse. However, even 18 to 20°C is sufficient to obtain good results, although rooting may require a few extra days. Air temperature must be lower than that at the base of the cuttings, otherwise

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precocious bud-bursting can result and this would have a negative effect, because of (i) the resulting movement of nutrients at the expense of the roots, and (ii) moisture imbalance due to increased transpiration. High temperatures (50Â°C and above) in the rooting area of the greenhouse, which may occur in warm periods, need not be cause for great concern if care is taken to maintain the relative humidity of the air around the leaves at a level not lower than 95%.

6.4.2 Humidity The control of the humidity of the rooting environment is an extremely delicate matter with regard to the olive (leaves, base of cuttings, air above the rooting benches). The cuttings must be kept in optimal conditions to maintain the right degree of leaf turgor. Loss of leaves, especially in the first few days, compromises rooting. The right conditions are achieved by ensuring that a light film of moisture remains on the leaves and when this coverage is reduced to 40 to 50% it must be augmented with more water (mist or fog, see box â&#x20AC;&#x2DC;Water distribution techniquesâ&#x20AC;&#x2122;). Suggested watering times and intervals for olive cutting propagation are reported in Table 6.1. These conditions must be maintained for at least 1 to 2 weeks, after which it is possible to increase the time interval between waterings, as long as the percentage of leaf moist surface does not go below 40%. The leaves should never be allowed to become completely dry. WATER DISTRIBUTION TECHNIQUES Mist This method has been in use all over the world for the last 50 years. Water is supplied to the cuttings in very small drops which remain in the air for several seconds (Fig. 6.15). Water is nebulised using not very high pressure (kept constant at 4 to 6 atm in an autoclave) through nozzles positioned (depending on the type of nozzle, width of rooting bench etc.) 30 to 150 cm above the cuttings. Mist enables the maintenance for the required time of optimal amount of moisture on the leaves as well as in the substrate.

Figure 6.15 Olive semi-hardwood cuttings under mist.

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Figure 6.16 Olive semi-hardwood cuttings under fog. Fog This has not been used very much with the olive. The technique makes use of water in the form of a thick cloud of droplets which can remain suspended in the air for up to a few minutes (Fig. 6.16). This is possible with water at high pressure (about 30 atm) which must, however, be filtered and demineralised to avoid blockage of the nozzles. The method would be ideal for the maintenance of the right amount of moisture on the leaves but not in the substrate.

Table 6.1 Common durations of mist and intervals for olive semi-hardwood cuttings in intermediate environmental conditions. Week

Mist (seconds)

Interval (minutes)

15–12

10–12

12–10

12–15

10–7

15–18

7–5

18–20

5–12

18–20

The evaporation of the film of water on the leaves lowers the leaf temperature and consequently the rate of transpiration, which helps to create optimal conditions for rooting and survival. Care must be taken not to give too much water when maintaining moisture on leaves as this can lead to deposits of stagnant water in the substrate. In order to reconcile these opposing aspects, it is advisable to operate with a rooting environment that is reduced to a minimum volume. This can be achieved by keeping the roof of the

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greenhouse not too high, i.e. not more than 2 m at the eaves, or by making use of tunnels on the rooting benches (height not more than 1 to 1.5 m from the substrate surface). It must be remembered that the information given here is approximate, that not all greenhouses are the same and that experience acquired in one is not necessarily applicable in another. The choice of nozzles for misting depends on the width of the bench and the height at which the nozzles are placed. Different types of nozzles, for both mist and fog, are available, and used singly or together to create a denser mist. Higher pressures (about 30 atm) are necessary for fogging, as well as non-calcareous, filtered water to avoid clogging the nozzles, which have smaller holes than those used for misting. With fog alone, though, a very small amount of water is used, which is not enough to keep the substrate moist. It may therefore be necessary to alternate fogging with misting. There should be no draughts in the rooting environment. If any are created by the heating or cooling system, they must be very slight so as to avoid interference with evaporation from leaves and consequently with the mist. If there is excessive movement of air, the use of tunnels on the benches is necessary. These precautionary measures make it possible to maintain a high degree of relative humidity for long periods without excessive watering. In the first few days of rooting, relative humidity should not be allowed to go below 90 to 95%. After the first two weeks, the quantity of water must be progressively diminished in order to avoid deposits of stagnant water in the substrate, as this can cause root necrosis. If there is insufficient moisture on the leaves they soon drop off which, in turn, compromises the outgrowth of roots. These conditions must be maintained throughout the rooting period (about two months in warm climates and three in cold climates). The quantity of water supplied depends on the need to maintain a film of liquid on the leaves, and is regulated (watering times and the length of time between them) in various ways: timers (clocks), electronic leaves, scales, photoelectric cells and integrated computer systems (see box â&#x20AC;&#x2DC;Instruments for the control of water supplyâ&#x20AC;&#x2122;). Since the various techniques for water regulation are not perfect, they are unable to maintain constant optimal relative humidity; this quickly drops after each watering, thus causing a condition in which the substrate gets exceedingly moist, while the relative humidity needs frequent adjustments. These sudden changes in humidity are Figure 6.17 Rooting bed enclosed by a polythene cover. notably diminished when the rooting

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bench is closed in a tunnel of plastic sheeting (Fig. 6.17), either with or without an appropriate number of holes of a diameter of about 5 mm. In dry and/or hot climates the use of fog is a great help in keeping humidity more constant. INSTRUMENTS FOR THE CONTROL OF WATER SUPPLY Timers These provide the simplest method of controlling watering periods (in seconds: 1 to 60) and the intervals between them (in minutes: 1 to 60). Clocks are not a precise method of control because they do not take into consideration variations in atmospheric conditions (clouds/sun, heat/cold) which alter the humidity requirements of cuttings during the daytime. This can be dealt with by daily checks carried out by the greenhouse personnel. In this way, the method provides a maximum guarantee of efficiency. Electronic leaf This is a small piece of plastic or glass, horizontally positioned, of about the same dimensions as a leaf. The film of liquid which the mist leaves on it allows for the passage of electric current between two electrodes positioned at each end. When evaporation breaks the circuit, the system resumes watering. This device is not widely used because of the difficulties of setting it to mimic the evaporation rate from the olive leaves. Scales With this method, evaporation from a predefined quantity of water alters the trim of the scale which, in turn, starts up the mist. The calibration of the scales in accordance with evaporation from leaves is not easy. As for the electronic leaf, this method is not widely used with olives. Photoelectric cells This system functions on the basis of the intensity of light inside the greenhouse. Light intensity is measured by a photoelectric cell which controls a pulse counter on the control panel. Pulses follow one another at a speed that is linked to the quantity of light. The system modifies pulse speed, which regulates the intervals between mistings. At the moment of maximum light (which corresponds to maximum evaporation) the number of pulses necessary to maintain the right amount of moisture on the leaves is established (the greater the light the more intense the evaporation) by the operator. The duration of misting is controlled by a timer. Though the method is not very accurate, it is highly dependable. Integrated systems Here, a computer takes into account the variations of environmental conditions (light, temperature, relative humidity) and maintains what has been decided upon at the beginning and/or makes modifications over time. This is an extremely precise method of control which can be adjusted to the specific requirements of any type of cutting. It is affordable and competitively priced. However, repairs are difficult and must be carried out by very specialised personnel. Furthermore, availability of spare parts on the spot can be a problem. It is to be remembered that if the greenhouses are situated at some distance from a town from which ‘technical assistance’ for instruments has to come, a delay of two hours before necessary work is done on a misting system could compromise all the cuttings placed for rooting. Consequently, it is advisable for ‘commercial’ greenhouses to have two systems of control, or at least to use equipment that personnel on the spot can repair easily and quickly.

The best method of water distribution for the olive would be the use of both mist and fog. However, such a system would require considerable investment as well as qualified operators and so is suitable only for medium to large nurseries or at any rate greenhouses and/or nurseries which ‘produce’ without interruption (full use of

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greenhouses for other species also). An interesting alternative for the small business or olive farm can be the ‘closed frame technique’ (see box).

6.4.3 Light The role played by light on olive rhizogenesis (quantity and wavelength) has not been studied. However, it is advisable to work with a slight reduction in the amount of light in order to limit water evaporation from the leaf surface and to reduce temperature; the rooting area should be shaded by about 30% with nets or tinted glass. CLOSED FRAME TECHNIQUE The ‘closed frame technique’ consists of a heated bench, which is placed inside a metal and plastic box and covered with an airtight lid. The bench, 30–40 cm high, is raised to about 1 m and contains a rooting substrate or small plastic containers (Fig. 6.18) in which cuttings are placed. The substrate, usually perlite, must be kept very moist, although not pressed too much, to ensure the maintenance of 100% relative humidity without waterlogging. An electric resistance is inserted in the lower part of the bench, in order to maintain the rooting zone at 20–22°C for the entire rooting cycle. The frame is placed in a greenhouse, covered with a shading net. No extra watering is required in the rooting period (50–60 days), during which time the lid should never be opened.

Figure 6.18 Rooting bed for the ’closed frame technique’.

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6.5 Hardening (1st transplant) Rooted cuttings should be transplanted at the right time to avoid root decay. Since the intrinsic and extrinsic factors which influence the speed of root development are many, rooting time may vary quite widely, and therefore it is useful to have general criteria to refer to when deciding on the right moment to transplant cuttings. The cuttings must have roots of an average length of 3 to 5 cm, greybrown in colour and no longer fragile, so as not be easily damaged (Fig. 6.19). If these conditions are met, the cuttings can be taken up and transplanted (Fig. 6.20). If necessary, cuttings can be left longer under mist, but not for more than an additional month, and care must be taken to reduce the amount of water given (longer intervals). Rooted cuttings must be transplanted in a short time (not more than two hours from removal from the misting greenhouse) in order to keep the roots from drying out. Every Figure 6.19 precaution must be taken to avoid water stress. Immediately after transplanting, the plants must be watered and positioned where it is

Figure 6.20 Rootlings at transplant time.

Rooted cutting sufficiently developed for transplant. Notice the burst buds and ramifications in the roots; root darkening indicates their maturation towards full functionality.

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possible to provide brief but frequent irrigation. In practical terms, the rooted cutting must be moved from a very humid environment (mist) to one where humidity is gradually diminished. However, in the first few days after transplanting the leaves should also be kept moist; in any case, the substrate must never be waterlogged. The choice between one or two transplants before final in-field planting depends on whether or not a slow growth rate (one transplant) or a faster one (two transplants), and therefore different plant heights, is desired.

Figure 6.21 Potted rootlings in trays, in the hardening greenhouse.

6.5.1 Containers and substrates The volume of the containers should be about 500 mL and not less than 300 mL. A final pot, not less than 2000 mL, can be used from the start, but this procedure is not optimal because the growth rate of the canopy is slower: the plant tends to develop its root system first, in relation to the space available, before pushing the growth of the crown; the ensuing reduced quantity of foliage means less photosynthesis, and therefore the plant does not recover the missing vegetation by the end of the vegetative period. As a result, the plantlets develop faster and are better balanced if they undergo several transplants. If possible, containers should be square in shape so as to occupy less surface area. Trays should be used to make removal of containers to different locations within the greenhouse easier (Fig. 6.21). As regards the substrate, the box â&#x20AC;&#x2DC;Examples of substrates for the 1st and 2nd transplantâ&#x20AC;&#x2122; reports just one example of mixtures that can be used after rooting has taken place. However, the number of natural materials, and industrial and agricultural residues

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used in compost around the world is very large; hence, for a comprehensive description, specific texts or reports should be consulted (e.g. Bunt 1976; Bartolini & Petruccelli 1991a, 1991b). EXAMPLES OF SUBSTRATES FOR THE 1ST AND 2ND TRANSPLANT Substrate for the 1st transplant • peat moss 40% (use peat with the highest – i.e. the least acid – pH) • pumice (3–5 mm) 40% • soil compost 10% (over 6 month old) • sand 10% • lime (CaCO3, about 100g/m3 to raise the pH to about 6; Ca(OH)2, etc.) The materials must be thoroughly mixed. The addition of mineral fertilisers at this stage is not advised, because of the short duration of this stage (1–4 months), and because the root systems are still young and delicate. Substrate for the 2nd transplant • peat moss 30–50% (use peat with the highest pH) • pumice (3–5 mm) 10–40% • soil compost 10–40% (over 6 month old) • lime as above, to adjust pH • mineral fertilisers 3–4 kg/m3 It is advisable to use slow release fertilisers (such as 2 kg soluble over 3–4 months and 2 kg over 8–9 months). The use of these products, mixed within the substrate, reduces the need to repeatedly supply solid or liquid fertilisers. They are immediately soluble and used during plant growth. In this way, labour requirements are reduced, if compared to fertigation, and the residual salt concentration in the pot is never too high. The negative aspect is that the best mineral balance is never achieved this way, and because, over time, the plant has different nutritional requirements which can be better satisfied with fertigation.

The size of the pot and the amount of substrate it contains limit the length of time the plant can be kept in the container to about 1 to 2 months. After this the plant will suffer stress because of lack of space which will lead to the precocious ageing of the root system. The right moment to carry out the second transplant should be decided according to the development of the root system. The roots must protrude from the substrate, coming into contact with the container inner surface by a few millimetres or at most a few centimetres. Transplanting when the roots are excessively long and developed outside around the substrate should be avoided.

6.5.2 Transplanting environment Transplanting should not be carried out on particularly hot or windy days. Cuttings that have just been potted should be placed in one of the following environments: • a misting greenhouse, obviously only for small numbers or for particularly valuable plants • hardening greenhouses, tunnels or lath-houses, where it is possible to control watering automatically (frequency and quantity). If the transplant is done during

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cold periods of the year, a heating system will be necessary if and when the outside temperature goes below 5Â°C. If the greenhouse has a heated floor, this maintains an optimal temperature for the base of the pot and therefore for the root system which is stimulated towards more rapid growth, and ensures a sufficient temperature throughout the greenhouse. These must have a watering system and be sufficiently shaded (maximum 30% reduction of light) in order to limit evaporation and to avoid direct sunlight in the first few weeks after transplanting. Where the first transplant takes place, plant health must be carefully and constantly checked as a warm damp environment can be conducive to fungal diseases.

6.6 Plant growing (2nd transplant) The right time to carry out the second transplant is generally 1 to 2 months after the first. This is done to increase growth and to obtain the best possible balance between the size of the root system and the upper part of the tree. Generally, plants remain in these pots for 8 to 12 months before being planted out (Fig. 6.22). A longer period in a pot is not advisable even if the plant is given sufficient fertiliser because it results in a plant disproportionately large above ground in relation to the root system. In such cases the plant is slow to take root when it is planted out and slower to resume growth than younger, better balanced plants. The plant first needs to redress the imbalance between the root system and the part above ground before the latter resumes its vegetative growth.

Figure 6.22 Plants in the open, about 12 months after the 2nd transplant.

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6.6.1 Containers and location of plants The optimal volume of the container is between 2 and 3.5 litres depending on the length of time envisaged and the amount of growth desired before planting out. It should be square to save space in the nursery as well as during transport. A suitable area for plant growing should be located as close as possible to those areas previously described (mist, hardening) in order to reduce the time required to move the pots. It should be slightly sloping so that rain and irrigation water can run off. To control weeds, the ground should be covered with plastic for mulching, or if available, black permeable plastic. Occasionally the plants will need to be shaded (maximum 30%). The area must be accessible to vehicles for the transportation of plants and for all other operations. Plants are shipped in the 2 L pots, or in larger ones if kept in the nursery to grow for more than 2 years. In the past, when light plastic containers were not available, the plants were extracted from the soil in autumn with the earthball, which was wrapped in rye straw before shipping (Fig. 6.23).

Figure 6.23 Three-year-old plants (the Italian â&#x20AC;&#x2DC;Piantoniâ&#x20AC;&#x2122;) ready for shipping. Note the earthball, wrapped in rye straw and containing the plant roots.

6.6.2 Irrigation and fertilisation The area where the plants are grown must have an automatic irrigation system. The amount and frequency of water given depends on the climate where the nursery is located and on the system adopted. It is generally advisable to avoid excessive watering

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and the occurrence of waterlogging for any length of time in the pots; as a general guideline: 500 to 1000 mL per pot, 2 to 3 times a week in hot periods, should be supplied.

ADDITIONAL CONSIDERATIONS ON THE TIMING OF PROPAGATION CYCLES The choice of which and how many production cycles to undertake will depend on the market and the growth rate of the stock plants, as well as on the growth speed of the already propagated plantlets. It should be remembered that trading periods of olive plants coincide with planting out, an operation that as a rule occurs during a few relatively cold months and with the prospect of a rainy period to enable the plants to face the hot season which follows. If these apparently trivial facts are not remembered, a fixed irrigation system will become necessary, or at least an emergency one. If the cuttings root in spring, the plants would not be big enough (over 1 m) to plant out in the same year of production or within a year from the collection of cuttings. Thus the plant remains in the nursery until it is 18 to 24 months old before being put on the market (over 120 cm). The same plant can be grown in about 6 months less time by propagating at the beginning of autumn and selling the plant after 12 to 18 months (over 120 cm), ready for planting out. The choice of propagation by cutting in autumn gives the nursery greater organisational freedom for the propagation by grafting done in spring, as the two methods are thus separated and do not overlap. This choice of action is almost inevitable for small nurseries, while in medium to large nurseries organisational resources make it possible to bear the burden of work entailed by doing both activities together. In warm temperate climates (Southern Italy, Andalusia, North Africa, etc.) best rooting takes place at bud sprouting (Februaryâ&#x20AC;&#x201C;April). In these conditions the growing season facing the rooted cuttings is longer, and the resulting plants achieve the proper size for transplant (over 120 cm) within the same year. It should be remembered that it is preferable to plant out when plants are relatively small (1 to 1.5 m) and have been in the container for less than a year. This keeps production costs down (less time in the nursery), ensures a higher survival rate and, above all, a better recovery (growth of the plant after transplanting). This is greater and more immediate with small plants as compared with bigger ones that have spent more time in a pot, because the former have a better balance between the parts below and above ground. It has been noted that when a plant remains in the same pot, the roots do not grow as much in volume as the part above ground, and age and die. On the other hand, it is not advisable to transplant olives that are less than a year old (50 to 60 cm in height) because of the considerable amount of care they need (drip irrigation, protection from weeds and animals, etc.). The period in which scions are taken obviously coincides with the two peak times for rooting (see also Fig. 3.11). The first, in early spring (Marchâ&#x20AC;&#x201C;April, in the Mediterranean area), is concomitant with the onset of vegetative growth and precedes grafting by a few weeks; the second, in summer-early autumn (Augustâ&#x20AC;&#x201C;October), is when the plant begins to grow again after the summer pause. By taking material for rooting in the latter period it is possible to produce plants in about a year, while if it is taken in the spring period production time is lengthened, as there is only one period recommended for planting which is autumn-winter (November-March in the Mediterranean area), depending on when rain and cold spells are to be expected. When material is taken during the autumn period it is advisable to avoid drastic pruning of stock plants because this can lead to a late, second bursting of buds. This, in turn, causes extreme sensitivity to cold which could lead to plant death. For this reason, it is advisable not to prune trees after late summer (September, in the Mediterranean area) and no more than necessary. Also, severe pruning in the spring period can unbalance the development of stock plants and as a consequence limit the bursting of new buds and temporarily arrest vegetative growth.

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Long practical experience has led to the conclusion that, in order to maintain the growth of a potted plant for about a year, a complete (NPK) slow-release mineral fertiliser should be put into the substrate (for example 4 kg/m3 of OsmocoteÂŽ, with 2 kg after 3 to 4 months and another 2 kg after 8 to 9 months). The use of mineral fertilisers in the pots brings with it the risk that on dissolving rapidly they increase salinity inside the pot. This also happens when evaporation is high and particularly if hard water is being used (in hot periods or climates). When this happens, in order to diminish salinity, the pots should be abundantly irrigated periodically to â&#x20AC;&#x2DC;washâ&#x20AC;&#x2122; the substrate. The problem of the right supply (proportions and quantity) of the various nutrients can be solved by using fertigation. This technique makes automatic modification of dosages possible in accordance with the growth rate of the plant, so that fertiliser is given only when needed. The disadvantage of this technique is that it requires a drip irrigation system adapted to each single pot. The cost is usually too high to make it commercially viable.

6.7 Plant training The pots are generally arranged in double rows. Rows of three or four are not advisable except in particular environmental conditions where there is plenty of light and a long growing season (more than six months). Each pot must be supplied with a tutor, i.e. a cane about 1 m in length to support the plant and force it to grow vertically. Various materials are used to tie the plant to the tutor, such as plastic strings or tapes attached manually. Excision of unwanted adventitious shoots is usually not necessary for some months. This can become necessary from the fourth or fifth month, when growth below 50 cm is eliminated. Lateral buds growing higher up need not be pruned off unless they are so vigorous that they compete with the apical bud. If necessary, a few weeks after potting, when the plants have got over the period of stress, a thin layer of pine bark can be spread over the surface of the soil in the pot to limit or prevent the development of weeds. When the plants have reached a height of 80 to 90 cm from the collar (after approximately 6 months in the pot, depending on local conditions) it can be given their final shape in accordance with what is required from the nursery. If they are to be grown as open centre trees in the field, it is useful to cut back the plants at 80 to 100 cm to facilitate the growth of lateral shoots from which to select the main limbs. Otherwise the shape will be a central leader or monocone, and the plant will be left to grow without interference. The choice of the final training system is not the concern of the nursery but of the olive grower. However, information of a general nature on the issue is useful for the nursery. For manual or facilitated harvesting, the open centre, or vase, and similar systems are the most suitable. Usually these are shapes that have greater surface exposure to the light and consequently greater fruiting area and bigger yield. Machine harvesting with shakers is possible with the open centre training system and its efficiency is similar to that of the monocone. It is advisable to use cultivars with a compact habit and an early and short ripening period. Planting spacings, depending on environmental conditions, vary from 6 to 10 m between rows and 4 to 6 m between trees along the rows (e.g. 6 x 4, 6 x 5, 6 x 6,

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7 x 5, 7 x 6). The maximum height of the tree, as maintained by pruning, should not be greater than 4 to 5 m. For machine harvesting, there has been a preference in recent years for the monocone, with planting distances shorter than for the open centre, in the order of 6 m between rows and 3 to 4 m along the row. With due regard to local conditions, planting trees too close is not advisable as it results in a need for drastic pruning when the canopies come into contact; it may also be preferable to uproot the trees and replant them with greater spacings. These operations are always very expensive and the possibility of having to resort to them must be borne in mind when the distances between trees are being decided upon. Most of the ever accumulating experience available shows that the results from this training system have not been up to expectations.

6.8 Plant certification Olive domestication and cultivation determined the selection of plants adapted to the most varied environments in terms of soil conditions, climate, and pest and disease resistance, while producing satisfactorily in terms of quantity and quality. Such plants (genotypes) often show minor differences among themselves, and are not easily distinguished by non-specialists. The cultivars of the olive derived from natural and artificial selection are of two main types: â&#x20AC;˘ monoclonal, represented by a single clone. There are numerous examples, usually local cultivars traditionally confined to a small cultivation area. â&#x20AC;˘ policlonal, comprehending a variable number of clones. Differences among clones are for very few characters, usually of no commercial importance. Only expert technicians working in the region can tell the clones apart, otherwise biochemical and/or molecular techniques must be used. Besides the genetic differences among cultivars, a marked influence on character expression is exerted by the environment; the result is that, by varying the cultivation site, different cultivars may appear identical, or the same cultivar may display a different phenotype in differently located orchards. The confusion that may ensue from this is reflected in the naming of cultivars (errors in identification, synonimies, omonymies, etc.), and the resulting lack of certainty in the name of the plants of interest. This problem is increased by the growing expansion of olive cultivation outside the Mediterranean, in areas where growers are less tied to traditions and look for the best plants for their orchards; on the other hand, many new growers often uncritically accept what is available, and rely on secondhand information that is sometimes far from accurate. All the above therefore justify the urgent need to guarantee the genetic origin and the good sanitary condition of the traded olive plants, allowing the provision of plants accompanied by reliable commercial certifications.

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6.8.1 Genetic origin The indication of genetic origin guarantees that the material used for propagation was taken from a stock plant that has been described in detail, both from a morphological and agronomical point of view, that it certainly belongs to a given cultivar, and that it is conserved in experimental fields or in germplasm collections. A model for certification can be represented by what is proposed in Italy for voluntary certification. It begins with the transfer of propagating material (plants) from the breeder (or from whoever carried out a program of clonal selection) to an institution that is in charge of assessing the sanitary conditions of the plants and whether their morphological and agronomical characters are as formerly declared by the breeder. Plants that satisfy the above requirements are transferred to the multiplication centre, which has the task of growing the accessions as stock plants, which in turn will provide multiplied material for nurseries. The process can be split into more steps, each with a different function, as in the box. STEP

ACTIONS

Breeder

Primary source

Verification centre for premultiplication (pre-base material; usually a public research centre)

Evaluation of accessions subject to certification, from sanitary and genetic points of view

Premultiplication centre (usually a public research centre)

Conservation of accessions and distribution of propagation material to multiplication centres

Multiplication centre (can be a voluntary association of nurseries, or depending on local governments; in the first instance a public control can be envisaged)

Planting and growing of stock plants for the production of propagation material for individual nurseries

Commercial nursery

Production of certified olive plants

Certification must be witnessed on each plant by an attached tag, carrying all information needed (cultivar, rootstock and clone if applicable, name and location of nursery, and whatever the law requires). Certification programs can include both patented and non-patented cultivars. New plant selections undergo different regulations in the different European Union countries. The main guidelines used in this regard make reference to the Paris Convention of December 1961 (Act 722/74 in Italy), and to the UPOV Convention of 1991. As a rule, whatever the law adopted for certification, the main problem remains to guarantee the genetic identity of the plant, that is, strictly speaking, its name. The breeder and the institution conserving the germplasm are responsible for the genetic identity.

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Each of these has specific responsibilities: Breeder (public researcher, nurseryman, olive grower, etc.) • Origin: • Plants from conventional breeding • Plants from selection within known cultivars or clones • Transgenic plants • Identification of plants (this information must be published in specialised journals): • Morphological traits • Agronomical characteristics (by using internationally accepted pomological descriptors) • Biochemical and/or molecular traits Public research institution • Possession of a known cultivar collection, information on which accessions are available from pomological literature, internet, etc.; such collections must have been established in a due period of time by specialists. • Accessions should have been collected by expert persons, living in the area the accession comes from. • Morphological and agronomical description on internationally accepted descriptors; this may not be completely coincident with the breeder’s description. • The true-to-typeness of accessions should be periodically checked by means of biochemical and/or molecular analyses, published in specialised journals.

6.8.2 Good sanitary conditions The plants must be exempt from diseases whose symptomatology is visible and nonvisible (as from draft international protocols, European Plant Protection Organization, Ministries of Agriculture, Regional Authorities, Institutes of Plant Pathology, etc.). The European Union has promulgated directives concerning the commerce of propagation materials, indicating the sanitary prerequisites the plants should have (Council Directive n. 92/34): absence of quarantine pathogens (listed in the EEC Directive 77/93) and of other agents of diseases which can alter the quality of propagation material (material of European Agricultural Conformity). The European Union has not yet approved a protocol for the production of certified material. When the required sanitary conditions are fulfilled, the plant, kept in a sterilised substrate, can be transferred from one country to another without any particular risk. This international movement of germplasm is presently quite difficult, due to a marked heterogeneity in regulations and laws (such as different quarantine periods). Certification does not involve any guarantee of the quality of the product (oil or fruit), which can only be provided by the breeder.

References Bartolini, G. & Petruccelli, R, 1991a. I substrati nel vivaismo, Ia parte. Colture protette, 6: 47–64. Bartolini, G. & Petruccelli, R, 1991b. I substrati nel vivaismo, IIa parte. Colture protette, 7: 33–54. Bunt, A.C. 1976. Modern Potting Composts. The Pennsylvania St. Univ. Press, USA, pp. 278. Fontanazza, G. & Rugini, E. 1977. Effect of leaves and buds removal on rooting ability of olive tree cuttings. Olea (December): 8–28. Nelson, P.W. 1985. Greenhouse Operation and Management. Reston Publ. (Prentice-Hall Co.), Reston (USA), pp. 598.

7 Conservation of olive germplasm

Olive germplasm is usually preserved in clonal collections, located in different sites in several countries. There is nothing like a centralised â&#x20AC;&#x2DC;serviceâ&#x20AC;&#x2122; for conservation; however, several institutions (universities, research centres, local authorities, nursery associations, private firms, etc.) carry on, on their own, the recovery, propagation and maintenance of a number of accessions, mainly the autochtonous ones. Only recently large public Institutions at an international level, such as the Ministry of Agriculture and Forestry Politics (MiPAF) of Italy and the International Olive Oil Council (IOOC), have promoted a thorough conservation campaign, aimed at retrieving and also preserving accessions from distant locations and countries where conservation is not provided. The kernel of germplasm conservation still lies in stock plant collections that nursery operators, alone or associated, keep for their propagation needs. This means that such collections include a limited number of local cultivars, and a variable number of internationally renowned cultivars. This is the situation of international olive genotype conservation. It is a system that depends on the location of the most important nurseries, which are concentrated in particular areas that do not always coincide with those where production is concentrated and where olive biodiversity is highest. The number of olive cultivars propagated in each country is relatively small and, on the world scale, only about a hundred cultivars are propagated by nurseries in detectable amounts. Due to poor communication between research institutions and public authorities, on one side, and nurseries on the other, valuable local germplasm is often neglected. The need to have large clonal collections, ideally replicated in several different environments to reduce the risk of loss of valuable germplasm due to biotic and abiotic damage, involves considerable financial support. For instance, a theoretical collection with 1200 accessions, each represented by four trees, having a density of 333.3 trees/ha (6 x 5 m), needs as much as 14.4 hectares. Such a size can be considered sustainable if the

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olive grove is productive (oil or table olives), but its profitability is reduced by the complexity of the operations determined by the differences among genotypes; if the surface is reduced by reducing spacing, productivity is reduced, and possibly confined to the production of scionwood. In both cases, the income can never equal maintenance costs. Clonal collections of medium-to-large dimensions are therefore only justified as a support to research activities, both on a national and international level. Their funding must therefore be based on public support, possibly in conjunction with international organisations. In the near future, alternative strategies of germplasm preservation (see 5.5) could join the traditional in vivo approach, and pave the way to a larger and safer maintenance of olive biodiversity.

Appendix 1

Rooting ability of all olive cultivars of which scientific literature is available. The reported percentages are the average of results obtained by different authors, after IBA treatments (extracted from Bartolini et al. 1998).

High rooting ability: 66 to 100% of rooted cuttings Aglandau, Alfafara, Arancino, Arbequina, Ascolana Tenera, Ashrasy, Ayvalik, Bari, Barnea, Bashika, Blanquette De Guelma, Borgiona, Bouchouk, Bouteillan, Canetera, Carolea, Cellina Di Nardò, Chatawi, Ciliegino, Coratina, Corbella, Cordovil De Castelo Branco, Corniolo, Coroncina, Correggiolo, Correggiolo Di Pallesse, Correggiolo Di Villa Verucchio, Cuoricino, Curivell, Da 12 I, Degal, Dulzal, Duzica, Frangivento, Frantoio, Frantoio Di Montegridolfo, FS–17, Fulla De Salze, Gemlik, Gioufong N. 1, Gordal De Granada, Görvele, Haouzia, Itrana, Jaropo, Kalinjot, Kan Çelebi, Kilis Yaglik, Kokermadh I Elbasan, Kothreiki, Lastovka, Leccino, Lechin De Sevilla, Lucio, Lucques, Madonna Dell’impruneta, Maiatica, Mansino, Manzanilla Cacereña, Manzanilla De Jaën, Manzanilla De Sevilla, Manzanilla Prieta, Menara, Mignola, Mignolo Cerretano, Mission, Mixan, Mollar De Cieza, Moraiolo, Morellona Di Grecia, Morisca, Morona, Morrut, Nebbia, Negrillo De Arjona, Nichitskaia Ii, Nocellara Del Belice, Nocellara Messinese, Nostrana Di Brisighella, Oblica, Ogliarola Barese, Olivastra Di Suvereto, Olivo Del Palone, Olivo Di Casavecchia, Pendolino, Pesciatino, Picudo, Pignola, Pisciottana, Raio, Ravece, Razzaio, Romanella, Rosciola, Rosciola Colli Esini, Rossellino, Rossello, Rotondella, Roughani, Ruveia, Salella, Samanli, Sari Yaprak, Sevillenca, Swan Hill, Toccolana, Tondina, Valle Fiorana Clone N. 50, Verdal, Verdial De Alcaudete, Verdial De Badajoz, Verdiell, Zard.

Medium rooting ability: 33 to 66% of rooted cuttings Adramitini, Aggezi shami, Allorino, Americano, Amigdalolia, Arbosana, Arnasca, Azapa, Azeitoneira, Bardhe I Kruje, Bardhe I Tirane, Becarut, Bella Di Cerignola, Bical De Castelo Branco, Bidh El Hammam, Blanqueta, Cailletier, Çakir, Callosina, Canino, Canino Di Bagno, Cañivano Blanco, Capolga, Carbunción Di Carpineta, Carrasqueño, Carrasqueño De La Sierra, Casaliva, Castellana, Cayon, Çekiste, Chalkidiki, Chemchali, Chétoui, Chorrúo, Cobrançosa, Colombina, Colombino, Cordovil De Serpa, Cornetta, Cornezuelo, Cornicabra, Correggiolo Di Montegridolfo, Cucca, Da Cuccare, Dezful, Doebli, Dolce Agogia, Dolce Di Andria, Edincik Su, Egriburun, Erkence, Esek, Fadhel, Fecciaro, Frantoio Andrea Corsini, Frantoio Di Villa Verucchio, Frengu, Galega Grada De Serpa, Galega Vulgar, Gentile Di Chieti, Giogolino, Girit, Gremignolo Di Bolgheri, Grossa Di Cassano, Grossaio, Grossanne, Hamed, Hojiblanca, Imperial, Izmir Sofralik, Jabaluna, Kadesh, Kalamata, Kalembezi, Karamürsel Su, Karidolia, Khodeiry, Kiraz, Kokermadh I Berat,

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Konservolia, Koroneiki, Labeeb, Ladoelia, Larcianese, Lea, Leccione, Levantinka, Limli, Maçanilha Carrasquenha, Maelia 29, Maremmano, Marsaline, Mary, Mastoidis, Maurino, Megaritiki, Melaiolo, Memecik, Memeli, Menya, Merhavia, Meski, Meslala, Mignolo, Moraiolo Tommaso Corsini, Negrinha, Nera Di Gonnos, Nerba, Nevadillo Negro, Nizip Yaglik, Nostrale Di Rigali, Oblonga, Ogliarola, Olivago, Olivo Bufalo, Olivo Delle Alpi, Olivoce, Orbetana, Ornellaia, Ottobratica, Ottobrina, Pajarero, Palma, Palomar, Piantone Di Falerone, Piantone Di Mogliano, Picholine, Picholine Marocaine, Picual, Piturzello, Pizze Carroga, Pocciolo, Raia, Rapasayo, Razzo, Razzola, Redding-Picholine, Redonale, Redondil, Rojal, Rosino, Rossellino Cerretano, Saiali, Salicino, Sam, San Benedetto, Santa Caterina, Sant’agostino, Sari Hasebi, Sari Ulak, Sorani, Sigoise, Taggiasca, Tanche, Tavsan Yürégi, Teffahi, Tonda Iblea, Úlli I Kuq, Vera, Verdale De L’hérault, Verdale Des Bouches-Du-Rhône, Yag Çelebi, Yuvarlak Halhali, Zarza.

Low rooting ability: 0 to 33% of rooted cuttings Alameño De Montilla, Albatro, Argudell, Ascolana Dura, Azéradj, Bella Di Spagna, Bianchera, Biancolilla, Boçi, Branquita, Briscola, Büyük Topak Ulak, Cañivano Negro, Carmelitana, Carrasquenha, Cayet Roux, Çelebi, Changlot Real, Chemlal, Chemlali De Sfax, Çilli, Conserva De Elvas, Correggiolo Di Massa Martana, Dekkar, Djilt, Domat, Dritta, Emilia, Empeltre, Farga, Feglina, Filare, Firenzuolo, Gentile Di Larino, Gerboua, Ghiacciolo, Ghiandaro, Giarraffa, Ginestrino, Golbina, Gordal Sevillana, Grappolo, Grappuda, Gremigna Tonda, Gremigno Di Fauglia, Gremigno Di Montecatini, Gremignolo, Grossa Di Gerace, Grossolana, Halhali, Hollë I Himarë, Hurma, Intosso, Kaissy, Kallmet, Karamani-Kachabi, Lastrino, Lazzero, Lazzero Delle Guadalupe, Lazzero Di Prata, Leccio Del Corno, Leccio Maremmano, Lianolia Kerkyras, Maçanilha Algarvia, Madremignola, Madural, Managjel, Manzanilla De Montefrio, Marks, Marzio, Morcaio, Morchione, Morcone, Moresca, Mortellino, Mummiana, Nabali Baladi, Nocellara Etnea, Ocal, Ogliarola Messinese, Olivastra Di Populonia, Olivastra Seggianese, Olivo Del Mulino, Olivo Di San Lorenzo, Oueslati, Passulunara, Pendagliolo, Piangente, Punteruolo, Puntino, Quercetano, Racimal, Rasi’i, Real Sevillana, Redondal, Rossina, Salonenque, Sammartinara, San Francesco, San Lazzero, Sargano Di Fermo, Scarlinese, Selvatica Tardiva, Selvatico, Shami, Shatawi, Soury, Spagnola, Tondello, Tortiglione, Ulli I Zi, Unafka, Uslu, Valanolia, Varagen, Verdeal Alentejana, Verdeal Trasmontana, Verdial De Huévar, Verdial De Vélez-Malaga, Villalonga, Voce, Zaituna, Zalmati, Zarrazi.

Appendix 2

Olive germplasm collections throughout the world (Bartolini et al. 1998) Albania Instituti i Kerkimeve te Ullirit dhe Agrumeve (Vlorë)

Algeria Station Expérimentale de Sidi Aigh Station Oleicole (Cap-Djinet)

Argentina Colleccion de Variedades de Hintermeyer (La Rioja) Colleccion Vivero Nacional (La Rioja) Estacion Experimental Agropecuaria – INTA (Catamarca) Estacion Experimental de Quines (San Luis) Ministerio de Agricoltura de San Juan Substacion Experimental Junin (Mendoza)

Australia Horticultural Research Station (Mildura) Victoria National Olive Variety Assessment, University of Adelaide – Roseworthy Campus, (Roseworthy), South Australia Plant Biology, Faculty of Natural and Agricultural Sciences, University of Western Australia (Crawley) Western Australia Riverina College of Advance Education and Yanco Agricultural Institute (Wagga Wagga) New South Wales Charles Sturt University (Wagga Wagga) New South Wales

Azerbaijan Institute of Horticulture and of Subtropical Plants (Baku)

Brazil Fazenda Sain André (Castro)

China Kunming Botanical Research Institute of the Cafsr (Yunnan) Olive Garden (Shaanxi, Chegdu County)

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Cyprus Zyghi station

Egypt Faculty of Agricolture, Cairo University (Giza)

France Assoc. amis de l’olivier. Confrérie chevaliers de l’olivier de Vans. Syndacat oléicult. Ardèche Mérid. INRA, UR-Génétique et Amélioration des Plantes (Montpellier) S. E. I. (Ghisonaccia, Corse)

Greece Experimental Station of Agios Mamas Khalkidiki (Chalkidiki) Substropical Plants and Olive trees Institute of Chania (Crete)

India Department of Pomology, Dr. Y. S. Parmar Univ. of Horticulture and Forestry, Solan (H. P.) Directorate of Horticulture, Bajaura (Kulu), (H. P.) Directorate of Horticulture, Dakrani (Dhera Dun), (U. P.) Directorate of Horticulture, Jeolikote (Nainital), (U. P.) Directorate of Horticulture, Maitra (Doda), (J. & K.) Directorate of Horticulture, Panarsa (Mandi), (U. P.)

Iran Agriculture Research Center (Ahvaz Khozestan Province) Agriculture Research Center, Dezful Research Station-Safiabad (Khozestan Province, Dezful) Agriculture Research Center (Gorgan Province) Rodbar Olive Station (Gillan Province)

Israel Agricultural Research Organization The Volcani Center, Institute of Horticulture (Bet Dagan)

Italy Azienda Agricola Sperim. entale Dimostrativa ‘Incoronata’, A. L. S. I. A., Melfi (Potenza) Campo Carboj; Menfi, E. S. A., Castelvetrano (Trapani) Centro Agricolo Sperimentale ‘Monna Giovannella’, Antella (Firenze) Consozio Interprovinciale per la Frutticoltura, Villasor (Cagliari) Dipartimento di Arboricoltura e Protezione delle Piante, University of Perugia Dipartimento di Coltivazione e Difesa delle Specie Legnose, Sez. Coltivazioni Arboree University of Pisa Dipartimento di Colture Arboree, University of Palermo

A ppendices

135

Dipartimento di OrtoFloroArboricoltura e Tecnologie Agroalimentari, Sez. Arboricoltura, University Catania Dipartimento di Ortoflorofrutticoltura, University of Firenze Dipartimento di Produzione Vegetale – sez. Coltivazioni Arboree, University of Milano Dipartimento Scienze Produzioni Vegetali, University of Bari, Centro Didattico-Sperim. ‘P. Martucci’ (Bari) Istituto di Ecofisiologia delle Piante Arboree da Frutto, CNR (Bologna) Istituto di Ricerche sulla Olivicoltura, CNR (Perugia) Istituto per la Fisiologia della Maturaz. e Conserv. del Frutto delle Sp. Arboree Mediterranee, CNR (Sassari) Istituto per la Valorizzazione del Legno e delle Specie Arboree, Azienda Sperimentale S. ta Paolina, Follonica (Grosseto) Istituto Sperimentale di Frutticoltura (Verona) Istituto Sperimentale per l’Olivicoltura (Rende) Istituto Sperimentale per l’Olivicoltura (Spoleto) Istituto Sperimentale per la Elaiotecnica (Città S. Angelo) Istituto Tecnico Agrario Statale (Pescia) Istituto Tecnico Agrario Statale (Cividale del Friuli)

Japan Shozu Branch (Schozusun Ikedacho Ikeda 237) Tokusan no Kudamono Olive (Ideta, Takagi)

Jordan Khalidieh Agricultural Stations (Khalidieh)

Morocco Centre Régional de la Recherche Agronomique du Haouz Presahara, INRA (Marrakech) Station Centrale de Recherche sur l’Olivier, INRA (Rabat) Station Experimentale d’AHL Souss Station Experimentale d’el Maghrek (Doukkala) Station Experimentale de Ain Taoujdat, INRA (Mekhès) Station Experimentale de la Menara, INRA (Marrakech)

Nepal Bissingkhel and Chitlang (Makwanpur) Kirtipur (Katmandu) Marpha (Marpha) Thaiba (Lalitpur)

Portugal Direcção Regional de Agricultura do Ribatejo e Oeste (Vila Franca de Xira) ENFVN, Department Olivicultura (Elvas) Estacao Agronomica Nacional (Oeiras)

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Republic of South Africa Arc-Fruit, Vine and Wine Research Institute (Stellenbosch)

Serbia and Montenegro Station de Cultures Subtropicales (Bar)

Spain Coleccion de Batea. IRTA, Dept. Arboricoltura Mediterrània (Reus, Tarragona) Coleccion de Mas Bové. IRTA, Dept. Arboricoltura Mediterrània (Reus, Tarragona) Coleccion de Olesa de Montserrat. IRTA, Dept. Arboricoltura Mediterrània (Reus, Tarragona) Coleccion de Tortosa. IRTA, Dept. de Arboricoltura Mediterrània (Reus, Tarragona) Olive World Collection; C. I. D. A. Alameda del Obispo (Córdoba)

Tunisia Collection Bir Chbika. Cooperative centrale des semences et plants selectionnés (Tunis) Collection Borj El Amri 1. Institut de l’Olivier. IRESA, Ministère de l’Agriculture (Tunis) Collection Borj El Amri 2. Institut de l’Olivier. IRESA, Ministère de l’Agriculture (Tunis) Collection Chott el Ferik. Institut des Régions Arides (Medenine) Collection Chott Meriem. Ecole Superieure d’Horticulture (Chott Meriem, Sousse) Collection de Boughrara. Institut de l’Olivier (Sfax) Collection INAT 1. Institut National Agronomique de Tunisie (Tunis) Collection INAT 2. Institut National Agronomique de Tunisie (Tunis) Collection Mehrine. Institut de l’Olivier. IRESA. Ministère de l’Agriculture (Tunis) Collection Ksar Gheriss (Maknassy-Mezzouna). Inst. Nat. Rec. Genie Rural, eaux et Forets (Tunis) Institut National des Sciences Appliquées et de Tecnologie (Tunis)

Turkey Institut de Recerche Oléicole (Kemalpasa)

United States of America Department of Pomology, University of California (Davis)

Reference Bartolini, G., Prevost, G., Messeri, C. & Carignani, G. 1998. Olive germplasm: cultivars and world-wide collections. FAO, Rome, pp. 462.

Index

abscisic acid 27 acclimatisation of plantlets 88–89 adult trees, grafting on 62–64, 73 adventitious root formation 22–27, 33 annual cycle 11 anthesis 16 asexual reproduction 2, 6 auxin treatment, of cuttings 35–41 improved effectiveness of 39–40 used with root-promoting compounds 40–41 auxin-talcum powder mixtures 37 auxins 26, 35–39, 87–88 axenic culture 82 bark grafting 58–61, 64 basal wounding 40 bench grafting 73 biological cycle 8–10 branches 28–30 bridge grafting 44 budding 45 callus 45, 62 certification 126–128 clonal rootstock, 27, 45 production of 73–74 closed frame technique 118 collar rot 53 collection of explants 81 commercial rooting preparations 39 containers, for in vitro propagation 87 for transplanting 120–121, 123 cryopreservation 92, 93 cultivars, certification of 125–128 rooting ability 38, 131–132 culture, axenic 82

culture medium, addition of zeatin formulation 84, 85, 87 oxidation 83 preparation 86 cultures, initiation of 82–83 cuttings 6–7 auxin treatment 35–41 basal wounding 40 grafted 73 plant material 27–33 preparation 34–41 propagation 6, 22–42 root development 22–27 soaking 39–40 cytokinins 26–27, 41, 84, 87 disinfection of explants

82–83, 94

embryo 20 embryogenesis, somatic 90–91 empty seeds 53 environmental conditions for rooting 113–118 ethylene 27, 41 explant, collection 81 disinfection 82–83, 94 oxidation 83 feather grafting 58 fertigation 123–125 fertilisation at second transplant 123–125 flower differentiation 13 flower induction 12–13 flowering period 17 flowers 9, 14–16, 18 fog 115, 117

fruit drop 18 fruit growth 19–21 fruit set 18 fruiting cycle 11 fungicide treatment of seedbeds fungicides 41

genetic origin, certification 127–128 genotype 2 germinability 46–49 germination 48–49, 51–52, 53–54 germplasm collections 127, 133–136 germplasm conservation 129–130 gibberellins 27, 41 graft incompatibility 58 graft union, histology 55–58 grafted cuttings 73 grafted plants, 74–75 care of 62 grafting, 6, 45, 56–58 on adult trees 62–64, 73 bark 58–61, 64 bench 73 bridge 44 disadvantages of 45 environmental conditions 58 feather 58 micro- 91–92 preparation of rootstock 61 productivity 61 propagation by 43–76 purposes 43–44 scionwood collection for 59 on seedling rootstock 46, 58–62 of seedlings 46 on suckers 63–64 terminology 45 topworking 45, 4 on wild olive trees 64,73 grafting area 55 grafting time 59 grafting tools and accessories 62 grafting wax 62

greenhouses, 81, 99–111 beds 106 computers for ambient control 106 control zone 106–107 cooling 104–106 covering materials 99–102 glass 100 hardening zone 110, 121–122 heating 103–104, 108, 110 irrigation 114–118 location 102–103 misting 121 plastic film 101 propagation 106–110 rigid panel 101–102 rooting zone 107–108, 110 size 96 ventilation 104 waterlogging 108 zones 106–110 growth regulators 26, 36, 40–41, 84–88 hardening of rooted cuttings 119–122 harvesting, 125–126 of seeds 49 heating, for seed germination 53 greenhouses 103–104, 108, 110 humidity 114–118 hydro-alcoholic solutions 35–37 in vitro propagation 77–95 in vitro storage 92–93 inarching 44 inflorescence 13–14 initiation of cultures 82–83 irrigation, 120, 123–125 fog 115, 117 in greenhouses 114–118 misting 108, 114–117, 120 in nurseries 114–118 of seedbed 53 knelt seedlings

52–53

Index

life cycle 8–10 light 118 machine harvesting 124–125 Mediterranean maquis 64 micrografting 91 micropropagated plants, field performance 89–90 micropropagation, 77–92 acclimatisation of plantlets 88–89 collection of explants 81 containers 87 culture conditions 87 culture medium 83–87 features of 77 initiation of cultures 82–83 laboratory 78–80 root induction 88 root promotion 87–88 shoot elongation 87 shoot proliferation 83–87 shoot rooting 87–88 stages of 77–78, 81–89 subculturing 87 micropropagation medium, addition of zeatin 86 formulation 84, 85, 87 oxidation 83 preparation 86 misting 108, 114–117, 120 multiplication 2, 6 nurseries, 96–128 grafting area 55 greenhouses 99–111 irrigation 114–118 shelters 110 size 96–97 stock plants 97–99 storage areas 10–111 sundry materials 97 water requirements 97

139

olive biology 6–21 olive cycles 8–11 olive domestication 3, 4, 6 olive oil consumption 1 olive stones 46 cleaning 50 extraction 49–50 quantity required for sowing 53 soaking 51 sources of 47–48 storage 50 weight 53 olive trees, history 1–7 ovary abortion 16 ovules 30–32 oxidation of explants and medium 83 peat 112–113 perlite 111–112 phase change, of seedlings 98 phenolic compounds 27 phenotype 2 plant certification 125–128 plant growing (2nd transplant) 122–125 plant material, for cuttings 27–33 plant spacing 125 plant training 125–126 plantlets, acclimatization of 88–89 plants, grafted vs. self-rooting 74–75 pollards 33 pollination 16, 18 polyamines 27, 41 propagation, 2–3 by cutting 6, 22–42 by grafting 43–76 history 6–7 in vitro 77–95 by micropropagation 77–90 by seed 51–55 propagation cycles, timing of 124 propagation greenhouses 106–110 pruning 75, 125

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quick-dip treatments

35–37

reproduction, 2 stages of 12–21 reproductive structures 9, 12–16, 18–21 root development 22–27, 33, 74–75 root induction 35–37, 88 root initiation 26–27 root-promoting compounds 35–41, 87 rooted cuttings, transplant 119–125 rooting ability, of cultivars 38, 131–132 rooting bed, closed frame technique 118 rooting shoot 87 rooting substrates 111–113 rooting treatments 35–41, 88 rooting, environmental conditions 113–118 roots, adventitious 22–27, 33 rootstock, 27, 45, 46 clonal 72–73 preparation for grafting 61 sand 113 sanitary conditions 128 scarification 51–52 scions 45, 46 scionwood, collection and storage 59 sclerification 19 seed development 20 seed dormancy 46–47 seed propagation 51–55 seedbed 52, 53 seedling rootstock production 57 seedling rootstock, grafting on 46, 58–62 seedlings, production 46–55 growth 55 knelt 52–53 phase change 98 quality 49 transplanting 55 seeds, 20–21, 46 empty 53 harvesting 49

germinability 46–49 germination 48–49, 51–52, 53–54 quality 47–48 scarification 51–52 sowing 52 synthetic 91 valuable 54 self-rooted vs. grafted plants 74–75 shelters 110 shoot collection 33–34 shoot elongation 87 shoot proliferation 83–87 shoot propagation 27, 30 shoot rooting 87–88 slow growth storage 92 soaking cuttings 39–40 sodium hypochlorite 82, 94 somatic embryogenesis 90–91 sterility 18 stock plants, 45, 97–99 phase change 98 training systems 98–99 stones see olive stones storage areas 110–111 storage, in vitro 92–93 subculturing 87 substrates, 53, 97 for growing plants 113 rooting 111–113 for transplanting 120–121 suckers, 33 grafting on 63–64 synthetic seeds 91 talc formulations 37–38 temperature 58, 113–114 tissue disinfection 82–83 topworking 45, 62–64 transplanting, 55, 75 containers 120–121, 123 environment 121–122 first 119–122 pruning 75 second 122–125

Index

vermiculite

113

warm beds 53 water distribution techniques 114â&#x20AC;&#x201C;118 water supply, instruments for control of 117

water required by nurseries 97 watering see irrigation wild olive trees, grafting on 64, 73 wounding cuttings 40 zeatin

84, 86

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