ARISTOTLE UNIVERSITY of THESSALONIKI FACULTY of SCIENCES, SCHOOL of BIOLOGY DEPARTMENT of BOTANY SHARAF MUSA AL-TARDEH B. Sc. Bi

ARISTOTLE UNIVERSITY OF THESSALONIKI FACULTY OF SCIENCES, SCHOOL OF BIOLOGY DEPARTMENT OF BOTANY

SHARAF MUSA AL-TARDEH B. Sc. Biological Sciences M.Sc. Food Quality Management

MORPHOLOGICAL AND ANATOMICAL ADAPTATIONS OF THE PERENNIAL GEOPHYTE URGINEA MARITIMA (L.) BAKER (LILIACEAE) TO THE MEDITERRANEAN CLIMATE

DOCTORAL DISSERTATION

Thessaloniki 2008

SHARAF MUSA AL-TARDEH B. Sc. Biological Sciences M.Sc. Food Quality Management

MORPHOLOGICAL AND ANATOMICAL ADAPTATIONS OF THE PERENNIAL GEOPHYTE URGINEA MARITIMA (L.) BAKER (LILIACEAE) TO THE MEDITERRANEAN CLIMATE

DOCTORAL DISSERTATION

Submitted to the School of Biology, Faculty of Sciences, Aristotle University of Thessaloniki

Presented for public criticism on the 28th of November 2008

MORPHOLOGICAL AND ANATOMICAL ADAPTATIONS OF THE PERENNIAL GEOPHYTE URGINEA MARITIMA (L.) BAKER (LILIACEAE) TO THE MEDITERRANEAN CLIMATE

Figures on the cover page: (A) TEM micrograph of nectary of Urginea maritima showing the nectary epithelium cell with nectar in crystallized form at the old stage; (B) Photo of inflorescence stalk; (C) SEM micrograph of anther; (D) LM micrograph of leaf with parallel veins and veinules; (E) TEM micrograph of leaf with sieve pore between the sieve element; (F) LM micrograph of bulb scale revealing cells stained red with Schiff’s reagent; (G) LM micrograph revealing cells stained brown-black with Sudan Black B; (H) LM micrograph showing idioblastic cortical cells with bundles of raphides and mucilage stained with TBO; (I) LM micrograph of root showing the 10-arch vascular cylinder; (J) LM micrograph of root showing series of idioblastic cells with bundles of raphides.

To my family

To my lovely wife “WOROD”

To those who defend their rights

SHARAF

ELABORATION OF THE DOCTORAL DISSERTATION Place Aristotle University of Thessaloniki, Faculty of Sciences, School of Biology Department of Botany

Time February 2004 – November 2008.

Examining committe • Stylianos Delivopoulos, Professor, Department of Botany, School of Biology, A.U.Th. Supervisor • Thomas Sawidis, Associate Professor, Department of Botany, School of Biology, A.U.Th. Cosupervisor • Barbara-Evelin Diannelidis, Assistant Professor, Department of Botany, School of Biology, A.U.Th. Cosupervisor

• Bosabalidis Artemios, Professor, Department of Botany, School of Biology, A.U.Th. Examiner • Eleftheriou Eleftherios, Professor, Department of Botany, School of Biology, A.U.Th. Examiner • Moustakas Michael, Associate Professor, Department of Botany, School of Biology, A.U.Th. Examiner

• Pantis John, Associate Professor, Department of Ecology, School of Biology,

A.U.Th. Examiner

Morphological and anatomical adaptations of the perennial geophyte Urginea maritima (L.) Baker (Liliaceae) to the mediterranean climate. ISBN

Approval of the present Doctoral Dissertation by the Department of Botany, School of Biology, Aristotle University of Thessaloniki does not imply the acceptance of the opinions of the author (N. 5343/1932, law 202, paragraph 2).

Sharaf Al-Tardeh Table of contents

TABLE OF CONTENTS

ELABORATION OF THE DOCTORAL DISSERTATION……………………...... i TABLE OF CONTENTS…………………………………………………………...... iii PROLEGOMENA ……...………………………………………………………...... vii LIST OF ORIGINAL ARTICLES……….………...... ix LIST OF ABBREVIATIONS AND SYMBOLS………………………………..…… x SUMMARY...………………………………………………………..…………...... xii ΠΕΡΙΛHΨΗ…………………………………………………………………………... xvi CHAPTER I: INTRODUCTION…………...……………………………………… 1 1.1 Mediterranean climate……………………………………………………………. 1 1.1.1 Introduction…………………...…………………………………………….… 1 1.1.2 Spatial patterns of temperature in the Mediterranean climate…………….…... 1 1.1.3 Spatial patterns of precipitation in the Mediterranean climate………………... 2 1.2 Geophytes……………………………………………………………………….... 3 1.2.1 Introduction…………………………………………………………………… 3 1.2.2 Taxonomy and origin of geophytes…………………………………………... 3 1.2.3 Diversity of geophytes………………………………………………………... 3 1. 2.3.1 Bulbous plants………………………………………………………...... 4 1.2.3.2 Tuberous plants…………………………………………………………… 4 1.2.4 The body of geophytes……………………………………………………...... 5 1. 2.4.1 Storage organs and tissues…………….…………………………………. 5 1.2.4.2 Aerial organs and tissues………..………………………………………… 6 1. 2.4.3 Roots……………………………………………………………………… 6 1.2.4.4 Tunics…………………………………………..…………………………. 7 1.2.5 Periodicity and dormancy of flower bulbs…………………………………….. 7 1.2.5.1 Definitions…………………………………………………………………. 7 1.2.5.2 Factors that affect bulb dormancy…………………………………………. 8 1.2.6 The flowering process……………………………………………………...... 9 1.2.7 Types of hysteranthous geophytes…………………………………………….. 9 1.2.8 Factors affecting bulb growth and development……………………………… 10 1.2.8.1 The size of the bulb……………………………………………………...... 10 1.2.8.2 Temperature………………………………………………………………... 10 1.2.8.3 Light……………………………………………………………………….. 11 1.2.8.4 Soil features……………………………………………………………...... 11 1.2.9 World production of geophytes……………………………………………..... 12 1.3 Chlorophyll contents and fluorescence…………………………………………... 13 1.4 Nectaries and nectar…………………………………………………………….... 17 1.4.1 Introduction…………………………………………………………………… 17 1.4.2 Definitions of nectary…………………………………………………………. 17 1.4.3 Basic types of floral nectaries.…..…………………………………………….. 18 ii

Sharaf Al-Tardeh Table of contents

1.4.4 Floral and extrafloral nectaries………………………………………………... 20 1.4.5 Gynopleural (septal) nectaries……………………………………………….... 20 1.4.5.1 Distribution of the gynopleural nectaries………………………………….. 20 1 4.5.2 Structure of the gynopleural nectaries…………………………………...... 21 1.4.5.3 Nectar secretion in the gynopleural nectaries…………………………….... 22 1.4.5.4 Fate of the gynopleural nectaries…………………………………………... 22 1.4.6 Nectar production and presentation…………………………………………… 23 1.4.6.1 Nectar secretion mechanism……………………………...... 23 1. 4.6.2 Nectar presentation………………………………………………………... 24 1.4.6.3 Fate of nectar and nectarines………………………………………………. 26 1.5 The summer flowering geophyte: Urginea maritima (L.) Baker (Liliaceae)…...... 27 1.5.1 Urginea maritima overview…………………………………………………... 27 1.5.2 Morphology…………………………………………………………………… 27 1.5.3 Growth, development and flowering……………………………………….… 28 1.5.4 Nectary of U. maritima……………………………………………………….. 29 1.5.5 Pollination in U. maritima as a model of hysteranthous geophytes………...... 29 1.5.6 Bulb production and/or propagation………………………………………….. 30 1.5.7 Horticultural usage……………………………………………………………. 32 1.5.8 Biochemical principles and their action………………………………………. 32 1.5.8.1 Toxic principle……………………………………………………………... 32 1.5.8.2 Defense mechanisms………………………………………………………. 33 1.5.8.3 Pharmaceutical advantages of U. maritima……………………………….. 34 1.6 Aim of the thesis…………………………………………………………………. 35 CHAPTER II: MATERIALS AND METHODS ……………………………….... 37 2.1 Materials………………………………………………………………………….. 37 2.1.1 Plant samples………………………………………………….……………..... 37 2.1.2 Apparatuses……….…………………………………………...……………… 38 2.1.3 Experimental design…………………………………………………………... 40 2.2 Methods…………………………………………………………………………… 42 2.2.1 Water content………………………………………………………………….. 42 2.2.2 Fixation………………………………………………………………………... 43 2.2.2.1 Solution preparations………………………………………………………. 43 2.2.2.2 Procedure…………………………………………………………………... 44 2.2.3 Sectioning……………………………………………………………………... 45 2.2.4 Staining techniques…………………………………………………………… 46 2.2.4.1 Staining with toluidine blue O (0.1%)…………………………………….. 46 2.2.4.2 Staining with Sudan Black B (1%)………………………………………... 46 2.2.4.3 Staining with Periodic acid-Schiff’s reaction (PAS reaction)…………….. 47 2. 2.4.4 Staining with uranyl acetate and lead citrate…………………………….. 48 2. 2.5 Scanning Electron Microscopy (SEM)……………………………………….. 48

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Sharaf Al-Tardeh Table of contents

2.2.6 Maceration…………………………………………………………………….. 49 2.2.6.1 Maceration of soft material………………………………………………... 49 2.6.2 Maceration of hard material…………………………………...…………….. 50 2.2.7 Leaf chlorophyll assay………………………………………………………... 50 2.2.8 Chlorophyll a fluorescence…………….……………………………………… 50 2.2.9 Morphometry…………………………………………………………………. 52 2. 2.10 Mathematical analysis of morphological and anatomical features of the plant tissues………………………………………………………………….... 52 2.3 Statistical analysis………………………………………………………………... 52 CHAPTER III: RESULTS ………………..……………………………………….. 53 3.1 ANATOMICAL STUDIES ON THE ADVENTITIOUS ROOTS……………... 53 3.1.1 Root morphology ………………………………………………………..……. 53 3.1.2 Root anatomy………………………………………………………………….. 53 3.1.3 Root histochemistry…………………………………………………………… 56 3.1.4 Root morphometry……………………………………………………………. 56 3.2 Biomass and reserve allocation patterns within the bulb………………………… 58 3.2.1 Bulb morphology……………………………………………………………… 58 3.2.2 Bulb anatomy…………………………………………………………………. 58 3.2.3 Bulb histochemistry…………………………………………………………… 60 3.2.4 Bulb morphometry and reserves allocation patterns………………………….. 61 3.2.5 Biomass allocation patterns and water status…………………………………. 62 3.3 MORPHO-ANATOMICAL FEATURES OF THE LEAVES…………………. 65 3.3.1 Leaf anatomy………………………………………………………………….. 65 3.3.2 Leaf morphology and morphometry…………………………………………... 67 3.3.3 Leaf histochemistry…………………………………………………………..... 69 3.3.4 Leaf chlorophyll assay……………………………………………………….... 69 3.3.5 Chlorophyll a fluorescence……………………………………………………. 70 3.4 MORPHO-ANATOMY OF THE INFLORESCENCE…………………………. 73 3.4.1 Flower morphology……………………………………………………………. 73 3.4.2 Anatomy and development of the gynopleural (septal) nectary………………. 74 3.4.2.1 Young stage nectary……………………………………………………….. 74 3.4.2.2 Nectary at the intermediate stage…………………………………………... 75 3.4.2.3 Nectary at old stage……………………………………………………….... 77 3.4.3 The ovary and ovules………………………………………………………..…. 78 3.4.4 Obturator gland……………………………………………………………….... 79 CHAPTER IV: DISCUSSION…………………………...... 170 4.1 ADVENTITOUS ROOTS………………………………………………………... 170 4.1.1 Root system……………………………………………………………………. 170 4.1.2 Special adaptation of roots…………………………………………………….. 170 4.1.3 Velamen………………………………………… ……………………………. 171

Sharaf Al-Tardeh Table of contents

4.1.4 Dimorphic hypodermis………………………………………………………... 172 4.1.5 Cortex…………………………………………………………………………. 173 4.1.6 The storage tissue……………………………………………………………... 173 4.1.7 Water economy…………………………………………………………...... 174 4.1.8 Raphides………………………………………………………………………. 175 4.1.9 Vascular cylinder…………………………………………………..………….. 177 4.2. BIOMASS AND RESERVE ALLOCATION PATTERNS WITHIN THE BULB………………………………… …………………………………………… 177 4. 2.1 Morpho-anatomy of the bulb………………………………………………….. 177 4.2.2 Tunics…………………………………………………………………………. 178 4.2.3 Biomass, resources allocation and plant growth……………………………… 178 4.2.4 Histochemistry and the storage tissue………………………………………… 181 4.2.5 Defense mechanisms…………………………………………………………... 181 4.2.6 Water contents………………………………………………………………… 183 4.3 MORPHO-ANATOMICAL FEATURES OF THE LEAVES ………………….. 184 4.3.1 Leaf anatomy………………………………………………………………….. 184 4.3.2 Epidermis and cuticle…………………………………………………………. 184 4.3.3 Stomata as an adaptive strategy………………………………………………. 184 4.3.4 The mesophyll ground tissues……………………………………………...... 185 4.3.5 The vascular system…………………………………………………………… 186 4.3.6 Leaf morphology………………………………………………………………. 187 4.3.7 Leaf histochemistry……………………………………………………………. 189 4.3.8 Photosynthetic efficiency of the leaf of U. maritima…………………………. 190 4.3.8.1 Chlorophyll a content…………………………………………….………... 190 4.3.4.2 The basis of chlorophyll fluorescence measurements……………………... 191 4.4 MORPHO-ANATOMY OF THE INFLORESCENCE ………………………… 194 4.4.1 Gynopleural (septal) nectary………….……………………………………….. 194 4.4.1.1 Nectary epithelial cells……………………………………………………... 194 4.4.1.2 Subsidiary tissue…………………………………………………………… 195 4.4.1.3 Amyloplasts and nectar secretion………………………………………….. 196 4.4.1.4 Fate of the nectary…………………………………………………………. 199 4.4.1.5 Nectar presentation and pollinators………………………………………... 200 4.4.2 Contents of raphides………………………………………………………….. 201 4.4.3 The obturator gland…………………………………………………………… 202 CHAPTER V: CONCLUSIONS…………………………………………………… 203 ACKNOWLEDGEMENTS...... 207 CHAPTER VI: LITERATURE CITED.…………………………………………... 208 ORIGINAL PUBLISHED ARTICLES………………………………………..…….. i

Sharaf Al-Tardeh Prolegomena

PROLEGOMENA

The thanks I owe for this PhD are rather as many as the words that it consists of, probably equally important and definitely harder to express. I would, nevertheless, like to express my gratitude to everyone, who was, as meant to be, present or absent.

I would like to express my deep gratitude to my supervisor Prof. Stylianos Delivopoulos for many things, most of which are difficult to express in words. I thank him for his continuous assistance, guidance, encouragement and friendship during the time I have spent at AUTH, Greece. Moreover, I would like to express my deep gratitude for his kindness and hospitality, for his quantity of enthusiasm and quality of interest. But most of all I thank him for having been both a teacher and a ‘father’ to me.

I would also like to express my gratitude to Dr. Thomas Sawidis for helping me in the subject and scientifical revision of my work. I thank him for his kind heart which has boosted up my enthusiasm and emotions. I will never forget his assistance, discussions and joke especially about the “noble price”. I also thank Dr. Barbara-Evelin Diannelidis for her endless assistance and encouragement.

I am deeply grateful to the following botanical staff; Prof. Ioannes Tsekos who gave me the opportunity to elaborate my Ph.D at the Aristotle University of Thessaloniki before his retirement. In addition, I would like to express my very deep gratitude for his assistance and critical suggestions that he has made on my research. I also thank Dr. George Nikolaidis, Maria Moustaka and Michael Moustakas for their help and technical support especially in the use of their microscopes. Thanks are also due to Dr. Stylianos Kokkini, Artemios Bosabalidis and Eleftherios Eleftheriou for their moral support and encouragement. I am deep grateful to Dr Emmanuel-Nicholas Panderis for his friendship, jokes and some scientific assistance. Finally, I would like to thank everybody in the Department of Botany for the joyful atmosphere created in the lab during my research work.

I also thank the SEM technician Mr. Stavros Oikonomidis for his help in carrying out the SEM protocol. I would like to express my deep gratitude to Mr. Tasos Makrantonakis and Thanasis Nikodimos for their help, moral support and lovely atmosphere created in

Sharaf Al-Tardeh Prolegomena the TEM laboratory. I also thank Mrs. Soula Sapountzoglou (mama tou kosmou), Pavlos Gaithatzis and the secretary of the Department Mrs Arete Dimopoulou.

I express my sincere thanks to my colleagues who, with their behaviour, gave a unique meaning of the concept “team work” and academic ethics. Those folks are Ioannes Akdamakis, Spyros Gelis, Natasa Tsirika, Giorgos Kofidis, Katerina Aligizaki, and Kalopesa Eleni for their constant encouragement and making a friendly atmosphere in the lab. They made sure not let a day pass without joking, smiling and drinking as well.

I also wish to thank my friends, Iyad, Khalid, Gada, Alaa, Haroon, Dr Saed, Dr Amjad, Dr Feras, Dr Rezq and Amjad Awad for their endless moral support and encouragement. They showed me the ethics of friendship and the brotherhood.

Finally, I would like to express my endless gratitude to my father Musa and my mother Mahbouba (may God bless her and meet herin the paradise, ameen), my 9 brothers and 6 sisters, all my uncles, nephews and nieces for their love, help, moral support and encouragement. In addition, I would like also to thank the family of Al-Sharabati whom my wife belongs to, especially her mother “Boshra”. Moreover, I would like to give a warm and very special thanks to my wife “Worod” for her having waited for me to finish my PhD thesis, and also for her encouragement, enthusiasm, and confidence for big heart and love.

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Sharaf Al-Tardeh List of original articles

LIST OF ORIGINAL ARTICLES

I. Al-Tardeh, S., Sawidis, T., Diannelidis, B-E., and Delivopoulos, S. 2006. Anatomical studies on the adventitious roots of the geophyte Urginea maritima (L.) Baker. J. Biol. Res., 5: 61-70.

II. Al-Tardeh, S., Sawidis, T., Diannelidis, B-E., and Delivopoulos, S. 2008. Water content and reserve allocation patterns within the bulb of the perennial geophyte Urginea maritima (Liliaceae) in relation to the Mediterranean climate. Botany, 86: 291-299.

III. Al-Tardeh, S., Sawidis, T., Diannelidis, B-E., and Delivopoulos, S. 2008. Morpho- anatomical features of the leaves of the Mediterranean geophyte Urginea maritima (L.) Baker (Liliaceae). J. Plant Biol., 51: 150-158.

IV. Al-Tardeh, S., Sawidis, T., Diannelidis, B-E., and Delivopoulos, S. 2008. Nectary structure and nectar presentation in the Mediterranean geophyte, Urginea maritima (Hyacinthaceae). Botany, 86: 1194 -1204.

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Sharaf Al-Tardeh List of abbreviations and symbols

LIST OF ABBREVIATIONS AND SYMBOLS

Abc: abaxial cuticle Gc: guard cell Abe: abaxial epidermis Gi: Golgi body Adc: adaxial cuticle Gr: grana Ade: adaxial epidermis HCL: hydrochloric acid AL : actinic light Id: idioblastic cell As: air space Is: intercellular spaces

ATPase: adenosine triphosphatase K2S2O5: Potassium metabisulfite AUTh: Aristotle University of Ld: Lower epidermis Thessaloniki Lg: ledges Ba: bacteria LM: light microscope Bs: bundle sheath LTD: leaf tissue density Ca: callose LWC: leaf water content Cc: companion cell Mc: mucilage cell Ch: chloroplast Mi: mitochondria Chl b: chlorophyll b ML: modulated measuring rate Chla: chlorophyll a Ms: myelin-like structure Ci: crystalloid inclusion Mx: metaxylem

Cn: crystal needles Na2B4O7.10 H2O: boric acid

CO2: carbon dioxide NaOH: sodium hydroxide Cu: cuticle Nc: nucleus d. H2O: double distilled water Ne: nectary epithelium DM: dry mass Ny: nectary Ed: endodermis O: ovule Edm: electron dense material Ob: obturator Edr: electron dense remnants oC: Celsius EM: electron microscope Oc: oil cell Em: embryo OD: obtical density Ep: epidermis Od: oil droplets ER: endoplasmic reticulum Os: osmiophilic droplet

EtOH: ethanol OsO4: Osmium tetroxide ETR: electron transport rate PAR: photosynthetic active radiation FR: far red light PAS: periodic acid-Schiff’s reagent

Sharaf Al-Tardeh List of abbreviations and symbols

Pc: parenchyma cell SLM: specific leaf mass PCD: programmed cell death SP: saturated light pulse Pd: plasmodesmata Sp: sieve pore Ph: phloem St: subsidiary tissue Pl: plastid Sv: secondary veins (veinules) PSI: photosystem I TBO: toluidine blue PSII: photosystem II TEM: transmission electron Pv: parallel vein microscope Px: protoxylem TWC: Total water content qN: non photochemical quenching Ud: upper epidermis qP: photochemical quenching UV: ultra violet light Rb: raphide bundle Vb: vascular bundle Rh: root hair Vc: vacuole Rn (Rb): raphide needles Ve: velamen SA: surface area Vs: vesicles Sc: sclerenchyma fibers WC: water content Sch: stomatal chamber Ws: water-storing cell SD: standard deviation X: xylem SE: sieve element ΦPSII: yield

SEM: scanning electron microscope. (C6H5O7)2Pb3.3H2O: lead citrate Sg: starch granules

Sharaf Al-Tardeh Summary

SUMMARY

Urginea maritima is a winter plant characterized by three distinct phenophases, consisting of the inflorescence, leaves and no above-ground biomass, respectively. The inflorescence stalk appears during August or September (late summer). Leaves first appear after the flowers have wilted in response to winter rains onset during November or December, and may remain green until late spring (May), depending on rainfall and temperature. The plant falls into dormancy from late spring to late summer.

The root system of U. maritima consists of adventituos roots which are attached to the basal plate disk. The bulb of about 10 cm diameter possesses 5-10 adventitous roots which could have up to five branches. The roots are characterized by the presence of a multi-layered velamen and a dimorphic hypodermis. The velamen provides mechanical protection to the cortex and reduces water loss. It may also function in the absorption of water and minerals. Both of the epidermis and endodermis of the root are uniseriate.

The cortex is 741.3 ± 51.34 µm thick and is composed of numerous large parenchyma cells with a storing character. Scattered idioblastic cells with bundles of raphides of calcium oxalate of 72 ± 22 µm length, occur in the cortical cells. The latter are vertically oriented and located in rows around the central cylinder. Cells containing in their vacuoles soluble polysaccharides or lipids are also present. Morphometrical analysis shows that idioblastic cells occupy 19.83 ± 1.10% of the cortex relative volume, cells containing lipids 14.38 ± 0.71%, and cells containing polysaccharides 11.27 ± 1.62%. The cortex storage cells occupy 34.11% of the total root volume. The average volume of the cortical cells is 73143 µm3.

The vascular cylinder is usually 10-arched. The root xylem consists of vessels in short radial rows. The phloem consists of sieve elements located between the vessel rays. The core of the central cylinder is sclerenchymatous in the older roots. U. maritima possessing adventitious roots proves to be efficient in storing water during the long summer drought, less susceptible to climatic stress and well synchronized with the climatic fluctuations.

Sharaf Al-Tardeh Summary

The different bulb parts serve different ecological functions in terms of their resources and their importance for these functions. The basal plate is the active centre, develops one or two apical meristems and roots in autumn, as well as the flowering bud in late summer. The middle region of the bulb (almost the 3rd bulb-scale) stores the resources and the tunics provide mechanical defense.

The bulb-scales are covered by hard modified bulb-scales, the tunics. The bulb-scales are coated with a smooth cuticle 12 ± 4 µm thick. The cells containing raphides (432 ± 54 µm long) are 2-6 fold longer than the neighbouring ones in longitudinal section. Bulb- scales consist of cells with storage character that contain mucilage, polysaccharides and lipophilic substances. The vascular bundles are collateral, scattered and numerous.

The middle region of the bulb contains a considerable relative volume of cells with polysaccharides (12.91 ± 2.43%) and the highest one of those with lipids (21.92 ± 2.43%) and mucilagenous material (9.34 ± 1.03%). The middle region of the bulb is the most efficient part in storing reserves, since the total relative volume of its storage cells is 50.66%. The basal plate exhibits the highest relative volume of polysaccharidic contents (15 ± 1.62%) and raphide inclusions (13.09 ± 1.67%). However, the lipid gradient decreases in the outermost part of the bulb and the basal plate (0%). In addition, biomass allocation patterns synchronize the plant’s phenological development with the seasonality of the Mediterranean climate. The synchronization of the plant timing with the periodicity of the Mediterranean- type of climate is to be accomplished by means of biomass and nutrient trade-offs which take place in the different plant parts.

The leaf of U. maritima is 714 ± 33.6 µm thick and possesses moderate specific leaf mass (8.564 ± 3.139 mg cm-2) and low leaf tissue density (136.5 ± 14.41 mg cm-3). The epidermal cells are compactly arranged and covered by cuticle (10.25 µm thick). The average density of sunken anomocytic stomata (45.4 ± 2.3 µm length) in the upper and lower epidermis (amphistomaty) of the leaf is 736 ± 105.35 mm-2 and 867 ± 66.09 mm-2 respectively. The mesophyll is 594.7 ± 33.3 µm thick. The mesophyll cells occupy 52.96 ± 1.42% of the total leaf volume, while the mesophyll intercellular spaces and the air spaces occupy 30.41 ± 1.13% of the total leaf volume. Idioblastic cells with bundles of raphides and crystalloid inclusions of different phenotypes, embedded in polysaccharides, occur in the lower side of the mesophyll. osmiophilic droplets occur in xii

Sharaf Al-Tardeh Summary the chloroplasts and the cytoplasm. Large bundles are associated with sclerenchyma for mechanical support and/or defense. However, in the smaller bundles only the phloem is associated with sclerenchyma, while the xylem is subtended by parenchyma with oil and polysaccharide contents for chemical defense. Bundle sheath does not possess chloroplasts which are of a pronounced C3 plant character.

U. maritima is a deciduous plant possessing mesomorphic leaves with a flat lamina. The leaves are adapted to the Mediterranean climate by possessing equifacial leaves in order to achieve a possible higher photosynthetic efficiency. This could be achieved by shortening the distances of CO2 diffusion, the high density of the sunken stomata on both sides, the fine structure of the mesophyll cells, the distribution of the chloroplasts parallel to the cell wall facing the intercellular spaces, the large air spaces in the mesophyll area and the low LTD and SLM.

The flowering shoot of U. maritima is characterized by an interesting one-dimensional growth system and rapid pattern of subapical elongation. Elongation of the inflorescence axis is accompanied by opening of the flowers in an acropetal direction. Stomata are present on the epidermis of both the anther and the ovary. When the flowers senesce, the petals close up again to protect the growing fruit capsule. When the fruit is ripe, it splits open to release its black winged seeds.

U. maritima has a typical gynopleural (septal) nectary with secondary presentation. The nectary consists of one layer of epithelial secretory cells and 1-4 layers of subsidiary tissue followed by 2-6 layers of parenchyma cells. The nectary outlets the nectar at a distance of about two-thirds from the top of the ovary by means of the carpellary suture. Nectar secretion mechanism largely depends on the hydrolysis of the starch grains stored in the plastids (amyloplasts) at the intermediate age nectary.

The young stage of the nectary is characterized by the endoplasmic reticulum dominating the cytoplasm of the secretory epithelium. The cytoplasm is densely packed with ribosomes attached to the ER. Development of the starch granules commences in the secretory epithelium cells, which contain mostly elongated plastids. The intermediate stage of the nectary is characterized by the dense and granular cytoplasm of the epithelial cells with an active and well-developed rough ER. Mitochondria and dictyosomes, also, xiii

Sharaf Al-Tardeh Summary dominate the cytoplasm of the epithelium. Plastids are of various shapes and contain osmiophilic granules. Numerous secretory vesicles, myelin-like figures and vacuoles with electron dense materials are also present.

The old stage of flower and/or nectary starts 19 hours after the flower opening. This stage is characterized by a crystallized form of nectar inside the epithelial cells, collapse of the cortical parenchyma cells, complete starch hydrolysation and disappearance of the amyloplasts and the ER. The epithelial and subsidiary cells undergo cytological modifications; some subsidiary cells degenerate and/or transform into parenchyma cells, while the epithelial cells become more elongated. However, the nectary is still persist and thinner; it is approximately six cells wide from the epithelium to the ovary wall. The microchannels are more conspicuous in the cell wall of the epithelial cells

The ovary of U. maritima is partitioned into three locules formed by the fusion of three carpels. The latter contain several ovules, four per each one at the mature stage. After fertilization about 12 seeds per capsule are produced. The ovary wall consists of approximately 3-10 layers of parenchyma cells. The ovarian vascular system is simple and looks like that of the leaf, bulb and root. However, the vascular tissue consists of only xylem vessels at the inferior side of the nectary, only sieve elements on both sides of the nectary and disappears at the high level of the ovary. The obturator is a placental protuberance secreting mucilage of a polysaccharidic nature, which may be involved in the heterotrophic growth of the pollen tubes or in controlling the direction of their growth.

The presence of cells with polysaccharidic content is evident in the tissues of the ovarian body, the obturator and the nectary. However, lipophilic substances are absent. Moreover, idioblastic cells containing bundles of raphides occur in the ovarian wall and the subsidiary tissues. The highest relative volume of idioblastic cells containing bundls of raphides is recorded at the young stage (7.41 ± 1.8 %), it decreases at the intermediate stage (4.33 ± 1.8 %) and increases again at the old stage (6.45 ± 1.55 %).

Keywords: adaptation, adventitious root, amyloplast, bulb, geophyte, gynopleural nectary, leaf, Mediterranean climate, morphometry, nectar secretion, starch, Urginea maritima, water content. xiv

Sharaf Al-Tardeh Summary

ΠΕΡΙΛHΨΗ

Η Urginea maritima είναι ένα χειμερινό μονοκόωτο φυτό χαρακτηριζόμενο από τρεις διακριτές φαινοφάσεις που αποτελούνται αντίστοιχα από την ταξιανθία, τα φύλλα και καθόλου υπέργεια βιομάζα. Ο ανθοφόρος άξονας εμφανίζεται αργά το καλοκαίρι (Αύγουστο ή Σεπτέμβριο). Τα φύλλα εμφανίζονται, αρχικά, αφού τα άνθη έχουν μαραθεί με την έναρξη των βροχοπτώσεων το Νοέμβριο ή Δεκέμβριο και μπορεί να παραμένουν πράσινα έως αργά την άνοιξη (Μάϊο), γεγονός που εξαρτάται από τις βροχοπτώσεις και τη θερμοκρασία. Το φυτό πέφτει σε λανθάνουσα κατάσταση από αργά την άνοιξη έως το τέλος του καλοκαιριού.

Το ριζικό σύστημα της U. maritima αποτελείται από πρωτογενείς ρίζες που είναι προσκολλημένεs στο δίσκο της βασικής πλάκας. Ο περίπου 10 εκ. διαμέτρου βολβός έχει 5-10 πρωτογενείς ρίζες, καθεμία από τις οποίες μπορεί να φέρει έως 5 κλάδους. Οι ρίζες χαρακτηρίζονται από την παρουσία ενός πολυστρωματικού ριζάμφιου και μιας διμορφικής υποδερμίδας. Το ριζάμφιο παρέχει μηχανική προστασία στο φλοιό και μειώνει την απώλεια νερού. Μπορεί, επίσης, να συμμετέχει στη διαδικασία απορρόφησης του νερού και των μεταλλικών στοιχείων. Η επιδερμίδα και η ενδοδερμίδα της ρίζας είναι μονοστρωματικές.

Ο φλοιός έχει πάχος 741.3 μm και αποτελείται από πολλά μεγάλα κύτταρα αποταμιευτικού παρεγχύματος. Στα κύτταρα του φλοιού υπάρχουν διάσπαρτα ιδιόβλαστα κύτταρα που φέρουν δέσμες βελονοειδών κρυστάλλων οξαλικού ασβεστίου μήκους 72 ± 22μm. Τα κύτταρα του φλοιού είναι κατακόρυφα προσανατολισμένα και σε σειρές γύρω από τον κεντρικό κύλινδρο. Στο φλοιό, είναι εμφανής, επίσης, η παρουσία κυττάρων που περιέχουν στα χυμοτόπιά τους διαλυτούς πολυσακχαρίτες ή λιπίδια. Η μορφομετρική ανάλυση δείχνει ότι τα ιδιόβλαστα κύτταρα κατέχουν 19.83% του σχετικού όγκου του φλοιού, τα κύτταρα που περιέχουν λιπίδια 14.38% και τα κύτταρα που περιέχουν πολυσακχαρίτες 11.27%. Τα αποταμιευτικά κύτταρα του φλοιού καταλαμβάνουν 34.11% του συνολικού ριζικού όγκου. Ο μέσος όγκος των κυττάρων του φλοιού είναι 73143 μm3.

Ο κεντρικός κύλινδρος της ρίζας είναι συνήθως 10-αρχικός. Το ξύλωμα της ρίζας αποτελείται από αγγεία τοποθετημένα σε μικρού μήκους ακτινωτές σειρές. Το φλοίωμα xv

Sharaf Al-Tardeh Summary

αποτελείται από ηθμοστοιχεία τοποθετημένα ανάμεσα στις ακτίνες των αγγείων. Στις μεγαλύτερης ηλικίας ρίζες η κεντρική περιοχή του κεντρικού κυλίνδρου καταλαμβάνεται από σκληρεγχυματικά κύτταρα. Η U. maritima με τις επιγενείς ρίζες της αναδεικνύεται σε ένα φυτό που είναι ικανό ν’αποθηκεύει νερό κατά τη διάρκεια της καλοκαιρινής ξηρασίας, λιγότερο ευαίσθητο στην κλιματική καταπόνηση και καλά εναρμονισμένο στις κλιματικές διακυμάνσεις.

Τα διαφορετικά τμήματα του βολβού εξυπηρετούν διαφορετικές οικολογικές λειτουργίες σε σχέση με τις πηγές τους και την σπουδαιότητα που έχουν όσον αφορά τις λειτουργίες αυτές. Ο δίσκος της βασικής πλάκας αποτελεί το ενεργό κέντρο, αναπτύσσει ένα ή δύο επάκρια μεριστώματα και ρίζες το φθινόπωρο, καθώς επίσης και τον ανθοφόρο οφθαλμό στο τέλος του καλοκαιριού. Το μεσαίο τμήμα του βολβού (σχεδόν το τρίτο λέπιο του βολβού) αποταμιεύει τα θρεπτικά στοιχεία, ενώ οι χιτώνες παρέχουν μηχανική άμυνα.

Τα εξωτερικά λέπια του βολβού (χιτώνες) είναι λεπτότερα και σκληρότερα για λόγους προστασίας. Tα λέπια του βολβού καλύπτονται από λεία εφυμενίδα πάχους 12 ± 4μm. Τα ιδιόβλαστα κύτταρα που περιέχουν ραφίδες (μήκους 432 ± 54) είναι 2-6 φορές μεγαλύτερου μήκους των γειτονικών κυττάρων σε κατά μήκος τομή. Τα λέπια του βολβού φέρουν επίσης ιδιόβλαστα αποταμιευτικά κύτταρα που περιέχουν βλέννα, πολυσακχαρίτες και λιπόφιλες ουσίες. Υπάρχουν, επίσης, άφθονες, διάσπαρτες, αμφίπλευρες αγωγές δεσμίδες.

Η μεσαία περιοχή του βολβού περιέχει ένα σημαντικό, σχετικό όγκο κυττάρων με πολυσακχαρίτες (12.9 ± 2.43%) και το μεγαλύτερο όγκο κυττάρων με λιπίδια (21.92 ± 2.43%) και βλεννώδες υλικό (9.34 ± 1.03%). Η μεσαία περιοχή του βολβού αποτελεί το αποτελεσματικότερο τμήμα αναφορικά με την αποταμιευτική ικανότητα, δεδομένου ότι ο συνολικός σχετικός όγκος των αποθηκευτικών κυττάρων της είναι 50.66%. Ο δίσκος της βασικής πλάκας παρουσιάζει το μεγαλύτερο σχετικό όγκο κυττάρων πολυσακχαριτικού περιεχομένου (15 ± 1.62%) και βελονοειδών εγκλείστων (13.09 ± 1.67%). Ωστόσο, η διαβάθμιση του περιεχομένου των λιπιδίων μειώνεται στο εξωτερικότερο τμήμα του βολβού και ο δίσκος της βασικής πλάκας (σχεδόν 0%). Επιπρόσθετα, τα πρότυπα κατανομής της βιομάζας συγχρονίζουν τη φαινολογική ανάπτυξη του φυτού με την περιοδικότητα του μεσογειακού κλίματος. Ο συγχρονισμός του φυτού με την περιοδικότητα του μεσογειακού τύπου κλίματος επιτυγχάνεται με τη xvi

Sharaf Al-Tardeh Summary

βοήθεια των ανταλλαγών της βιομάζας και των θρεπτικών υλικών που συμβαίνουν στα διάφορα τμήματα του φυτού.

Το φύλλο της U. maritima είναι πάχους 714μm, έχει μέτρια ειδική μάζα (SLM) (8.564 ± 3.139 mg cm-2) και χαμηλή πυκνότητα ιστού (LTD) (136.5 ± 14.41 mg cm-3). Τα επιδερμικά κύτταρα είναι συμπαγώς διατεταγμένα και καλύπτονται από εφυμενίδα πάχους 10.25μm. Η μέση πυκνότητα των βυθισμένων, ανομοιοκυτικών στομάτων (μήκους 45.4μm) στην άνω και κάτω επιδερμίδα του αμφιστοματικού φύλλου είναι 736 mm-2 και 867 mm-2 αντίστοιχα. Το μεσoφύλλο έχει πάχος περίπου 594.7 μm. Τα κύτταρα του μεσόφυλλου καταλαμβάνουν 52.96% του συνολικού φυλλικού όγκου, ενώ οι μεσοκυτταρικοί και οι αεροφόροι χώροι του μεσoφύλλου καταλαμβάνουν 30.41% αυτού. Στην κάτω περιοχή του μεσόφυλλου υπάρχουν ιδιόβλαστα κύτταρα με δέσμες βελονοειδών κρυστάλλων και κρυσταλλοειδή έγκλειστα διαφόρων φαινοτύπων βυθισμένα σε πολυσακχαρίτες. Στους χλωροπλάστες και στο κυτόπλασμα απαντώνται ελαιοσταγονίδια. Οι μεγάλες δεσμίδες τόσο του ξυλώματος όσο και του φλοιώματος συνοδεύονται από σκληρέγχυμα για μηχανική υποστήριξη ή και αμυντικούς λόγους. Όμως, μόνον οι μικρές δεσμίδες του φλοιώματος συνοδεύονται από σκληρέγχυμα, ενώ οι αντίστοιχες του ξυλώματος συνοδεύονται από παρέγχυμα που φέρει έλαια και πολυσακχαριτικής φύσης περιεχόμενο ως μηχανισμό χημικής άμυνας. Τα κύτταρα του δεσμικού κολεού δεν περιέχουν χλωροπλάστες, που αποτελεί ένα χαρακτηριστικό

γνώρισμα των C3 φυτών.

Η U. maritima είναι ένα φυλλοβόλο φυτό με μεσομορφικού τύπου φύλλα και επίπεδο έλασμα. Το φυτό είναι προσαρμοσμένο στο μεσογειακό κλίμα, έχοντας ισόπλευρα φύλλα ώστε να επιτυγχάνεται υψηλός δείκτης φωτοσύνθεσης. Αυτός μπορεί να

οφείλεται στις μικρές αποστάσεις διάχυσης του CO2, στην υψηλή πυκνότητα των βυθισμένων στομάτων και στις δύο πλευρές του φύλλου, στη λεπτή δομή των κυττάρων του μεσoφύλλου, στην κατανομή τω χλωροπλαστών παράλληλα προς το κυτταρικό τοίχωμα που βρίσκεται προς τους μεσοκυτταρικούς χώρους, στους μεγάλους αεροφόρους χώρους στην περιοχή του μεσoφύλλου, καθώς επίσης και στη χαμηλή ειδική μάζα και πυκνότητα ιστού των φύλλων (LTD και SLM).

Ο ανθοφόρος άξονας της U. maritima χαρακτηρίζεται από ένα ενδιαφέρον, μονοδιάστατο σύστημα ανάπτυξης και ένα πρότυπο ταχείας επιμήκυνσης του επάκριου xvii

Sharaf Al-Tardeh Summary

τμήματος. Τα ανοικτά άνθη, πολύ πιθανόν, αποτελούν μια ζώνη διαθέσιμου άνθους κινούμενη συνεχώς προς την κορυφή. Η επιδερμίδα τόσο του ανθήρα όσο και της ωοθήκης φέρει στόματα. Όταν το άνθος μαραίνεται, τα πέταλα κλείνουν ξανά για να προστατεύσουν την αναπτυσσόμενη κάψα του καρπού. Μόλις ο καρπός ωριμάσει, η κάψα διαρρηγνύεται και ανοίγει για να απελευθερώσει τα σπέρματα, τα οποία φέρουν μαύρα πτερύγια.

Η U. maritima έχει ένα τυπικό γυνόπλευρο (διαφραγματικό) νεκτάριο με δευτερογενή παρουσία. Το νεκτάριο αποτελείται από ένα στρώμα επιθηλιακών εκκριτικών κυττάρων, υποστηριζόμενο από 1-4 στρώματα κυττάρων υποαδενικού ιστού και ακολουθούν 2-6 στρώματα παρεγχυματικών κυττάρων. Το νεκτάριο εκκρίνει το νέκταρ σε απόσταση περίπου 2/3 από την κορυφή της ωοθήκης με τη βοήθεια της ραφής του καρπόφυλλου. Ο μηχανισμός έκκρισης του νέκταρος εξαρτάται, κυρίως, από την υδρόλυση των αμυλοκόκκων που είναι αποταμιευμένοι στα πλαστίδια (αμυλοπλάστες) του μέσης ηλικίας νεκταρίου.

Το νεαρό στάδιο του νεκταρίου χαρακτηρίζεται από την καταλυτική παρουσία του ενδοπλασματικού δικτύου στο κυτόπλασμα των κυττάρων του εκκριτικού επιθηλίου. Το κυτόπλασμα είναι ασφυκτικά γεμάτο με ριβοσώματα προσκολλημένα στο ΕΔ. Η ανάπτυξη των αμυλοκόκκων αρχίζει στα κύτταρα του εκκριτικού επιθηλίου, τα οποία ως επί το πλείστον περιέχουν επιμηκυσμένα πλαστίδια. Το ενδιάμεσο στάδιο του νεκταρίου χαρακτηρίζεται από το πυκνό και κοκκιώδες κυτόπλασμα των επιθηλιακών κυττάρων, το ενεργό και πολύ αναπτυγμένο ΕΔ, καθώς επίσης την κυρίαρχη παρουσία των μιτοχονδρίων και των δικτυοσωμάτων. Συνυπάρχουν, ακόμα, ποικιλόσχημα πλαστίδια που περιέχουν οσμιόφιλα κοκκία, ενώ απαντώνται πολυάριθμα εκκριτικά κυστίδια, μεμβρανικοί σχηματισμοί και χυμοτόπια με υλικά υψηλής ηλεκτρονικής πυκνότητας.

Το ώριμο στάδιο αρχίζει 19 ώρες πριν ανοίξουν τα άνθη. Το στάδιο αυτό χαρακτηρίζεται από την παρουσία νέκταρος με κρυσταλλοποιημένη μορφή μέσα στα επιθηλιακά κύτταρα, τον εκφυλισμό των υποαδενικών παρεγχυματικών κυττάρων του παρεγχύματος, την υδρόλυση όλων των αμυλοκόκκων, καθώς επίσης και την πλήρη εξαφάνιση των αμυλοπλαστών και του ΕΔ. Τα επιθηλιακά και τα υποαδενικά κύτταρα υφίστανται σημαντικές αλλαγές. Μερικά επικουρικά κύτταρα εκφυλίζονται και/ή διαφοροποιούνται σε παρεγχυματικά, ενώ τα επιθηλιακά κύτταρα επιμηκύνονται xviii

Sharaf Al-Tardeh Summary

περισσότερο. Υπάρχει, όμως, ακόμα το νεκτάριο, αν και είναι λεπτότερο, πλάτους περίπου 6 κυττάρων από το επιθήλιο έως το τοίχωμα της ωοθήκης. Τα μικροκανάλια είναι περισσότερο εμφανή στο κυτταρικό τοίχωμα των επιθηλιακών κυττάρων.

Η ωοθήκη της U. maritima χωρίζεται σε τρεις κοιλότητες που σχηματίζονται από την ένωση τριών καρπόφυλλων. Οι κοιλότητες περιέχουν αρκετές σπερματικές βλάστες , περίπου τέσσερες για κάθε μία στο ώριμο στάδιο. Μετά τη γονιμοποίηση, όμως, παράγονται περίπου 12 σπέρματα ανά κάψουλα. Το τοίχωμα της ωοθήκης αποτελείται από περίπου 3-10 στρώματα παρεγχυματικών κυττάρων. Το αγωγό σύστημα της ωοθήκης είναι απλό και μοιάζει με αυτό του φύλλου, του βολβού και της ρίζας. Ο αγωγός ιστός, όμως, αποτελείται μόνο από αγγεία του ξυλώματος στο κατώτερο μέρος του νεκταρίου, μόνον από ηθμοστοιχεία και στις δύο πλευρές του νεκταρίου, ενώ απουσιάζει από το ανώτερο επίπεδο της ωοθήκης. Από μια προεξοχή του πλακούντα αναπτύσσεται ένας βλεννώδης αδένας (obturator) που εκκρίνει βλέννα πολυσακχαριτικής φύσης. Ο αδένας αυτός μπορεί να εμπλέκεται στην ετερότροφη ανάπτυξη των γυρεοσωλήνων ή/και στον έλεγχο της κατεύθυνσης της ανάπτυξής τους.

Στους ιστούς της ωοθήκης, του βλεννώδη αδένα και του νεκταρίου είναι εμφανής η παρουσία κυττάρων με πολυσακχαριτικό περιεχόμενο. Δεν υπάρχουν, όμως, λιπόφιλες ουσίες. Επιπλέον, στο τοίχωμα της ωοθήκης και στους υποαδενικούς ιστούς, παρατηρούνται ιδιόβλαστα κύτταρα που περιέχουν δέσμες βελονοειδών κρυστάλλων. Ο υψηλότερος σχετικός όγκος έχει καταγραφεί στο νεαρό στάδιο (7.41 ± 1.8%), μειώνεται, όμως, στο ενδιάμεσο (4.33 ± 1.8%) και αυξάνεται πάλι στο ώριμο στάδιο (6.45 ± 1.55%).

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Sharaf Al-Tardeh Introduction

CHAPTER I INTRODUCTION 1.1 Mediterranean climate 1.1.1 Introduction In the Köppen (1936) classification, a Mediterranean climate is defined as one in which winter rainfall is at least three times the summer rainfall. Indeed, over much of the Mediterranean, summer rainfall is virtually close to zero. This strong summer-winter rainfall contrast is echoed by a pronounced seasonal cycle in almost all climate variables. In particular, climatologists recognize a “Mediterranean climate” with winter rainfalls (between September and April), mild winters and high summer temperatures. The rain is characteristically intense, especially in the drier season and in the drier parts of the region. Rainfall regimes are mainly cyclonic or convectional (Barry and Chorley, 1992).

The climatic characteristics vary greatly over short distances in the Mediterranean, because of the basin and range character of the landscape and because of the impact of the sea on the coastal areas. Inset storms in mid-October, onshore and offshore breezes play an important role in microclimate near the coast (Conte and Giufredda, 1989). Moreover, the Mediterranean mountains play a key role in providing a great diversity of land types. The great basins are normally filled with the erosional products of the mountain-building and are therefore even younger and may be unconsolidated.

Under any circumstances, the vegetation cover plays a key role in land degradation (Francis and Thornes, 1990) and in fact, reduction in the perennial plant cover is regarded as an indicator of the onset of desertification. Rackham (1983) stated that in Boetia (Greece) ‘the landscape and the vegetation appear to have changed less in 2500 years than those of England in the last 1000 years or of New England in the last 180 years’. This is probably becoming less true for many other parts of the Mediterranean, where the abandonment of grazing is leading to an increase in scrubland and hence woodland.

1.1.2 Spatial patterns of temperature in the Mediterranean climate The spatial patterns of temperature and precipitation correspond to the records during the hydrological years 1961-1985. The highest mean temperatures are found in the south and southeast, exceeding 18 oC over it in Libya and Egypt. There is a gradient towards the 1

Sharaf Al-Tardeh Introduction north and northwest, to temperatures below 12 oC. In winter (December, January and February), the lowest mean temperatures in the true Mediterranean region are experienced over Italy, the north coast of the Adriatic Sea and western Greece (below 6 oC). The highest temperatures of course, occur in summer. In this season, the most extensive areas of high temperatures for northern Mediterranean are over Crete and southern Turkey, rising to over 26 oC.

1.1.3 Spatial patterns of precipitation in the Mediterranean climate Precipitation in the region varies from about 1000 mm (it does not exceed 1200 mm per annum) in the more northerly areas and areas above about 800 m altitude, to 250 mm in the southern dry lands. The lowest values are found in southeastern Spain, (between 200 and 400 mm per annum), and over western Turkey and the western shore of the Black Sea, with less than 400 mm.

In winter, precipitation is generally above 60 mm over the Mediterranean islands (except the Balearics) and the land north of the Mediterranean Sea. Spring precipitation is over 40 mm in most areas north of the Mediterranean Sea. The summer season demonstrates a pronounced gradient in the north Mediterranean, from less than 20 mm in southern Spain and Turkey, to over 60 mm over northern Italy. Autumn shows a maximum over the central Mediterranean of over 60 mm and, in isolated areas, over 90 mm. Finally, it is clear from this description that most of the regions conform to the Köppen definition (see section 1.1.1) of a Mediterranean climate (Palutikof et al., 1996).

Sharaf Al-Tardeh Introduction

1.2 Geophytes 1.2.1 Introduction The “geophyte” is a herbaceous plant with an underground storage organ. It is derived from the Greek words, geo = earth, and phyton = plant (Raunkiaer, 1934; Galil, 1981). Geophytes are plant species that survive not only by seed but also by specialized underground storage organs. The primary function of the underground tissue(s) is to store food reserves, nutrients and moisture for seasonal growth and development, thus, ensuring the survival of the species (Raunkiaer, 1934). Since the early seventies of the 20 century, there has been a tremendous and significant expansion in the scientific knowledge related to this diverse group of plants. In addition, worldwide bulb production and utilization have increased three-fold over the past twenty years and the spectrum of bulbs that has been researched and commercially grown has greatly expanded. Lastly, the demand by consumers for knowledge on the utilization of flowering bulbs has increased tremendously.

1.2.2 Taxonomy and origin of geophytes Geophytes are classified using the binomial classification system (Bryan, 1989) into 42 families. But, most genera are included in only three families: Amaryllidaceae, Iridaceae, and Liliaceae. The families in which specific geophytes are classified have been changed from time to time. Thus, some genera have been placed in two or more different families.

The origin of many geophytes has been summarized by Bryan (1989). Many occur in 23o to 45o North and South latitudes (Du Plessis and Duncan, 1989; Rix, 1983; Rees, 1972). There are temperature, altitude, moisture and soil type variations within the specific localities (Du Plessis and Duncan, 1989; Rix, 1983). This range of origins has many climatic conditions (warm/cold; wet/dry), as well as various altitudes. Thus, there is a diversity not only among the storage organs of the geophytes but also winter hardness and summer drought tolerance.

1.2.3 Diversity of geophytes The geophytes genera include both monocotyledonous and dicotyledonous species (Bryan, 1989; Genders, 1973; Herbert, 1970; Liberty Hyde Bailey Hortorium, 1976;

Sharaf Al-Tardeh Introduction

Mathew, 1973, 1978, 1987), and they can be separated into two groups: bulbous and tuberous. Collectively, the ornamental geophytes are commonly called “Flower bulbs” (Halevy, 1990; Rees, 1989). U. maritima is located in the bulbous group in which the scales and leaf bases are the primary storage tissues.

1.2.3.1 Bulbous plants The bulbous plant group consists of true bulbs and corms. Most bulbs and corms are monocots; however, Oxalis cernua (Galil, 1967), a dicot, also possesses a bulb as the storage organ. Morphologically, a bulb has a shortened stem called the basal plate that has one or more apical meristems and is enclosed by several fleshy scales. The basal plate also contains adventitious root initials. The scales are the primary storage tissue in true bulbs. Depending on the species, the scales can be either enlarged leaf bases (Hippeastrum; Rees, 1972) or enlarged scale leaves.

Bulbs can be either tunicated, e.g., Tulipa (de Hertogh and le Nard, 1993) and Narcissus, or non-tunicated, e.g., Fritillaria and Lilium. In addition, some bulbs are replaced annually, e.g., Tulipa, while others are perennial in nature, e.g., Narcissus and U. maritima (Al-Tardeh et al., 2006; Dafni et al., 1981a & b; Bruneton, 1996). Thus, there is great diversity among the true bulb species. Small underground bulbs are called either bulblets or, if they occur at the periphery of the mother bulb, offsets or offset bulblets. Aerial bulblets are called bulbiles. They may occur in either the leaf axils or in the floral parts.

A corm has an enlarged stem (basal plate) that has distinct nodes and internodes. The basal plate is enclosed by several dry, scale-like leaves (tunics) and contains the adventitious root primordial(s) (Hartmann et al., 1990). In contrast to true bulbs, the primary storage organ of the corm is the basal plate. Small corms are called cormlets or cormels. Most corms are monocotyledonous and are reproduced by annual corm replacement; however, Liatris is an example of a dicotyledonous corm.

1.2.3.2 Tuberous plants The tuberous plant group consists of tubers, tuberous roots, rhizomes, and enlarged hypocotyls. Most tuberous plants are dicotyledonous; however, several are monocotyledonous, e.g., Alstroemeria and Convallaria, in this extensive group of 4

Sharaf Al-Tardeh Introduction geophytes. A tuber is comprised primarily of enlarged stem tissues (Harbaugh and Tjia, 1985). It can have one or more apical buds and develops root primordial(s) on the basal part of the tuber, e.g., at the base of the stems produced by the buds as is the case of Gloriosa.

A tuberous root is comprised primarily of enlarged root tissues (De Hertogh, 1989). It has a crown containing one or more apical shoot meristems. Root primordia develop from the distal end of the enlarged roots. Most tuberous roots are dicotyledonous; Hemerocallis, however, is monocotyledonous.

A rhizome is a specialized, horizontally growing stem (Mathew, 1986). In general, both shoots and roots arise at right angles to the horizontal stem, which typically has a nonuniform unjointed appearance. Both monocots and dicots have rhizomes. There are few species that have enlarged hypocotyl tissue as the primary storage organ. This tuberous type is generally called a “Tuber”, but the storage tissues arise from the hypocotyl. Typically, these genera arise from seed-propagated species and they are dicotyledonous, e.g. Begonia Tuberous Hybrids and Cyclamen (Mathew, 1986).

1.2.4 The body of geophytes 1.2.4.1 Storage organs and tissues Scales are the primary storage organs of true bulbs. They are modified leaves bearing stomata. The latter have not been considered in physiological studies such as the major air exchanging apparatuses for respiratory processes, but they should be a contributing factor. The cuticle on scales has also not been determined. Both stomata and cuticle are very important for bulbs such as tulips that require rather high rates of ventilation during dry bulb storage.

Dropper is unusual plant organ defined as hollow diverticulum containing and surrounding a bulb in its entirety, and which, when dry, constitutes the tunic of the bulb. Rees (1972) stated that they are a common juvenile trait of tulips, but they also occur with certain tulip cultivars and in Erythronium as well. In addition to serving as a storage organ, they also help, through developmental processes, to position the bulb at lower soil depths. This serves to protect the bulb against adverse environmental conditions.

Sharaf Al-Tardeh Introduction

1.2.4.2 Aerial organs and tissues Geophytes exhibit a wide diversity of flowers, leaves and stems (Bryan, 1989; Rix, 1983; Rix and Phillips, 1981). The flowers may be single, double, semi-double, or multiflowered. They are leafed and leafless scapes and there are multi-stemmed as well as single stemmed plants. Floral development can be determinate or indeterminate. Many genera in the Amaryllidaceae, Araceae, Hypoxidaceae, Iridaceae and Liliaceae have subterranean ovaries (Rix, 1983). Du Plessis and Duncan (1989) indicate that the appearance of the flowers can be hysteranthous (leaves appear after flowers), proteranthous (leaves appear before flowers), or synanthous (leaves and flowers appear simultaneously). In addition, they classify geophytes as either evergreen or deciduous based on leaf persistence through the year.

1.2.4.3 Roots There are three distinctive root features for geophytes: 1. The contractile root habit. 2. The branching habit. 3. The presence or absence of root hairs. Bulbs generally have one of four types of root systems. They can be performed with fixed root numbers and non-branching. They can be performed with fixed root numbers, but the roots will branch. They can be performed but will form additional basal roots as they grow, e.g. Hippeastrum. Lastly, like most other plant species, they can form totally normal root systems, e.g. Begonia Tuberous Hybrids, Cyclamen, etc.

Many bulbs have contractile roots. However, this feature is not only unique to the ornamental geophytes but also to other plants such as alfalfa, carrots and many perennial herbaceous plants (Esau, 1965). The contractile roots play an analogous role to that of droppers (see section 1.2.4.1). Both assist the plant in positioning the storage organs at the proper depth and in the survival of the species.

Some bulbs have non-branching characteristics (monofilamentous). This is an important feature because if the root is injured, e.g. burned by salts or mechanically damaged, there will be no branching of the remaining roots. Also, tulip roots, unless exposed to ethylene (De Munk and De Rooy, 1971), do not have root hairs. This is an unusual root feature.

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Schuurman (1971) confirmed that the number of basal roots formed per bulb or corm can be fixed and is related to the bulb size. Thus, the larger the basal plate the more primordial roots are formed. Since the number of basal roots is fixed, any injury to the basal plate results in an irreversible loss of roots. This, in turn, can lead to remarkable changes in growth and development of the bulb. For example, if the basal plate of Dutch Iris becomes infected with Fusarium, the plant produces shoots with a pronounced curvature. Therefore, it is critical to always protect the basal plate from both disease and mechanical damage.

1.2.4.4 Tunics The tunic is an interesting bulb tissue surrounding the storage organ and possessing stomata. It is defined as a scale that has totally been modified as the bulb or corm mature. It must be noted that tunics do not necessarily cover the basal plate. For example, in Hyacinthus and Narcissus the basal plates are covered only by scales. The tunic provides some functions such as chemical effects, respiration and protection against disease infection and mechanical damage of the outer scales and/or the basal plate that contains the root initials. However, Kamerbeek (1962) showed that after the tunic was removed from Iris hollandica (Dutch irises), bulb respiration was stimulated. It returned, however, to steady state after 6-7 days. Rodrigues Pereira (1962) found a similar response but reported that the steady rate was double that of the bulbs with tunics. Algera (1947) and Rees (1972) reported a similar tunic effect on tulips. It has been reported (Ruzin, 1979; Rees, 1972; Mansour, 1968) that not only the tunics of Freesia and Tulipa have stomata but also the scales of tulips do. The only exception is being the inner surface of the innermost scale of the tulip. In almost all instances the stomata of these tissues were found to be open. It is obvious that these morphological features affect bulb respiration, but further investigations are needed to determine their precise effects.

1.2.5 Periodicity and dormancy of flowering bulbs 1.2.5.1 Definitions In their native habitat, geophytes are subjected to a wide range of climatic conditions (see section 1.1.1). They are characterized by marked seasonal changes in temperature, rainfall, daily irradiation and photoperiod. An exception to this situation would be the equatorial and subtropical areas, where more or less uniform conditions are present all 7

Sharaf Al-Tardeh Introduction year round at a given altitude. In the latter case, many genera do not show any marked rest period and they continuously possess foliage leaves (evergreens or non-deciduous). Well known examples of these non-deciduous bulbs are Hippeastrum (Hartsema, 1961; Rees, 1972) and Clivia (Du Plessis and Duncan, 1989).

In order to withstand the Mediterranean climate fluctuation, geophytes have undergone dormancy (rest period). During the rest period the various genera show a great diversity of behavior. Kamerbeek et al. (1972) and Amen (1968) considered the ability of the shoot to develop new organs and to elongate as the major criteria to define dormancy. Rees (1974, 1981, 1985), depending on his studies on Tulipa, suggested that periods of meristem inactivity could be regarded as true dormancy. While Le Nard (1983) suggested that the term of dormancy could be applied to the entire bulb and not specifically to the apical bud or meristem. The period of bulb dormancy could, therefore, be defined as being the period during which it is not able to react to environmental factors which induce and/or allow the expression of growth process. This period of dormancy is characterized by the fact that organogenesis is not active and it is also in agreement with Rees’ proposal (1985). In such a physiological state, the bulb reserves are not available for growth of the daughter organs. The period of dormancy mainly corresponds to the period of bulbing (bulb enlargement) (Aoba, 1974a, b; Tsukamoto, 1974; Uemoto et al., 1983; Okubo et al., 1988).

The latter concept of dormancy is based on studies which demonstrate that following their enlargement, bulbs or corms are never physiologically or biochemically at complete rest. They exhibit a continuous physiological evolution even in the absence of visible organogenesis (Le Nard, 1983; Roberts et al., 1985). Finally, dormancy could be defined as a complex and dynamic physiological, morphological, and biochemical state, during which there is no apparent external morphological changes or growth. Internally, however, many physiological and/or morphological events are occurring (Interbulb developmental period).

1.2.5.2 Factors that affect bulb dormancy Temperature is the major factor that affects bulb growth. Moreover, it is commonly used to hasten or delay development. However, other factors can influence the duration of bulb dormancy. The existence of a solid and hard tunic, which probably limits the 8

Sharaf Al-Tardeh Introduction gaseous exchanges, contributes to a longer period of dormancy in Gladiolus cormels

(Apte, 1962). Denny (1938) has stated that the moisture prolongs the rest period in Gladiolus corms. The fact that most bulbs contain fairly high moisture levels and have dry matter contents of about 30%, probably constitutes one of the major factors that prevent true physiological rest during storage.

1.2.6 The flowering process The order in which the different organs of the plants are differentiated is important. When the leaves are differentiated before the floral parts, growth is usually synanthous¸ i.e. the foliage is produced before flowering. This is very a common situation in bulbous and tuberous plants, e. g. Hyacinthus, Lilium, Narcissus and Tulipa. However, in some genera or species the flower bud is initiated before the vegetative bud that produces the leaves. This generally leads to the production of hysteranthous plants, i.e. flowering takes place in the absence of foliage and before it emerges. Hysteranthous geophytes include: Amaryllis belladonna, Gladiolus carmineus, Nerine sarniensis, Pancratium maritimum (Halevy, 1990), Scilla autumnalis, Urginea maritima (Al-Tareh et al., 2008b), and Watsonia hysterantha, as well as many species of Colchicum, Crocus, and Sternbergia, and the majority of Amaryllidaceae of Southern Africa (Du Plessis and Duncan, 1989).

In hysteranthous plants, from flower emergence until anthesis, the photosynthesis rate is very low. Only the flower stem is present, but the bulb reserves are sufficient to allow flower stem elongation and flowering. The contrast to hysteranthous growth is proteranthous growth, i.e., the foliage dies down before the flower is produced, e.g., Boophane haemanthoides (Du Plessis and Duncan, 1989). Thus, all photosynthesis takes place before flowering.

1.2.7 Types of hysteranthous geophytes Most of the geophytes with hysteranthous leaves in the Mediterranean flora are from the Liliales: Liliaceae (U. maritima), Amaryllidaceae and Iridaceae. Some other genera with similar phenological behavior are Primulaceae (Cyclamen) and Araceae (Birarum). Depending on the basis of their storage organs, life cycles, and several other features, two patterns can be distinguished among hysteranthous geophytes; The Urginea type and the Crocus type. 9

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In the hysteranthous geophytes of “Crocus type”, such as Crocus, Merendera, Colchicum, and Sternberhia, the capsula is kept below the ground level during the winter, thus being protected from winter damages (Arber, 1910). Seed dispersal starts in spring, and they go into dormancy of up to one year. The first chance for germination is during the following autumn.

The hysteranthous geophytes of “Urginea type”, such as Scilla autumnalis, S. hanburyi, Urginea maritima (L.) Baker, U. undulata, Pancratium maritimum, and P. parviflora, are characterized by possessing perennial storage organs, a distinct flowering stem, onset of flowering proceeding leaf development in winter, seed dispersal immediately after flowering, germination without dormancy, and tropical origin (Dafni et al., 1981).

1.2.8 Factors affecting bulb growth and development 1.2.8.1 The size of the bulb Like other plants, bulbs must reach a certain physiological stage before they acquire the capacity to flower. This stage, analogous to a “ripeness to flower”, is reached after a “juvenile” period, which can be from less than a year to as long as six years. The juvenile period of Tulipa is 4-7 years, Gladiolus is 1-2 years and U. maritima is almost six years (Al-Tardeh et al, 2008b; McCorohan, 1990).

Bulb size is the major, and easily measured, factor that determines the capacity to flower. The critical size is genus or species dependent. The minimum flowering size for Tulipa and Gladiolus is 6-10, 3-6, cm in circumference, respectively. Moreover, for a given genus or species, the critical size can also varies with cultivars and the environmental conditions (Hartesma, 1961). Finally, the possible effect of root activity on flower induction is another factor. In the case of Tulipa, it appears that the presence of actively growing roots unfavorably affects flower induction in the daughter bulbs (Le Nard, 1985, 1986).

1.2.8.2 Temperature In bulbs that exhibit a marked periodicity, growth and development are mainly affected by seasonal thermoperiodicity. Moreover, in some bulb species, such as Scilla autumnalis and U. maritima, daily thermoperiodic changes are required to obtain flowering (Halevy, 1990). Changing the seasonal thermoperiodism can lead to an 10

Sharaf Al-Tardeh Introduction important modification of growth and development. Halevy (1990) has reported that Ornithogalum thyrsoides, a synanthous-deciduous plant in its native habitat in South Africa, does not produce a normal bulb when grown in tropics. It flowers constantly and is evergreen.

Root differentiation and growth are absolutely necessary to obtain optimal bulb growth and development. It is generally observed that temperatures favorable to flower differentiation are also favorable to root initiation and differentiation (Kawa and De Hertogh, 1992). Moreover, temperatures allowing prolonged growth of the aerial parts are generally favorable to bulbing (Le Nard, 1983).

High temperatures could lead to physiological disorder known as “spitting” in Hyacinthus. The major effect of high temperature may be desiccation of the bulbs and causes abnormalities such as floral abortion or damage to the basal plate of Narcissus (Bergman, 1978).

1.2.8.3 Light The effects of light may be due to photoperiod (e.g., Dahlia, Gladiolus, Lilium) or light intensity. Very low light intensities usually result in flower abortion or blasting in genera such as Freesia, Gladiolus, Iris, and Lilium (Einert and Box, 1967; Hartsema and Luyten, 1955; Mansour, 1968; Wassink, 1965). Generally, the higher the growth temperature, the higher the light intensity required to avoid flower abortion.

1.2.8.4 Soil features For almost all flower bulbs, drainage is the most critical factor for selection of a production field or planting medium (De Haan and Van der Valk, 1971; Wiersum, 1971). Most bulbs need to be kept moist through the growing season, but they do not grow well under poorly drained conditions. Some bulbs, e.g., hyacinths and/or even specific cultivars of tulips, do best when grown on sandy soils, while others prefer a clay soil. Although there are some exceptions, the pH for most bulb soils should be 6-7. Very high or low pH can lead to root burn. For disease control, it is important to rotate the bulb crops with other agronomic crops.

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1.2.9 World production of geophytes Nowadays, much attention and interest have been focused upon geophytes. The world flower bulb production has been concentrated on six genera and their production is centered in three countries: the Netherlands, being the world leader with about 75% of the total world production of geophytes, the United States and the United Kingdom (Gould, 1957; Kiplinger and Langhans, 1967; Moore, 1984; Schenk, 1984; Smith and Danks, 1985; Vijverberg, 1982). The genera produced in excess of 900 hectares are Gladiolus, Hyacinthus, Iris, Lilium, Narcissus, and Tulipa. These six genera represent about 90% of the world’s flower bulb production acreage.

The major uses for flower bulbs are outdoor usage and forcing. The outdoor usage includes the home gardens, parks, arboreta, commercial landscapes, roadsides, cut flowers, resorts, golf courses and containers. The forcing includes cut flowers, potted flowering plants, growing (sprout) plants, home forcing, house plants and interiorscapes.

Sharaf Al-Tardeh Introduction

1.3 Chlorophyll contents and fluorescence The principle underlying chlorophyll fluorescence analysis is relatively straightforward. Light energy absorbed by chlorophyll molecules in a leaf can follow one of three routes: (1) it can be used to drive photosynthesis (photochemistry), (2) excess energy can be dissipated as heat or (3) it can be re-emitted as light-chlorophyll fluorescence. These three processes occur in competition, so that any increase in the efficiency of one will consequently result in a decrease in the yield of the other two. Hence, by measuring the yield of chlorophyll fluorescence, information about changes in the efficiency of photosynthesis (photochemistry) and heat dissipation can be gained. Although the total chlorophyll fluorescence is very low (1 or 2% of total light observed), measurement is quite easy (Maxwell and Johnson, 2000).

The measurement of chlorophyll a fluorescence is elaborated by dark-adapting of leaves of the same age for 30 min by means of leaf clips. Leaf clips are put in the centre of each leaf where the mesophyll chloroplasts were almost of the same age, in order for results to be comparable. Leaf clips were made of white plastic material, which reduces heat absorption by the plant tissue under study. They were also equipped by a particular bolt to prevent any light inlet to the studied tissue. Adaptation of the plant tissue to the dark makes electrons to move away from the reaction centres of photosystem II (PSII), which have already been characterized as open reaction centres. As a result, measurement of the minimum fluorescence value (Fo) (minimum fluorescence level) and of its maximum value (Fm) before and after the first exposure of the tissue to the light is made possible (van Kooten and Snell, 1990).

After opening of the bolt a weak modulated measuring light (6 nmol m-2 s-1 to 660 nm intensity) goes through a hole and allows measurement of Fo. A saturating light pulse (10.000 μmol m-2 s-1 intensity of a 400 nm < λ, 700 nm wavelength for 0.5-2 seconds) follows in order to measure Fm in tissues already adapted to the dark (van Kooten and Snell, 1990). The difference between Fm and Fo is termed as “variable component” (Fv). The Fv/Fm ratio (Genty et al., 1989) is estimated to express the maximum quantum response of PSII during photochemistry, when all centres of PSII reaction are open and in relation to the stable status of fluorescence response to the light (Ft) information is obtained regarding the ability of photochemical quenching and the PSII response (Maxwell and Johnson, 2000). 13

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The Fv/Fm ratio is a sensitive indicator of the photosynthetic ability of plants. It is known that in healthy plants its value is equal to 0.83 in most plant species (Björkman and Demming, 1987; Johnson et al., 1993). Values lower than 0.83 are observed for plants under stress conditions and in association with changes in the value of Fo parameter, which become reliable indices of the photo-inhibition phenomenon (He et al., 1996; Maxwell and Johnson, 2000). In the period following lighting (SP), transport of enzymes from PSII to PSI commences ensures the change of radiant energy to chemical. As a consequence, a significant amount of QA plastoquinone recipients reduces during the first seconds of lighting which results in fluorescence increase. However, as photochemistry and the subsequent energy loss to heat form, fluorescence response decreases reaching a value corresponding to a stable status of fluorescence response (Ft).

Through a continuous actinic light (AL), which activates photosynthetic processes, lighting of the photosynthetic tissue occurs and after some time the tissue is lit with actinic light of different intensities (66-3111 μmol m-2 s-1) with wavelength λ > 700 nm (FR) (Far Red light) for 10 min with saturating light pulse every 20 s. In this way, the minimum fluorescence values (Fo’) with PSII reaction centres open and the maximum fluorescence values (Fm’) with PSII centres closed are assessed for every light intensity applied (van Kooten and Snell, 1990). At this point, reduction of plastoquinone QA recipients (PQ) follows during the first seconds of lighting. The values of parameters Fm’ and Ft are then used in estimating the fluorescence response at each period of time, which is indicative of the photochemical quenching of fluorescence (qP). Parameters Fm and Fm’ are employed for the determination of non-photochemical quenching of fluorescence (qN) which is attributed to energy lost in the form of heat.

Schematically, van Kooten and Snell (1990) measured chlorophyll fluorescence by using the saturated pulse method. Measurement takes place by means of a fluorimeter of weak modulated measuring light. Depending on the light conditions five characteristic curves of distinct fluorescence were applied: modulated measuring light (ML), saturating light pulse (SP), actinic light (AL) which activated photosynthetic processes, wavelength λ > 700 nm light. They assessed fluorescence quenching at a given time by comparing it to a reference status adapted to the dark, which is characterized as qP = 1 and qN = 0 presenting value changes of the above parameters after application of light of specific 14

Sharaf Al-Tardeh Introduction intensity and wavelength. The values of Fo, Fm, Ft (Fs), Fo’ and Fm’ fluorescence indices were measured. Moreover, the photosynthetic parameters ETR (J) (Electron Transport Rate), Yield (ΦpsII), qP (Photochemical quenching), qN (Non-photochemical quenching), l-qp and the ratio Fv/Vm were estimated. These parameters are typical of different points of event sequence of the photosynthetic process. Estimation of these parameters is necessary to predict the photosynthetic ability of plants of different genotypes and plants of stress exercised of the same and of different genotypes. More concretely, the photosynthetic parameters represent:

1. Fv/Fm It express the major yield of PSII photons in photochemistry and the ability of photon yield, when all PSII reaction centres are oxidized (open reaction centres) (Maxwell and Johnson, 2000). The value of the ration Fv/Vm is determined by the equation: Fv/Vm = (Fm-Fo)/Fm which renders information about those processes which have caused changes in the ability of PSII photon yield (Maxwell and Johnson, 2000). Values of the Fv/Vm ratio in tissues adapted to the dark show the possible ability of PSII photon yield and are sensitive indices of the photosynthetic yield of plants subjected to stress conditions (Krause and Weis, 1991; Maxwell and Johnson, 2000).

2. Quantum yield (ΦPSII) It is the most useful parameter since it represents the total quantum yield of PSII according to photochemistry (Genty et al., 1989). It informs about the light analogy absorbed by chlorophyll molecules in relation to PSII which is used in photochemical processes. Moreover, it shows, in a way, the rhythm of electron transport thus constituting an indication of total photosynthesis (Maxwell and Johnson, 2000). It is determined by the following equation: ΦPSII = (Fm’ – Ft)/Fm’

3. ETR (J) (Electron Transport Rate) It represents the apparent rhythm of electron transport through PSII measured in μmol electron m-s s-1. It is determined based on the Yield and PAR parameters by the equation: 15

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ETR = Yield x PAR x 0.5 x 0.84 where the Yield and PAR parameters represent the total photon yield and the flow density of the photosynthetically shedding active radiation measured in μmol photons m-2 s-1, respectively. Factor 0.5 is also estimated due to the fact that absorption of two photons is required for the transport of one electron, since PSII and PSI are involved. Factor 0.84 is taken into account in the above relation, because as it is known 84% of the incident photons are absorbed by the leaf.

4. qP (Photochemical quenching) It constitutes a broadly used parameter of chlorophyll fluorescence, which reflects the amount of radiant energy transformed into chemical in photochemistry (Maxwell and Johnson, 2000). Its values range between 0 and 1 (Krause and Weis, 1991) and are determined by: qP = (Fm’ – Ft)/(Fm’ – Fo’) Despite of its superficial similarity to the Yield parameter, it represents different phenomena of photosynthetic mechanism. More specifically, Yield parameter represents the analogy of absorbed energy used in photochemistry, whereas qP parameter informs about the analogy of the PSII reaction centres which are oxidized (open reaction centres) (Maxwell and Johnson, 2000).

5. qN (Non-photochemical quenching) This parameter reflects the energy released in the form of heat and is determined by: qN = (Fm-Fm’)/(Fm-Fo’) and qN = 1-(Fm’-Fo’)/(Fm-Fo) Its values range between 0 to 1 (Krause and Weis, 1991) and represents the changes in energy loss to heat, which are associated with a reference situation adapted to the dark (Maxwell and Johnson, 2000).

6. 1-qP It is an alternative way of expressing photochemical quenching and gives information about the analogy of the PSII reaction centres (closed reaction centres) (Maxwell and Johnson, 2000). It is determined as excitation pressure of PSII (Maxwell et al., 1994) as follows: 1-qP = 1-(Fm’-Ft)/(Fm’-Fo’)

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1.4 Nectaries and nectar 1.4.1 Introduction The original pollinators were most probably beetles (insects of the order Coleoptera, having biting mouthparts and forewings modified to form horny coverings that protect the underlying pair of membranous hind wings when at rest). The original attractant in insect pollination was the pollen. However, due to the high cost of pollen production, the flowers started to produce a cheaper foodstuff, nectar, as an alternative. For the production of nectar, special structures were formed, the nectaries. They originated independently in the most diverse lines of angiosperms evolution and on a most widely varying morphological basis (Takhtajan, 1980). Later, Peter Endress (1994) stated that the reward(s) to pollinating insects in early angiosperm flower evolution were floral secretions not pollen. Nectaries may have multiple evolutionary origins and can be induced in a wide range of positions and tissues in the flower (Simpson and Neff, 1983).

Earlier, it was assumed that nectaries originated as excretory organs to rid the flower of superfluous liquid (Lorch, 1978). de la Barrera and Nopel (2004) gave a physiological explanation for nectar-secreting structures in the context of carbon and water relations of flowers. According to their “leaky phloem” hypothesis, nectar secretion could result from high hydrostatic pressure in the phloem and the structural weakness of the developing phloem tissue. Their complementary “sugar excretion” hypothesis is based on sugar accumulation due to rapid growth and associated high transpiration rates to be ecological rather than physiological, as sites where liquid substances involved in interactions with animals were produced and offered in exchange for benefits to the plant.

1.4.2 Definitions of nectary Nectaries, from a functional point of view, are plant secreting structures that produce nectar. However, from the anatomical point of view nectaries vary widely in ontogeny, morphology and structure (Fahn, 1979a, 1988; Durkee, 1983; Smets et al., 2000; Nepi at al., 1996a; Küchmeister et al., 1997; Fahn and Shimony, 2001; Pacini et al., 2003). Morphological differences exist between flowers of the same plant and between plants of the same species with different ploidy (Davis et al., 1996), and morphological characters may be affected by environmental conditions such as water availability. Nectary structure in Lamiaceae species in a Mediterranean shrub community is largely shaped by 17

Sharaf Al-Tardeh Introduction phylogenetic and climate constraints (Petanidou et al., 2000). Moreover, the frequency of species with stomatal nectar secretion should be much higher in hot and arid climates like the Mediterranean and deserts than in temperate ones (Petanidou, 2007).

According to Schmid (1988), nectary is more or less localized, multicellular glandular structure that occurs on vegetative or reproductive organs and that regularly secretes nectar, a sweet solution containing mainly sugars and generally serving as a reward for pollinators or for protectors (e.g. ants) against herbivores, or, in carnivorous plants, as a lure for animal prey.

Schmid’s definition implies that the nectary may be an organ, e.g., the rudimentary carpellodia of staminate flowers of Buxus (Schmid, 1988), or commonly only part of an organ and can be applied correctly when the nectary is conspicuous, continuous, and occupies a well-defined area. However, small discontinuous nectar-secreting structures scattered over a large area have been found among floral and extrafloral nectariferous organs of Peperomia (Piperaceae), Cabomba (Cabombaceae), Sarracenia (Sarraceniaceae), Cephalotus (Cephalotaceae), Chimonanthus (Calycanthaceae), Aristolochia (Aristolochiaceae). Vogel (1998) termed such small secreting structures as nectarioles. In such cases it is unclear whether the term nectary refers to the individual nectar-secreting areas or to all of them as a whole. The term nectarium, introduced by Linnaeus (1735) and used also by Davis et al. (1998) for the complex nectary of Brassicaceae, can be used to describe all separated nectaries in a flower, whereas nectary represents the single unit (Bernardello, 2007).

1.4.3 Basic types of floral nectaries The essential topographical classification of nectaries indicates two types: floral (on the flowers) and extrafloral (on the vegetative organs) (Caspary, 1848). These terms were replaced by a functional classification into nuptial (related to the pollination process) and extranuptial (not related to pollination) nectaries, respectively (Elias and Gelband, 1975). This situation creates a terminological inconsistency because some “extrafloral” nectaries are located in the “flower”, i.e. nectaries located abaxially on sepals and petals. To avoid this problem, the more recent classification (Schmid, 1988) favours the use of reproductive (on any reproductive structure from inflorescences, bracts, pedicals, to flowers and fruits) and extrareproductive (strictly on the vegetative organs) nectaries. 18

Sharaf Al-Tardeh Introduction

Despite this, the traditional use of the terms floral and extrafloral nectaries (Elias and Gelband, 1975) is still standard in botanical publications all over the world.

There are a few general nectary classifications that are helpful for floral nectaries (Zimmermann, 1932; Vogel, 1977; Fahn, 1979a; Smets, 1986, 1988; Smets and Cresens, 1988; Smets et al., 2000; Bonnier, 1878; Ewert, 1932; Brown, 1938; Fahn, 1953, 1979a & b, 1982; Daumann, 1970). However, Schmid’s (1988) classification would be the most acceptable one, since it depends strictly on positional criteria and extremely practical investigations, covering most requirements to categorize nectaries. He proposed several additional appropriate terminologies: • Inflorescences (inflorescence nectaries) • Peduncles or pedicles (peduncular or pedicellar nectaries) • Bracts, bracteoles, or involucra (bracteal, bracteolar, or involucral nectaries) • Flowers (floral nectaries) • Ovules in gymnosperms (ovular nectaries) • Fruits (post-floral or fruit nectaries) Among the floral nectaries in the strict sense, Schmid (1988) recognizes nectaries on the following flower parts: • Receptacles (receptacular nectaries) with three types: 1. Extrastaminal nectaries 2. Intrastaminal nectaries 3. Interstaminal nectaries • Hypanthia (hypanthial nectaries) • Tepals (perigonal or tepal nectaries) • Sepals (sepal or calyx nectaries) • Petals (petal or corolla nectaries) • Stamens (staminal or androecial nectaries) with three main possibilities: 1. On filaments (filament nectaries) 2. On anthers (anther nectaries) 3. On staminodes (staminodal nectaries) • Pistils (gynoecial nectaries) with four possibilities: 1. On stigma (stigmatic nectaries) 2. On styles (stylar nectaries)

Sharaf Al-Tardeh Introduction

3. On pistillodes (pistillodal or carpellodial nectaries) 4. On ovaries with two variants: On the outer regions (ovarian nectaries) In the septal regions (septal nectaries) between adjacent carpels that result from incomplete intercarpellary post-genital fusion (Rao, 1975; Schmid, 1985; van Heel, 1988; Simpson, 1993; Rudall, 2002). Smets and Cresens (1988) and Smets et al. (2000) recommended the use of gynopleural instead of sepal, because it is more specific and covers inner, outer and confluent septal nectaries.

1.4.4 Floral and extrafloral nectaries Nectaries are specialized tissues that secrete a sugary solution involved in interactions with animals (Fahn, 1979a; Pacini, et al., 2003). Two types of nectaries, floral and extrafloral, were recognized (Bonnier, 1879). They may differ considerably in anatomical structure, source of nectar components, and mode of presentation (Davis et al., 1988; Pacini et al., 2003). Nevertheless, both present reward for animals and their exudates are chemically similar. However, floral nectaries are better known than extrafloral ones, because they are important sources of food for honeybees and are involved in the reproduction of many plants of economic significance and in the production of many fruit and seed crops.

Floral nectaries are located in different parts of flower such as ovary, stamen, calyx, corolla and receptacle. Their nectar is consumed by insects (Hymenoptera, Diptera, and Lepidoptera), birds (hummingbirds, sunbirds) and mammals (bats, small marsupials) that are vectors for pollen dispersal. Nevertheless, the extrafloral nectaries are common in leaves (petiole, stipule, and blade) and less common in developing inflorescence and developing or mature fruit (Thomas and Dave, 1992). Extrafloral nectaries reward a more limited set of animals, mainly ants that keep herbivores away.

1.4.5 Gynopleural (septal) nectaries 1.4.5.1 Distribution of the gynopleural nectaries A topographical classification of floral nectaries indicates nine different types (Fahn 1979a). Among them, the “ovarial nectary” type includes nectaries that are placed in the septal region between adjacent carpels, the so-called septal nectaries or gynopleural nectaries as they have been more recently defined by Smets and Cresens (1988). They 20

Sharaf Al-Tardeh Introduction result from the incomplete fusion of a small region of the carpel margins, which are otherwise fused (Brown, 1938; Daumann, 1970; Rao, 1975; Schmid, 1985; van Heel, 1988; Smets et al., 2000; Rudall, 2002). The gynopleural nectary, being a cavity inside the ovary, is not directly exposed to nectar-feeding animals and the site of nectar emission is often different from the site of nectar production (Smets et al., 2000).

Gynopleural nectaries are restricted to monocotyledons, where they represent the most common type of floral nectary (Smets et al., 2000). They have been lost several times in monocot evolution due to the development of different pollination systems (in apostasioid orchids, some Tecophilaeceae, some Xanthorrhoeaceae, some Asparagaceae) or the development of perigonal nectaries (in Liliales, some Iridaceae, some Orchidaceae) (Daumann, 1970; Vogel, 1981, 1998; Dressler, 1990; Smets et al., 2000; Rudall, 2002). In addition, gynopleural nectaries are absent in taxa with a gynostemium; a compound structure formed by adnation (The adhesion or cohesion of different floral verticils or sets of organs) of stamens and style (Rudall and Bateman, 2002). Moreover, they are largely absent in dicotyledons, although there are non-secretory septal slits in Saruma (Endress, 1994), Cneorum tricoccum, Koelreuteria paniculata, Ruta bracteosa and a few other dicodyledons (Schmid, 1985).

1.4.5.2 Structure of the gynopleural nectaries Gynopleural nectaries have structural variations that are correlated to (1) the ovary position: superior, semi-inferior, or inferior, (2) the slits position, and (3) the internal structure (Simpson, 1993; Daumann, 1970; Schmid, 1985, 1988; van Heel, 1988; Vogel, 1998; Smets et al., 2000; Rudall, 2002). A very thin and sometimes apparently discontinuous cuticle is present on the surface of the epithelial cells (Fahn and Benouaiche, 1979; Nepi et al., 2006). The nectary cavity is lined by a layer of secretory epithelial cells that may overlie a subsidiary glandular tissue, characterized by smaller cells with denser cytoplasm than the ground parenchyma cells, thus resembling the nectary parenchyma of floral nectaries.

Wall ingrowths are very common in epithelial cells that for this reason are regarded as transform cells. The differentiation of transform cells in gynopleural nectaries is supposed to be an anatomical device to increase nectar output via eccrine secretion (Schmid, 1985). Cell wall ingrowths are highly developed in Aloe and Gasteria (Schnepf 21

Sharaf Al-Tardeh Introduction and Pross, 1976; Nepi et al., 2006), but are not so abundant in nectaries of banana and Tillandsia (Fahn and Benouaiche, 1979; Cecchi-Fiordi and Palandri, 1982), where predominantly granulocrine secretion seems likely. Different extents of the subsidiary tissue were observed in different species of Tillandsia (Cecchi-Fiordi and Palandri, 1982) and were related to nectar production rates.

1.4.5.3 Nectar secretion in the gynopleural nectaries A very complex type of secretion has been reported in several species with gynopleural nectaries. A mixture of proteins and polysaccharides was found in the septal nectaries of banana (Fahn and Benouaiche, 1979). A ring of mucilage canals around the infralocular nectary of some Bromeliaceae was reported (Sajo et al., 2004). Poor nectar production and the presence of amorphous, hydrophilic, acid polysaccharides suggest that the nectariferous tissue may have a role in water and nutrient accumulation in Tillandsia, where nectaries are more developed in species growing in dry habitats (Cecchi-Fiordi and Palandri, 1982).

1.4.5.4 Fate of the gynopleural nectaries The development of gynopleural nectaries follows two patterns that differ mainly in the fate of nectary after the secretion phase: • Breakdown of the nectary epithelium as in male and female Musa paradistiaca flowers (Fahn and Kotler, 1972). The cytoplasm becomes very electron-dense, plastids and mitochondria degenerate, and the vacuole increases gradually in volume until it occupies most of the cell (Fahn and Kotler, 1972). • Transformation of nectary tissue into parenchyma as in Aloe, Gasteria, and Tillandsia (Schnepf and Pross, 1976; Cecchi-Fiordi and Palandri, 1982). It occurs by elongation of the epithelial cells and occlusion of the nectary cavity by acidic polysaccharides.

Schnepf and Pross (1976) demonstrated differentiation of transfer cells in the epithelium of septal nectaries in some Aloe species. A short time before anthesis, they formed an elaborate system of wall protuberances along their outer walls. They redifferentiated in the developing fruit, losing the wall protuberances, increasing in size, and becoming parenchymatous cells. Rearrangement of these

Sharaf Al-Tardeh Introduction

cells was accompanied by transformation of amyloplasts into chloroplasts, probably involved in photosynthesis to help fruit development.

1.4.6 Nectar production and presentation 1.4.6.1 Nectar secretion mechanism Two main types of secretion can be recognized in plants: the holocrine type, in which the process involves cell death at the moment of secretion, and the merocrine type, in which the secreting cells survive and continue their secretory activity. In most cases nectar secretion is merocrine, but in a few cases it is holocrine implying the death of the cells (Elias et al., 1975; Vesprini et al., 1999; Horner et al., 2003; Nepi and Stpiczyńska, 2007).

Phloem sap is considered the “raw” material of nectar. Pre-nectar unloading is favoured by phloem companion cells that often have wall ingrowths of the transfer cell type as observed in Vicia faba, Pisum sativum (Fabaceae) (Davis et al., 1988; Razem and Davis, 1999), and Eccremocarpus scaber (Bignoniaceae) (Belmonte et al., 1994). Pre-nectar passes through the plasmodesmata from the phloem parenchyma cells to the nectary parenchyma cells by symplastic route. Alternatively, pre-nectar flows from the sieve elements and the companion cells via intercellular spaces and cell walls to the secretory cells (apoplastic route). Nectar secretion, i.e. the transport of nectar outside the protoplast of the secretory cells, may occur by two mechanisms (Fahn, 2000): 1. Eccrine secretion is the molecular transport of individual sugar molecules across the cell membrane, possibly by a carrier molecule. 2. Granulocrine secretion is the transport of a sugar solution into vesicles derived from dilated cisternae of the endoplasmic reticulum (ER) or from dictyosomes that fuse with the plasmalemma, releasing nectar into the wall area.

Pre-nectar transport follows the symplastic path in nectaries with trichomes as secretory structures. The apoplastic route of the pre-nectar is impeded by lignification or complete cutinization of the radial walls of the stalk cell, i.e. the second cell of the hair (Fahn, 1979b; Sawidis et al., 1987a; Davis et al., 1988). In Hibiscus rosa-sinensis (Malvaceae) the basal cells of the trichomes (i.e. the first cells of the trichomes), situated at the level of the epidermal cells, have a greater number of plasmodesmata, playing a role in the collection and conveyance of pre-nectar from the nectary parenchyma cells towards the

Sharaf Al-Tardeh Introduction secretory hairs (Sawidis et al., 1987b). Great density of plasmodesmata also occurs in the walls of the stalk cells, which, besides providing a barrier to apoplastic transport, favour symplastic flow of the pre-nectar. After entering the secreting hairs, the pre- nectar flows from cell to cell through the plasmodesmata to reach the tip cell (Sawidis et al., 1987b). Here the nectar accumulates in spaces between the cell membrane and the cuticle prior to its “pulsed” release (Sawidis et al., 1987b).

According to Fahn (1988), hydrolysis of sucrose in the nectary cells maintains a sucrose concentration gradient that could cause a passive flow of sucrose from sieve elements to nectary secreting cells. This model may also explain the preferential flow of the pre- nectar towards the secretory cells rather than the neighbouring cells; it is, however, not applicable to nectaries where sucrose is the dominant sugar secreted.

Starch hydrolysis products may be transformed in the vacuole by specific enzymes, such as invertase. Peng et al. (2004) also demonstrated that pre-nectar transport in C. sativus follows the apoplastic route. ATPase activity in nectary parenchyma cells is required for the transport of pre-nectar from the secretory cells to the intercellular spaces and also for nectar secretion on the surface of the nectary. ATPase activity occurs not only in the plasmalemma of the secreting cells, but also in vesicle membranes in intercellular spaces during nectar secretion.

The presence of invertase activity in the nectar is still debated. It occurs in the nectar of Tilia (Malvaceae) and other species (Baker and Baker, 1983). However, invertase is not found in the nectar from the extrafloral nectaries of R. communis (Nichol and Hall, 1988). Pate et al. (1985) reported that the diluted extrafloral nectar of Vigna unguiculata (Fabaceae) contained inverted sucrose, whereas the undiluted nectar did not. This finding suggests that nectar contains freely soluble invertase, the activity of which is inhibited osmotically at high sugar levels, or that invertase is associated with nectariferous cells and cell debris in nectar, being leached from these materials when nectar is diluted.

1.4.6.2 Nectar presentation • Nectar presentation in floral nectaries The position of floral nectaries with respect to the organ bearing them could be (Fahn, 1979):

Sharaf Al-Tardeh Introduction

1. At surface level 2. Forming an outgrowth, as in Brassicaceae and Fabaceae 3. Sunken, as in the ovary septal nectaries of monocots

Pacini et al. (2003) proposed the new term “nectar presentation” in analogy to the better- known term pollen presentation (Faegri and van der Pijl, 1979), to indicate how and where nectar is offered for consumption. Nectar presentation can be primary, when nectar is offered in the nectary itself and secondary presentation when it is presented elsewhere, e.g., stored in spurs or other reservoirs (Weryszko-Chmielewaska et al. 2006; Nepi et al. 2006).

Spurs are cavities commonly derived from the corolla and are present in at least 15 angiosperm families (Hodges, 1997; Bernardello, 2007). In certain families, such as Scrophulariaceae, spurs are typical of almost all members; in other families, such as Ranunculaceae, they may occur in some members only (the genera Aconitum, Aquilegia, Delphinium, Consolida and Nigella). Spurs may be directed upwards as in Ranunculaceae, or downwards as in Linaria vulgaris P. Mill (Scrophulariaceae) and many orchids.

The “cuculli” of Asclepias (Kevan et al., 1989) are another type of nectar reservoir in which the nectar flows from the nectary by a capillary system. Vogel (1998) describes auxiliary structures, named nectar ducts, the function of which is to conduct nectar from the source towards the site of presentation, as in septal nectaries. Nectar flows along these ducts driven by capillary forces, secretion pressure and gravity, depending on the orientation of the nectar and organs bearing it.

Secondary presentation (spurs, cuculli, septal nectaries) may imply one or more of the following functions (Neiland and Wilcock, 1995):

1. Protection against evaporation 2. Consumption by a limited number of animals with long sucking mouthparts that can reach into the spurs 3. Protection against contamination by fungal spores and bacteria 4. Long exposure (several days), increasing the chances of the flower being visited, especially when pollinators are few as is the case of many orchids

Sharaf Al-Tardeh Introduction

The different ways of nectar emergence and presentation may be related to one or more of the following: 1. Nectary position in the flower 2. Nectary accessibility from outside 3. The path of flower visitors 4. Mouthparts of flower visitors 5. Number of ovules per ovary

• Nectar presentation in extrafloral nectaries

Extrafloral nectar, unlike floral nectar, is always present on the surface of the nectary (primary presentation only). The most common extrafloral nectaries are situated on leaves and stems, rarely on fruit. The secreted nectar is always derived directly from photosynthesis by the nectary or other contiguous tissues, generally without storage of starch. Floral nectar can often be seen as drops by the naked eye, whereas extrafloral nectar, owing to its reduced volume, is not perceived as drops but as a shiny surface. The nectar generally does not flow and is rarely lost. Feedback occurs in some species in which no further nectar is produced, if that present is not collected (Cruden et al., 1983).

1.4.6.3 Fate of nectar and nectaries The fate of the secreted nectar has different fates. It can either:

1. Be consumed by a pollinator 2. Be consumed by a nectar thief 3. Drop from the flower 4. Remain in the nectary or flower if not removed

When the nectary parenchyma has chloroplasts, there are two possibilities at the end of secretion: the nectary either abscisses or persists. Abscission occurs if the nectary is a small inconspicuous protuberance, as in many members of Brassicaceae, Fabaceae, and Asteraceae (Horner et al., 2003). Persistence means that photosynthesis by the nectary parenchyma continues, though products of photosynthesis are shifted to benefit the developing fruit. This call for a rearrangement in the manner of conveying parenchyma photosynthate, i.e. a reorientation of cell polarity and flux must occur in phloem cells.

Sharaf Al-Tardeh Introduction

1.5 The summer flowering geophyte: Urginea maritima (L.) Baker (Liliaceae) 1.5.1 Urginea maritima overview Urginea species belong to the subclass Monocotyledonae and the family Liliaceae, which is about 3700 species in 250 genera (Liberty Hyde Bailey Hortorium, 1976). There are about 40 species that originate in South Africa, the Mediterranean countries and India. The most common species is U. maritima (Mitrakos et al., 1974) and the storage organ is a large bulb.

Urginea is derived from the name Beni Urgin, a native ancient Arab tribe of Annaba (Bellakhdar, 1997). It has several common names such as Mountain Squill, Red Sea Squill, White Squill, Sea Onion, Squill (English); Mountain Slangkop, Red Slangkop, Transvaal Slangkop (Africans); Drimia maritima (L.) Stearn, Scilla maritima L. Urginea scilla Steunh. (English synonymy); Cebolla albarrana, Cebolla de grajo, Ceba marina, Ceborrancha, Escila, Esquila (Castellian Spanish); Beni Urgin, Ein Sit, El-Ansal or Bassal-El-Far (rat’s onion) or Bassal-El-Khanzir (wiled boar’s onion) (Arabic) (Makhlouf, 1978; Bellil, 1983); Skilokrémidha (Greek), Vana-palândam (Sanscrit); Hatzav (Hebrew).

U. maritima is a perennial bulbous geophyte of the family Liliaceae (Bruneton, 1996), native to the Mediterranean basin and well adapted to its type of climate (Polunin and Huxley, 1965; Kopp et al., 1996). It generally occurs in the slopes of hills, in sandy grounds and dry coastal areas of the Mediterranean such as Southern Spain, Portugal, Morocco, Algeria, Corsica, southern France, Italy, Malta, Dalmatia, Greece, Syria, Palestine (Zohary, 1962) and Asia Minor (Battandier, 1893; Gentry et al., 1987), and in certain regions of Northern Africa, (Cuenod et al., 1954; Rogues, 1959; Bellakhdar, 1997) and Middle East.

1.5.2 Morphology U. maritima has two varieties: red and white. The red variety (red squill) is predominant in Tunisia (Makhlouf, 1978; Cuenod et al., 1954), Algeria (Battandier, 1893) and Greece (Al-Tardeh et al., 2006; 2008a,b,c). The white variety is predominant in Morocco (Bellakhdar, 1997).

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The plant can reach 50-150 cm in height. The bulb, which is usually only half buried in the soil, is tunicated and mostly globular. It can reach 5-15 cm in diameter and consists of fleshy, smooth scales (modified leaf bases), which overlap one another. It has a little odor, but its inner scales have a mucilaginous, bitter, acrid taste, owing to the presence of bitter glycosides (Kopp et al., 1996). The leaves are in a rosette form, wide (3-10 cm), long (30-100 cm), long-lanceolated, somewhat undulated, shining and dark-green. They emerge after flower spouting.

Mitrakos et al. (1974) have conducted a detailed study on the flowering process of the plant. They have stated that the inflorescence stalk is a long and dense raceme with more than 250 flowers. The flowers are white, small, numerous and arranged in terminal racemes. Bracts are subulate, often caducous and shorter than pedicels. The pedicels are 10-30 mm long and more or less erect. Perianth-segments are 6-8 mm long, oblong and whitish with a green or purple mid-vein. Anthers are c. 2.5 mm long. Styles are equal to the stamens. The fruits are loculicidal capsules containing numerous flattened seeds (Cuenod et al., 1954).

1.5.3 Growth, development and flowering Mitrakos et al. (1974) have conducted a detailed study on the periodicity of U. maritima. During the autumn, the leaves emerge and attain a length of 30-100 cm. They photosynthesize until April. By May, they are completely dry. According to Scaramuzzi and Bianco (1962), between April and May the leaves develop the necessary conditions for flowering. The bulbs remain dormant from May to late August. In late August, scape emerges and flowers in September. The flowering shoot of U. maritima constitutes an interesting one-dimensional growth. Total elongation of the scape takes only 25 (21-28) days to reach a length of about 140 cm. The maximum rate of elongation is 10 cm per day depending on temperature: the hotter the faster. However, this is an extremely rapid growth. The first floret opens from the bottom flower 15 days after shoot emergence, when it reaches about 80% of its height and the florets continue opening for 19 days.

There have been no reports till now about when the flower initiation begins, but it is complete by May. However, the flowering could be enhanced by applying a 28 °C temperature for one month with more than 48% relative humidity (Scaramuzzi and Bianco, 1962). Flowering can also be hastened by producing a higher than normal 28

Sharaf Al-Tardeh Introduction temperature differential between day and night (McCorohan, 1990). The plant has ornamental value, flowering after almost six years, when the bulb reaches a considerable size (Pascual-Villalobos and Fernandez, 1999).

1.5.4 Nectaries of U. maritima It has been reported by Dafni and Dukas (1986) that the nectar of U. maritima is secreted from three septal nectaries located at two-thirds of the ovary. Its secretion lasts from the opening of the flower at about 1:00 a.m. till about 5:00 a.m., and the maximal quantity per flower is 3 to 7 µl. The nectar drains to the base of the ovary and accumulates there into mucilaginous droplets, one for each nectary. The nectar is initially very dilute, but by mid-day it is almost in a crystallized form. Nectar concentration is affected directly by both temperature and relative humidity, being higher in dry conditions. The maximal nectar volume of about 7 µl comes at 5:00 a.m. and drops to 1 µl at 9:00 a.m. Dry, windy nights could show a maximum accumulated quantity of less than 1 µl.

1.5.5 Pollination in U. maritima as a model of hysteranthous geophytes The flowering season of the hysteranthous geophytes is characterized by poverty of insects (Herrera, 1982) and harsh meteorological conditions. Dafni and Dukas (1986) have described the different devices by which U. maritima faces its unfavourable pollination environment as follows: 1. Insects: The flowers exhibit a typical insect-pollination syndrome by offering abundant exposed nectar as well as pollen. A large gallery of various unrelated insects visit the flowers of U. maritima, but only a few of them can be regarded as potential pollinators: • Apis mellifera • Polistes gallicus • Vespa orientalis 2. Self-pollination 3. Wind pollination: It also occurs and is generally responsible for self- pollination. Development of extra wind-pollination accompanied by partial self-pollination is an adaptation to increase pollination in an unfavorable season when insects are scarce.

Sharaf Al-Tardeh Introduction

The flowers open at about 1:00 a.m. and close the following night at 19:00. Pollen is exposed immediately with the flower’s opening and is quite conspicuous. Every slight vibration of the flowering stem, especially at low humidity, may cause dispersal of pollen by wind. Counting of the number of pollen grains transferred by the wind to the stigmata shows an average of 202 ± 98 grain per stigma. Pollen availability is, thus, considerably reduced with time and by about mid-day the anthers are almost empty. The pollen grains of U. maritima are smooth and non-sticky (Carpenter, 1938) and the plants are usually in dense colonies due to vegetative reproduction. Both factors can promote wind-pollination.

1.5.6 Bulb production and/or propagation Few details in commercial bulb production are available. U. maritima has not been developed commercially, because its rate of asexual multiplication by bulb offsets is extremely low (El Garari and Backhaus, 1987; Van Horn and Domingo, 1950). Genders (1973) indicates that they can be propagated by offsets and seed. While McCrohan (1990) provides information on a twin-scaling system, which requires six years to produce a flowering bulb as follows: 1. 12.5 to 15 cm diameter bulbs are dug out of the field in May or June. 2. Bulbs are cut into 10 to 16 longitudinal sections that must include a portion of the basal plate. 3. The sections are placed into dry soil in a nursery about 5 cm apart and covered with 7.5 to 10 cm of soil throughout the dry southern California summer. 4. By October, small bulblets start to form between the bulb scales of the sections. Usually one bulblet forms per section, with an occasional double. An average is 110 bulblets per 100 sections. 5. By December small leaves, about the size of a small garden chive, appear from the bulblet. 6. After two years in the nursery, 2.5-5 cm bulbs are produced. 7. The bulbs are harvested from the nursery in the summer while dormant and then planted in the filed. Rows (86 cm) with a spacing of 37.5 cm between bulbs are used (300, 000 bulbs per hectare). 8. The first flowers appear about three or four years after being planted in the field.

Sharaf Al-Tardeh Introduction

El Garari and Backhaus (1987) developed a system of in vitro propagation of U. maritima as follows: 1. 6 to 15 cm diameter bulbs (ca. 2 to 3 years old) are dug out of the field in late December or early January and are used immediately. 2. Internal scales are cut from the basal plate, washed and sterilized. 3. The explants (bulb-scales and basal plate) are placed in basal medium*1 in dark for one week. After that, they produce bulblets. Bulblets initiation occurs on 100% of the basal segments and inner bulb- scales. Moreover, bulblet formation occurs only on the inner (adaxial) surface of bulb-scales. It is possible to dissect the bulblets longitudinally into four pieces and each is subcultured in light in order for shoot proliferation, while the bulb- scale could be returned to fresh basal medium regenerating new bulblets. 4. Root initiation*2 is attempted on bulblets arising from bulb-scale explants and on shoots produced from the subculture of longitudinally quartered bulblets within 4 to 6 weeks. 5. Rooted propagules are transferred to pots of vermiculite and placed under the fluorescent lights in the culture room for up to 10 weeks, or 6. They are placed and/or chilled in a 5 oC refrigerator for 3 to 4 weeks prior to transfer to culture room. Plants are irrigated twice a week, once with tap water and once with Hogland’s solution and placed in the greenhouse after 10 weeks.

*1 A medium containing Murashige and Skoog (1962) salts supplemented with a combination of 0.5 or 1.6 µM naphthaleneacetic acid and 0.4 or 1.3 µM 6-benzylaminopurine. *2 Rooting occurs in medium containing 0.5 or 1.6 µM naphthaleneacetic acid.

Sharaf Al-Tardeh Introduction

1.5.7 Horticultural usage Urginea species are not commonly available, but their primary use is as a garden plant. U. maritima can also be used as a cut flower (McCrohan, 1990). In North America, it is used in climatic zones 8 to 11 (De Hertogh et al., 1978). Good drainage is the most important criteria for the suitable soil. The sandier the soil the better. Therefore, clay soils are not recommended. The plant needs a good space apart for future growth and multiplication. The root system is quite extensive, therefore growing in containers is not recommended. The plant prefers full sun or the shade of a deciduous tree. Like most bulbs, a dry obligate dormancy is required. U. maritima seems to be a well adapted ornamental flower.

1.5.8 Biochemical principles and their action From the phytochemical point of view, it has been reported that the major constituents of U. maritima bulbs are glycosides (cardiac glycosides) of the bufadienolide type (Kopp et al., 1996; Krenn et al., 2000; Iizuka et al., 2001). Anthocyanins (Vega et al., 1972), flavonoids (Fernandez et al., 1972), fatty acids, polysaccharides (Spies et al., 1992; Praznik and Spies, 1993) and calcium oxalate (Cogne et al., 2001) are also present.

1.5.8.1 Toxic principle The plant’s toxic principles are several steroid glycosides classified as bufadienolides. The basic chemical structure consists of three components, a sugar moiety, and a steroid nucleus to which an unsaturated 6-membered lactone ring is attached. Scillaren, scillarenin, scilliroside and scillirubroside (Paris and Moyse, 1969) are concentrated especially in the bulbs of the red variety. The quantity of the bufadienolides (4%) varies with the variety of U. maritima (Bellakhdar, 1997).

Scillaren is a mixture of glycosides scillaren A and scillaren B in the proportions of about 2 parts of A to 1 part of B. Scillaren A is scillarenin 3-glucosylrhamnoside. Scillaren B is a water soluble mixture of glycosides remaining after extraction of Scillaren A (The Merck Index, 1996). Scillarenin forms from adaptive enzymic decomposition of proscillaridin A, while scilliroside is scillirosidin 3-glucoside.

On drying, the bulb loses 80% of its weight, its acridity is largely diminished, with slight loss of medicinal activity and increases the glycoside content significantly. A high yield 32

Sharaf Al-Tardeh Introduction of proscillaridin A is obtained from the bulb outer scales after exposure to indirect light for 30 days. Extracts from scales exposed to indirect sunlight are more active in guinea pigs than those obtained directly from the plant (The Pharmaceutical Society of Egypt, 1981).

1.5.8.2 Defense mechanisms The defense mechanisms of U. maritima are screened by the stored compounds of bafadienolides types and caustic saps in the bulbs (Foukaridis et al., 1995; Krenn et al., 2000). The cells of U. maritima contain raphides of calcium oxalate which produce mild inflammation and irritant contact dermatitis when rubbed on the skin (Cogne et al., 2001; Salinas et al., 2001; Makhlouf, 1978) as a major defense and tolerance against herbivores (Ruiz et al., 2002). In addition, Genders (1973) indicates that the juice of the bulbs can cause blisters and that gloves should be worn when handling the bulbs.

Poisoning occurs frequently in autumn; there are marked differences in the susceptibility of animals (Blood and Radostits, 1989; Makhlouf, 1978; Fitzpatrick, 1952; Basson, 1987; Nel et al., 1987; El Bahri et al., 2000) and humans (Touncok et al., 1995) to the U. maritima. Young calves are more susceptible, while goats (Blood and Radostits, 1989) and wild boars (Makhlouf, 1978) least. Signs of poisoning develop within 24 hours of ingestion of large quantities of the bulbs. Scilliroside, the major toxic glycoside, occurs in all plant parts including the leaves, flowers, stalks, scales and especially the roots and the core of the bulbous part (Verbiscar et al., 1986b).

The extract (scillioside) of bulbs shows a strong activity against rodents (Verbiscar et al., 1986a; Fitzpatrick, 1952; Heth et al., 2000; McCrohan, 1990). Meanwhile, the bulb extract is a strong insecticide (Pascual-Villalobos and Robledo, 1999; Pascual-Villalobs, 2002; Pascual-Villalobs and Fernandez, 1999; Civelek and Weintraub, 2004). Rivera and Obon de Castro (1991) mention that U. maritima bulbs have been planted in some cases touching the roots of fruit trees in Spain to avoid ant infestations. The extract gives positive reaction against the microbial agents (Sathiyamoorthy et al., 1999; Hoffmann et al., 1993; March et al., 1991) and fungi (Miyakado et al., 1975).

Sharaf Al-Tardeh Introduction

1.5.8.3 Pharmaceutical advantages of U. maritima Many extraction methods were applied to the plant’s bulb in order to obtain various natural products and to determine their functions (Verbiscar et al., 1986a,b; Pascual- Villalobos and Robledo, 1999; Kopp et al., 1996; Vega et al., 1972; Fernandez et al., 1972; Spies et al., 1992; Cogne et al., 2001).

Liberty Hyde Bailey Hortorium (1976) reports that U. maritima is used medicinally. The cardiac glycosides (scillaren and scillarenin) are used as a cardiotonic diuretic for the treatment of cardiac marasmus and oedema (Mitsuhashi et al., 1994; Harvey and Champe, 1992). The main effects of cardiac glycosides are on the heart, but their extra- cardiac effects on the gastrointestinal tract and the central nervous system are also important. U. maritima bulbs are used traditionally as fish poison in Spain (Arias, 2000).

Urginea sanguinea Schinz has been traditionally used as a blood purifier, abortifacient (Watt and Breyer-Brandwijik, 1962), a treatment for venereal diseases, abdominal pain and backache (Foukaridis et al., 1995). In addition, U. altissima (L.f.) Bak. has been used in African communities against skin problems, bruises, aches and rheumatism (Oliver-Bever, 1986).

Hot water infusions from pounded bulbs of Drimial urginea (D. robusta) Baker are used as enemas and an ingredient of protective mixes. They are also used as expectorants, emetics (vomiting agents), diuretics to treat bladder and uterus diseases, feverish colds (Hutchings et al., 1996), and promote the healing of broken bones (Pohl et al., 2001). Many medicinal bulbous plant species belong to monocotyledons geophytes of South Africa and their traditional relevance in controlling of infectious diseases have been reported (Louw et al., 2002)

Sharaf Al-Tardeh Aim of the thesis

1.6 Aim of the thesis

After reviewing the literature, it has been found that there is little information on U. maritima from the biological point of view (Carpenter, 1937,1938). Mitrakos et al. (1974) investigated the growth pattern of the flowering shoot and Grammatikopoulos et al. (1999) studied the site-dependent differences in transmittance and UV-B-absorbing capacity of isolated leaf epidermis and mesophyll. There are also a few elaborated studies such as caryological and genetical by Pfosser and Speta (2004), phytochemical studies by Kopp et al. (1996), Krenn et al. (2000) and Verbiscar et al. (1986a,b) and pharmacological studies by Pascual-Villalobos and Robledo (1999).

In addition, U. maritima is a consistent component of the Mediterranean vegetation, it is dominant over wide areas and important for homeopathic therapy. Therefore, it has been decided to investigate the morphological and anatomical adaptation of the plant to the Mediterranean climate, thus providing the background knowledge for understanding further experimental work implicating physiological parameters. Moreover, this study aimed at elucidating the plant’s adaptive strategies to the Mediterranean climate, in relation to its photosynthesis efficiency, its efficiency in water and nutrient storing during the long summer drought and its defense mechanisms against herbivores and environmental hazards.

This involved:

1. The anatomical features of the plant organs such as root, bulb, leaf and flower, studied by light microscope (LM) and transmition electron microscope (TEM). 2. The morphological characters of the plant organs such as leaf, root, bulb scales and flower parts, assessed by scanning electron microscope (SEM), morphological parameters and morphometrical studies. 3. Identification of the following components and their role in protecting the plant from environmental hazards and herbivores: • Carbohydrates (polysaccharides) content, identified by periodic acid-Schiff’s reagent (PAS).

Sharaf Al-Tardeh Aim of the thesis

• Lipids and oil content, identified by Sudan Black B reagent. • Total water content, measured by calculation of the moisture. • Raphide crystals content, determined morphometrically. 4. Biomass and resource allocation patterns within the bulb, determined by morphometrical analysis and water contents. 5. Nectary structure and nectar secretion, presentation and fate, explored by LM, TEM, SEM and histochemical analysis (PAS and Sudan Black B).

Sharaf Al-Tardeh Materials and methods

CHAPTER II MATERIALS AND METHODS 2.1 Materials 2.1.1 Plant samples Urginea maritima is classified as following (Mabberley, 1997): Kingdom : Plantae-Plants Subkingdom : Tracheobiota-Vascular plants Superdivision : Spermatophyta-Seed plants Division : Magnoliophyta-Flowering plants Class : Liliopsida-Monocotyledons Subclass : Liliidae Order : Liliales or Asparagales (new monocots taxonomy) Family : Liliaceae –Lily family or Hyacinthaceae (Judd et al., 2002) Genus : Urginea –Liverseed grass Species : U. maritima. Binomial name: Urginea maritima (L) Baker- red squill

Individuals (more than 200 samples) of U. maritima were collected from Souda, a village of about 10 km southeast of Chania, on the island of Crete (the fifth largest island in the Mediterranean), northeast Greece (Fig. 2.1). The meteorological data of the study site (Souda: 35o 33’ N and 24o 07’ E) were collected by the national metrological service of Greece, during the years 1915-1975. The climatological parameters of the study site are listed in Table 2.1. The altitude of the area is about 140 m above the sea level. The area’s climate is that of the Mediterranean type and most of its annual rainfall is distributed in autumn-winter. The annual precipitation is 669.4 mm. The mean annual air temperature is 18.1 oC. January (10.8 oC) and February (10.8 oC) are the coldest months, while July (26.3 oC) and August (26 oC) are the warmest ones (Pennas, 1975).

Sharaf Al-Tardeh Materials and methods

2.1

Fig. 2.1. Map of Crete showing the location of the study site (arrows) from where the plant samples were collected and examined for their adaptation to the Mediterranean climate.

2.1.2 Apparatuses • A Balzers CPD 030 critical-point drier. • A JEE-4X vacuum evaporator. • A JSM 840-A scanning electron microscope (SEM). • A Zeiss 9 S-2 transmission electron microscope (TEM). • An inverted photomicroscope ECLIPSE TE2000-S (Nikon) (LM). • Electronic balance, model: Mark 220. EiB-Italy. • Eppendorf centrifuge, model: 5415, Germany. • Knife maker, model: Leica EM KMR2. • Knife maker, model: LKB. • LKB Ultraspec II spectrophotometer

• MK2 area meter. Delta-T Devices Ltd, Cambridge, UK. • Olympus SZX12 stereo-microscope • pH meter model, Universal Pocket Meter Multiline Pu SET 3, Germany. • Portable fluorometer PAM-2000. Walz, Effeltrich, Germany.

• Reichert OM U2 ultra microtome. • Reichert-Jung Ultracut E microtome. • Rotatory evaporator, WB2001, Heidolph. • TAAB embedding oven, model: E062. • TAAB vari-speed rotator, model: R060. • UV-Visible diode array spectrophotometer 8452, Hewlett-Packard.

Sharaf Al-Tardeh Materials and methods

Table 2.1. A brief summary of the climate of the study site (Souda), showing the monthly and annual mean values of the climatological parameters according to the meteorological station records during the years 1914-1975 (Pennas, 1975).

Monthly and Maximum Minimum The highest The lowest Daily mean Mean Mean values Mean Mean values Mean values Number

annual mean mean mean recorded recorded air duration of duration of of air values of of rainfall of of

values of air values of values of air temp. temp. sunshine in sunshine in relative clouds (scale (in days) precipitation stormy

temp. air temp. air temp. hours hours humidity % from 0-8) days

January 10.8 14.5 7.2 26.0 0.0 3.5 108.0 75 5.5 18.9 156.4 3.5

February 11.1 15.1 7.3 24.8 1.2 4.8 135.6 72 5.2 14.7 114.2 3.1

March 12.7 16.7 8.2 32.4 0.4 5.7 175.8 70 4.7 11.6 75.5 1.9

April 15.7 19.9 10.7 32.2 4.0 7.5 226.2 66 3.9 7.5 39.0 1.0

May 20.1 24.5 14.5 37.2 6.8 10.3 318.9 59 3.0 3.7 13.0 0.6

June 24.3 28.5 17.9 40.0 12.0 11.1 332.4 53 1.8 2.0 6.8 0.4

July 26.3 30.3 20.1 44.5 13.0 11.6 361.0 51 0.8 0.3 0.2 0.1

August 26.0 30.2 20.1 41.4 14.5 11.5 355.7 54 0.6 0.7 0.7 0.1

September 22.9 27.2 17.7 36.2 12.2 9.3 278.2 61 2.0 3.2 16.4 1.3

October 19.0 23.2 14.7 35.0 8.2 6.2 193.0 69 3.9 11.3 92.6 4.0

November 15.9 20.4 11.9 32.8 2.2 5.8 173.3 72 4.0 10.5 59.7 3.3

December 12.8 16.7 9.0 28.6 0.4 4.1 126.7 73 5.2 17.0 94.9 4.1

Year 18.1 22.3 13.3 7.6 2784.8 65 3.4 101.4 669.4 23.4

Sharaf Al-Tardeh Materials and methods

2.1.3 Experimental design The study was carried out from February 2004 till January 2008 at the Aristotle University of Thessaloniki, School of Biology, Department of Botany, Greece. Plant samples were separated according to their age. Individuals of 10 to 15 cm bulb circumference of U. maritima (Fig. 2.2) were selected and proceeded for the experimental examinations. For root investigation, segments of the adventitious roots (2-3 cm from the root tips) were fixed and studied by LM and TEM. Morphometrical and histochemstrical investigations were also applied. The total water contents of the root were calculated in winter and summer in order to express the water storage capacity of the roots (Al-Tardeh et al., 2006).

Fig. 2.2. Photo of the geophyte U. maritima (L) Baker (Liliaceae), growing wild in Crete, Greece.

Individuals of U. maritima with a bulb diameter of 10 cm, growing wild in Souda, were collected at the end of spring of 2005, soon after the seasonal die-back of the leaves. This particular period was chosen for three reasons: (1) to quantify the total nutrient content in

Sharaf Al-Tardeh Materials and methods bulb’s component parts available for reproduction and growth; (2) by that time the plant has recycled the resources from the herbaceous above-ground plant parts for re-usage; and (3) it is before inflorescence initiation that begins in early September. The bulbs were sectioned through their longitudinal median axis to determine their chronological age in order to select plants of the same age. The bulb-scales were ordered from one to eight followed by the tunics, which are bulb-scales that are completely dry (Fig. 2.3). They were dissected from the basal plate, grouped according to their positions and dried at 70 ºC until constant mass (weight). The water content of constituent bulb parts was measured several times during the plant life cycle. Moreover, segments of the ordered bulb-scales, inflorescence stalks and basal plates were fixed (see section 2.2.2. of this chapter) for LM, TEM, SEM, morphometrical and histochemical investigations (Al-Tardeh et al., 2008b).

Segments of leaves (third leaf from the rosette basis) were fixed in order to investigate the leaf features under normal conditions by means of LM, SEM, TEM, morphometry and histochemistry. While some of the plant samples were collected during 2006, they were planted in natural fields into two groups; one was fully exposed to the sun, while the other completely excluded from the sun, in order to explore the differences in the transition zone of the plant leaves (Al-Tardeh et al., 2008b).

In order to investigate the morphology, anatomy and the ultrastructure of the floral nectary (flower glands) in relation to nectar secretion, from its commencement to cessation, three different flower stages were examined: (1) young flowers with the corolla starting to open but not all anthers dehisced; (2) intermediate flowers with all of the anthers dehisced and the corolla completely open; (3) old flowers in which the corolla had started to wilt. The floral parts of the selected flowers were also fixed and examined by LM, TEM, SEM, morphometry and histochemistry. Finally, fresh hand-cut sections of all plant parts were investigated under LM (Al-Tardeh et al., 2008c).

Sharaf Al-Tardeh Materials and methods

2.3

Fig. 2.3. Diagram of a longitudinal section of an U. maritima bulb sampled from a natural population in Souda (Greece). The bulb-scales are ordered from one to eight followed by the tunics and the basal plate.

2.2 Methods 2.2.1 Water content In order to express the water status in the plant tissues under natural environmental conditions, plant parts were dried at 70 ºC until constant mass. Total water content (TWC) was then measured (fresh tissue mass – dry tissue mass / fresh tissue mass x 100%) 42

Sharaf Al-Tardeh Materials and methods according to Cappelletti (1954). This procedure was applied to roots during both the active (winter-spring) and inactive (summer) periods. However, the water status of the bulb-scales was investigated at each stage of development. Biomass was estimated according to the Klapp (1929) and Wacher (1943) method. Their method is based on shoot dry weight proportions as assessed from fresh weight proportions under field conditions.

2.2.2 Fixation The fixation procedure (Table 2.2) was according to the Karnovsky (1965) method. 2.2.2.1 Solution preparations • Buffer stock solution (0.1 M, pH=7-7.2):10.7 g of sodium cacodylate were dissolved into 500 ml of double distilled water. • Buffer solution containing sucrose (pH=7-7.2): 8.56 g of 0.25 M sucrose were dissolved into 100 ml of buffer stock solution while 8 ml of 0.1 N HCl were added. • Fixative solution (pH=7-7.2): 1 g of 2% paraformaldehyde (w:v) was dissolved into 20 ml of distilled water followed by vortex at 60-70 ºC for 10 minutes. Two drops of 1N NaOH were added and subjected to vortex for 5 minutes again. The cocktail was

left to cold at room temperature. After that, two drops of 50 mM CaCl2, 5 ml of 2.5% glutaraldehyde and 25 ml of buffer solution containing sucrose were added. • Washing solutions: a) 50 ml of buffer solution containing sucrose b) 30 ml of (a) and 30 ml of buffer stock solution c) 15 ml of (b) and 15 ml of buffer stock solution d) & e) buffer stock solution.

• Osmium tetroxide 2% OsO4 (w:v): 0.25 g of OsO4 were dissolved into 7 ml of distilled water and were diluted in 7 ml of buffer stock solution (1:1). • Graded ethanol (EtOH) in distilled water (v:v): 30%, 50%, 70%, 90% and 100% • Agar Low Viscosity Resin (LV): the resin media were formulated in order to get blocks of medium hardness and it was consisted of: 48 g of LV Resin, 16 g of VH1 Hardener, 36 g of VH2 Hardener and 2.5 g of LV Accelerator. The mixture was rotated with a magnetic rod for 15 minutes.

Sharaf Al-Tardeh Materials and methods

2.2.2.2 Procedure A: Pre-fixation Small specimens (~9 mm2) of the adventitious roots (Al-Tardeh et al., 2006), bulbs (Al- Tardeh et al., 2008a), leaves (Al-Tardeh et al., 2008b) and floral parts (Al-Tardeh et al., 2008c) were fixed (Table 1) in the fixative solution at room temperature for 3-4 hours.

B: Post-fixation and dehydration After washing of the segments for 15 min in each of the washing solutions, they were postfixed in 2% OsO4 for 2-4 hours, followed by washing in buffer stock solution for 15 min. The specimens were dehydrated in EtOH (v:v) series 30% (15 min), 50% (15 min), 70% (30min), 90% (30min), 100% (1, 2, 2, 1 hours, respectively) and two baths of propylene oxide (15 and 30 min).

C: Infiltration and embedding The Spurr resin (Spurr, 1969) and Agar Low Viscosity Resin were used. The specimens were infiltrated at room temperature as follows: Propylene oxide : Resin 3:1 (v:v) for 2 hours 1:1 for 2 hours 1:3 for 2 hours Pure resin for 12 hours Pure resin for 2 hours Pure resin for 2 hours Embedding in freshly made pure resin for 20 hours at 70 oC.

Sharaf Al-Tardeh Materials and methods

Table 2.2. Chemical treatment of plant tissues for observation in LM and TEM. Treatment Solutions used Time duration Prefixation Fixative solution: 2.5 % glutaraldehyde, 3-4 hours and 2% paraformaldehyde* in 0.05 M cacodylate buffer

Washing Solution a, b, c, d, and e (see section 15 min each 2.2.2.1)

Postfixation Osmium tetroxide 2% OsO4 2-4 hours

Washing Buffer stock solution 15 min up to overnight

Dehydration 30% EtOH 15 min 50% EtOH 15 min 70% EtOH 30 min 90% EtOH 30 min 100% EtOH (4 repeats) 1, 2, 2, 1 hour respectively. Propylene oxide (2 repeats) 15 and 30 min respectively

Infiltration Propylene oxide/LV resin 3:1 2 hours Propylene oxide/LV resin 1:1 2 hours Propylene oxide/LV resin 1:3 2 hours Agar Low Viscosity Resin (LV) 12 hours Agar Low Viscosity Resin (LV) 2 hours Agar Low Viscosity Resin (LV) 2 hours

Embedding Agar Low Viscosity Resin (LV) 18 hours at 70 oC

* 2% of paraformaldehyde was not added into the solution used for the root and the leaf.

2.2.3 Sectioning For light microscopy, semi-thin sections (1-2 µm thick) from resin embedded tissues were obtained with a Reichert OM U2 (ultra) and Reichert-Jung Ultracut E microtome. The sections were obtained by glass knives, which they were cut using an (LKB) knife maker and mounted by immersing them on a drop of distilled water on glass slides. Then the slides were heated at 60 oC for 2 min.

For transition electron microscopy, silver grids were used. The grids were coated with a thin membrane of formvar according to Drummond (1950). For this view, a 0.4 g of formvar was dissolved into 100 ml of chloroform [0.4% of formvar (w:v)]. Clean slides were coated by

Sharaf Al-Tardeh Materials and methods immersing them in the solution using a graduated cylinder. The membrane was cut by a razor blade and laid on the water surface in 45o angle. The membrane was then isolated from the slides and floated over the water surface. The grids were transferred over the floating membrane. A piece of parafilm was laid over the grids and taken out of the water. Ultrathin sections (0.08 µm thick) were obtained and collected over the coated grids.

2.2.4 Staining techniques 2.2.4.1 Staining with toluidine blue O (0.1%) Solution preparation:

1 g of boric acid Na2B4O7.10H2O and 1 g of toluidine blue O (TBO) were dissolved into 100 ml of distilled water by hand shaking. The solution was filtrated with a filter paper Whatman no. 42 mounted on a funnel inserted in the volumetric flask. A dilution of the stain (0.1%) was used.

Procedure: The mounted specimens were stained with 0.1% TBO at 60 oC (Pichett-Heaps, 1969) for a few seconds, then washed with distilled water. They were dried off water by being heated again at 60 oC. Finally, they were photographed using an inverted photomicroscope ECLIPSE TE2000-S (Nikon)

2.2.4.2 Staining with Sudan Black B (1%) Solution preparation: 1 g of Sudan Black B was dissolved into 100 ml of 70% EtOH. The solution was kept in a closed container at 37 ºC for 12h and then filtrated with a filter paper Whatman no. 42 in a funnel inserted in the volumetric flask.

Procedure: To stain lipophilic substances, semi-thin sections (1-2µm) of fixed material and/or hand-cut sections of fresh material were stained with 1% Sudan Black B (Bronner, 1975). Glass slides with semi-thin sections were immersed in 70% EtOH for 1-2 min, and then transferred into the freshly filtered saturated solution of Sudan Black B at 60 ºC in an oven

Sharaf Al-Tardeh Materials and methods for 35 min. The slides were rinsed in 70% EtOH for one minute, and then washed with water (Al-Tardeh et al., 2006 & 2008 a & b).

2.2.4.3 Staining with Periodic acid-Schiff’s reaction (PAS reaction) For the identification of polysaccharide contents, semi-thin sections of fixed material and/or hand-cut sections of fresh materials were exposed to periodic acid-Schiff’s reagent (PAS) according to Nevalainen et al. (1972) and examined with an inverted photomicroscope ECLIPSE TE2000-S (Nikon).

Solution preparations: • Solution A (0.5% Periodic acid): 0.5 g of periodic acid was dissolved into 100 ml of distilled water using a volumetric flask.

• Solution B (Periodic-Schiff’s reagent): 0.5 g of pararosallin was dissolved into 15 ml of 1 N hydrochloric acid (HCl) and stirred slightly until the solution became brown to brown-green. This solution was combined with a solution made

by dissolving 0.5 g of Kalivmpyrosulfit (K2S2O5) into 85 ml of d.H2O. The resulted solution was kept in a refrigerator for 24 hours and was stirred from time to time. 0.5 g of active carbon was added to the yellowish solution which was stirred well for 2 min. The cocktail was filtrated with a filter paper Whatman no. 42 mounted on a funnel inserted in the volumetric flask. The Periodic-Schiff’s reagent is colorless or light yellowish and must be kept in the refrigerator.

• Solution C (SO2-water): this solution was prepared by mixing 200 ml of tap

water, 10 ml of 1N HCl and 100 ml of 10% K2S2O5 solution. This solution was

prepared just before applying the test and was used only if it had a strong SO2 smell.

Procedure: Sections of fixed material and/or hand-cut fresh material were transferred into the solution A at room temperature for 5 min. After sufficient washing with distilled water, they were deposited in Periodic-Schiff’s reagent (solution B) for 30-60 min. The sections were

Sharaf Al-Tardeh Materials and methods

washed, respectively, in three vials full of SO2-water, followed by washing using other three vials full of distilled water. With this stain, polysaccharide contents appeared red with different degrees depending on their concentration. The positively reacted-sections were photographed using an inverted microscope ECLIPSE TE2000-S (Nikon) photomicroscope.

2.2.4.4 Staining with uranyl acetate and lead citrate Grids containing sections as mentioned in section 2.3 were used. Solution preparations: • 4% uranyl acetate: 4 g of uranyl acetate were dissolved into 100 ml of 50% EtOH.

• 2% lead citrate: 1 g of lead citrate (C6H5O7)2Pb3.3H2O was dissolved into 30 ml of double distilled water by shaking for 20 min. After that, 8 ml of 1 N NaOH were added to the solution and the volume was made up to 50 ml using a volumetric flask. The cocktail was shaken for 30 min till it appeared transparent.

Procedure: Grids containing sections were inserted gently, under the stereoscope, in a plastic sheet containing groves for grids. The plastic sheet was immersed in uranyl acetate for 20-40 min. After washing with 50% EtOH for 5 min, the sheet was immersed again in lead citrate solution for 20-40 min in dark (Reynolds, 1963). The grids were washed well with double distilled water and were dried off by means of filter paper. The sections were examined by using a Zeiss 9 S-2 (TEM). For photographing, films of Agfa-Gevaert Scientia 23 D5AH of 7x7 cm size were used.

2.2.5 Scanning Electron Microscopy (SEM) To study the morphology of the U. maritima (tunics, bulb-scales, leaf and flower), a SEM was used. The fixation method for SEM has been described in section 2.2.2 and Table 2.2 with some modifications. After postfixation in osmium tetroxide, the segments were dehydrated in graded acetone series, i.e. 30% (15 min), 50% (15 min), 70% (30 min), 90% (30 min) and four repeats of 100% (1, 2, 2, 1 hours, respectively), while the specimens were critical-point dried in a Balzers CPD 030 device. They were coated with carbon in a JEE-4X

Sharaf Al-Tardeh Materials and methods vacuum evaporator. Observations were made with a JSM 840-A scanning electron microscope (Weryszko-Chmielewaska et al., 2006; Al-Tardeh et al., 2008a).

2.2.6 Maceration Maceration method is depended on the hardness of the epidermis of the plant tissue as follows:

2.2.6.1 Maceration of soft material This method was applied to the leaf of U. maritima.

Solution preparations: Basic fuchsin 1% (w:v): 1g of basic fuchsin was dissolved into 100 ml of boiled water. Fuchsin was added before the water was boiled. After cooling at room temperature, 10 g of solid NaOH were dissolved into the solution.

Procedure: Small pieces (2 cm2) of the leaves were dehydrated in 80% EtOH for several days and then they were rehydrated. After rehydration, they were placed in a solution of 1% basic fuchsin and 10% NaOH at 60 ºC for 10-14h. The specimens were washed for 12 h in frequently changed water followed by dehydration through graded EtOH series for 12h and they were kept into absolute EtOH for 1-2h. They were then placed into a mixture of EtOH and HCl (3:1). At this step, the lignified tissues appeared dark green and other tissues bleached, if they were left in the solution for 1-15 min. The samples were washed again with absolute EtOH for 24h; EtOH was changed several times. Finally, the samples were passed through two xylene baths till they were clear. They were mounted to glass slides and photographed using an inverted microscope ECLIPSE TE2000-S.

Sharaf Al-Tardeh Materials and methods

2.2.6.2 Maceration of hard material This method was used for the adventitious roots of U. maritima.

Procedure: Small pieces (2 cm2) of the adventitious roots of U. maritima were dehydrated into 95% EtOH for several days. For staining, the pieces were placed into basic fuchsin 1% at 60 oC for 24h. After that, the samples were washed in water till completely rehydrated. The hydrated samples were kept in 15% NaOH at 60 oC till they were completely clear. The 15% EtOH was changed every 24h for 3-5 days. The following steps used for soft materials (leaves) as the ones mentioned before (see section 2.2.6.1).

2.2.7 Leaf chlorophyll assay To determine the leaf chlorophyll a and b contents, 1 cm2 of fresh leaf material was homogenized with liquid nitrogen, extracted with 90% acetone, kept at minus 10 ºC for 24h and then centrifuged at 10 000 g for 15 min. The absorbance of the supernatant was measured at 664 and 647 nm with an LKB Ultraspec II spectrophotometer. Chlorophyll a and b content was calculated using the coefficient given by Jeffry and Humphry (1975): -1 Chla (µg ml ) = 11.93 x OD664 – 1.93 x OD647 -1 Chlb (µg ml ) = 20.36 x OD647 – 5.5 x OD 664 Where OD is the optical density This procedure was applied to plant leaves under natural conditions at stages of three and six months after leaf emergence. Moreover, chlorophyll assay was done for plants grown in habitats fully exposed to the sun and plants grown in the shade.

2.2.8 Chlorophyll a fluorescence Two groups of the plant U. maritima were grown up from leaf emergence till just before their senescence (approximately six months). Group one was fully exposed to the sun (light leaf), while group two was fully exposed to the shade (shade leaf). Chlorophyll a fluorescence was then measured after three and six months after leaf emergence. Chlorophyll a fluorescence was measured with a portable fluorometer PAM-2000 (Walz, Effeltrich, Germany). The fluorometer was connected to a portable computer which assessed

Sharaf Al-Tardeh Materials and methods by special software (DA-2000, Walz). All measurements were carried out at room temperature.

More concretely, the photosynthetic parameters represent: 1. Fv/Fm The value of the ration Fv/Vm is determined by the equation: Fv/Vm = (Fm-Fo)/Fm

2. Quantum yield (ΦPSII) is determined by the following equation: ΦPSII = (Fm’ – Ft)/Fm’

3. ETR (J) (Electron Transport Rate) is determined based on the Yield and PAR parameters by the equation: ETR = Yield x PAR x 0.5 x 0.84 where the quantum yield and PAR parameters represent the total photon yield and the flow density of the photosynthetically shedding active radiation measured in μmol photons m-2 s-1, respectively.

4. qP (Photochemical quenching) is determined by: qP = (Fm’ – Ft)/(Fm’ – Fo’)

5. qN (Non-photochemical quenching) is determined by: qN = (Fm-Fm’)/(Fm-Fo’) and qN = 1-(Fm’-Fo’)/(Fm-Fo)

Sharaf Al-Tardeh Materials and methods

2.2.9 Morphometry For the morphometric evaluation of the relative volume of the histological components of the adventitious roots (Al-Tardeh et al., 2006), leaves (Al-Tardeh et al., 2008b), bulb- scales (Al-Tardeh et al., 2008a), and flowers (Al-Tardeh et al., 2008c), a transparent sheet bearing a square lattice of points arrays, 10 mm apart, was laid over light micrographs. The point-counting technique analysis was then applied (Steer, 1981). To determine the density of stomata, stereoscopic photographs of the adaxial and abaxial leaf surfaces were used.

2.2.10 Mathematical analysis of morphological and anatomical features of the plant tissues The following procedures were applied to the U. maritima leaves (Al-Tardeh et al., 2008b). Leaf samples (n=10) of almost the same age and size were investigated under natural conditions. The leaf surface area (SA) was measured using the MK2 (Image Analysis System) area meter (Delta_T Devices Ltd, Cambridge, UK) connected to a TC7000 Series Camera (Burle Industries Inc. Lancaster, PA, USA). Leaf thickness was measured with an electronic digital caliper. Leaf dry mass (DM, mg) was determined by drying at 70 oC to constant weight. Leaf water content was measured (LWC). Specific leaf mass (SLM, mg cm-2) was calculated as the ratio of leaf dry weight to unifacial leaf area. The specific leaf area (cm2 g-1) was determined as the leaf area per unit leaf mass (Reich et al., 1992). Leaf tissue density (LTD, mg cm-3) was calculated as the ratio of DM to leaf volume (V, leaf area x leaf thickness cm-3) (Witkowski and Lamont, 1991), in order to express leaf compactness (Christodoulakis and Mitrakos, 1987). The total leaf thickness and the thickness of the histological components, viz. of the adaxial and abaxial cuticles, the adaxial and abaxial epidermises and the mesophyll were measured from 10 light micrographs of leaf cross- sections (x50).

2.3 Statistical analysis All statistical tests were performed using the statistical software package SPSS for Windows (11.5.1, SPSS Inc., USA). Differences in means of anatomical and morphological variables were assessed by means of the analysis of variance (ANOVA). Correlation analysis was used to examine relationships among plant tissue variables.

Sharaf Al-Tardeh Results

CHAPTER III RESULTS

3.1 ANATOMICAL STUDIES ON THE ADVENTITIOUS ROOTS Individual plants of U. maritima with a bulb diameter of 10 cm, growing wild in Souda (Crete, Greece), were collected and used as experimental material. Segments of the adventitious roots at the region 3 cm above the root tip were fixed for microscopical studies (LM and TEM). Some roots were used for fresh hand-cut sections. All of the rest roots of the selected plants were dried to their constant mass in order to calculate their total water content and their biomass.

3.1.1 Root morphology The root system of U. maritima is an adventitious root system in which most or all the roots develop adventitiously from the underground short stem called basal plate. A bulb of 10 cm diameter possesses 5-10 adventitous roots (Fig. 1). The adventituos roots usually have five branches (Fig. 2). The roots of U. maritima are approximately 30 cm long (Fig. 2) and approximately 1 cm thick (Fig. 3). They have unlimited growth downwards, while the lower part breaks down. The root tips are white to red in colour, while, their upper part turns to dark brown. The above-ground structures (leaves) are completely dry by June, but the roots and the bulb survive the dry summer (dormancy). The average of total water contents of the root during the photosynthetic period (autumn-spring) and dormancy period (summer) are 91.2% and 87.27%, respectively. There is convincing evidence of an intense shrinkage of the velamen in the roots of the dormant period (summer) which is clearly evident in comparison to those of the photosynthetic period (autumn-spring) (Figs 4 & 5).

3.1.2 Root anatomy The adventitious roots of U. maritima are characterized by the presence of a multiple- layered velamen, an epidermal system of 2 to 4 cells wide, and a dimorphic hypodermis with regularly alternating long and short cells (Fig. 3). The velamen epidermis is usually uniseriate, devoid of cuticle, thick-walled and sometimes bears root hairs (Fig. 4). The cells

Sharaf Al-Tardeh Results of the velamen are dead, and therefore may become saturated with water for storage purpose (Fig. 5). The epidermal cells contain different myelin-like structures and other peculiar membranous configurations (Figs 6-8). Putative bacteria are occasionally present within the vacuoles of the epidermal cells (Fig. 9). Electron-dense remnants of putative dead fungi and/or fungal pelotons appear around the outer surface of the epidermal cells (Figs 10 & 11). Electron-dense materials of various shapes are also present inside the intercellular spaces (Fig. 12). All of the exodermal (hypodermal) cells, are usually thin-walled almost elongated and tend to be anticlinally orientated. Many of these cells are frequently shorter (anticlinally) and wider than those of the velamen with a didtinct nucleus (Fig. 3).

The cortex region of the adventitious roots of U. maritima is approximately 20 cells wide in a transverse section (Fig. 3). The cortical cells size decreases gradually from the area close to the velamen towards the endodermis (Fig. 3). The cortical cells are large, globular to oval in shape and thin-walled reflecting a water-storing character (Fig. 13). Thick–walled idioblastic cells containing bundles of raphides (profiles) in the central vacuole are present among ordinary cortical cells (Fig. 14). Bundles of raphide occur within the central vacuole. Under the electron microscope (TEM), the vacuolar content appears foamy and each crystal needle seems to be embedded in a translucent homogeneous substance (Fig. 15). Observations of hand-cut sections under the polarized light reveal open bundles of calcium oxalate needles with an average length of 72 ± 22 µm. These cortical cells which contain bundles of raphides are vertically oriented and located in subtended rows around the central cylinder (Figs 16 & 17).

Idioblastic cells conataining bundles of raphides are observed at the active zone of the root during the active phase (autumn-spring) of the plant life. The development starts by building up the central vacuoles (Fig. 18), which contain immersed raphide crystals and are connected to the tonoplast (Fig. 19). The growing up of the idioblastic cells could be interpreted by the fact that the content of the homogenous substances stored in the intercellular spaces (Fig. 20) is translocated into the central vacuole through the cell walls. The homogeneous substance accumulates around the bundles of raphides forming a sheath- like structure (Figs 13-17). This is strongly evident because the intercellular spaces become

Sharaf Al-Tardeh Results devoid of their content during the next developmental stage (inactive stage). Moreover, the homogeneous substance of the intercellular spaces could also be used in building up the cell walls and/or blocking the present plasmodesmata.

It is also worthy to note that many organelles such as endoplasmic reticulum, the dictyosomes, and mitochondria are accumulate close to- or around the intercellular spaces at the active stage (Fig. 20). This aggregation of organelles and the presence of plasmodesmata indicate that they possibly have a role in the development of the idoblastic cells (Figs. 20). The cortical cells often contain myelin-like structures, irregularly shaped nuclei (Fig. 21), and straight arrays endoplasmic reticulum (Fig. 22).

Roots of U. maritima possess auniseriate endodermis consisting of oval shaped and periclinally oriented cells facing the central cylinder (Fig. 23). The endodermal cell wall does not greatly differ from that of the neighboring tissues (Fig. 24). The pericycle is also uniseriate possessing circular to oval, O-thickened elongated cells (Fig. 23), but can be distinguished with difficulty (Fig. 23). In young roots, the pericycle is composed of parenchyma cells with primary walls, but as the root ages, the pericycle cells may develop secondary walls.

The vascular cylinder of the root of U. maritima is typical of monocots. The xylem is ordinarily 10-arched (Figs 13 & 23). It consists of vessel members arranged in short radial rows, alternating with circular, oval and elliptical clusters of phloem (sieve element) cells (Fig. 25). Vascular tissues are embedded in thin-walled parenchyma cells. The vessel members are typical of those of angiosperms which are obviously seen after maceration (Figs 26 & 27). The vessel members have pitted walls and perforated plates. The protoxylem cells (the first formed xylem differentiating from the procambium) have their secondary walls deposited in the form of rings (annular) or spirals (helical). During the active phase of the plant life cycle (autumn-spring), parenchyma cells in the central cylinder possess nuclei of different shapes (Figs 25, 28 & 29). Moreover, many organelles, such as mitochondria, dictyosomes, endoplasmic reticulum etc., aggregate close to the intercellular space, which is filled with a homogeneous substance.

Sharaf Al-Tardeh Results

The parenchyma cells of the central cylinder possess a few plasmodesmata showing a bulge on their inner side, especially at the stage of leaf emergence and inflorescence stalk (Fig. 30). Loss of water during the summer time results in the shrinkage of these parenchyma cells that have a water-storage character (Fig. 31). In addition, a peculiar, long uniseriate endoplasmic reticulum and myelin-like structures also occur inside the parenchyma cells of the central cylinder (Figs. 32 & 33). Moreover, the pith of the central cylinder is sclerenchymatous with circular to oval and thick-walled cells (Fig. 23).

3.1.3 Root histochemistry In order to investigate the presence of polysaccharide contents in the adventitious roots of U. maritima, Schiff’s reagent was applied to transverse sections. The presence of cells with polysaccharidic content (red-stained cells) is clearly evident in the cortex region of the adventitious roots (Fig. 34). The idioblastic cells, that contain bundles of raphides, also react positively with this stain (small micrograph in the corner of Fig. 34).

In order to investigate the presence of lipid contents in the adventitious root of U. maritima, Sudan Black B technique was applied to transverse sections of the plant roots. When semi- thin or hand-cut sections were treated with Sudan Black B numerous cortical parenchyma cells appeared intensely stained brown to black (Fig. 35). Cortical cells with lipids occur scatterely and they do not differ in size from the ordinary cortical cells. The mucilage cells positively react with Sudan black and negatively with the Schiff’s reagent. The mucilage could be the bitter acrid glycosides.

3.1.4 Root morphometry In order to investigate quantitatively the histological components of the adventitious roots of U. maritima, a morphometric study was applied to the cortex region (Tables 3.1 & 3.2). The relative volume of the idioblastic cells containing bundles of raphides is the highest (19.83%), that of the cells containing polysaccharides the lowest (11.27%), while the corresponding one of the cells containing lipids is intermediate (14.38%). The cortical storage cells occupy in total 45.48% of the root cortex volume.

Sharaf Al-Tardeh Results

Table 3.1. Relative volume (%) of the histological components of the root cortex. (± SD, n= 10).

Histological component of the root cortex Relative volume%

Parenchyma cells 43.99 ± 3.30

Idioblastic cells containing bundles of raphide 19.83 ± 1.10

Cells containing lipids 14.38 ± 0.71

Cells containing polysaccharides 11.27 ± 1.62

Intercellular spaces 10.53 ± 1.28

The cortical cells are mostly oval to slendercal, with an average diameter of 52.32 µm (Table 3.2) and an average volume of 73143µm3. Measurements revealed that the cortex is the thickest tissue (741.3 µm) being almost twice as thick as the central cylinder (318.58 µm). The thicknesses sum of the root partial tissues results in a total thickness of 1976.56 µm (0.198 cm) of the U. maritima adventitious roots at the region 3 cm above the root tip. Consequently, dividing the cortex thickness by the root radius and multiplying by 100% it is concluded that the cortex contributes to the total root thickness by 75% and the central cylinder by 16.12%. Correspondingly, the cortical storage cells occupy approximately 34.11% of the total root volume. The thickest wall is the nacreous wall of the vessel members (2.893 µm), whereas the thinnest one is that of the endodermal cells (0.5014 µm).

Table 3.2. Average thickness (µm) of the histological components of the adventitious root (cross sections) at the region 3 cm above the root tip (Mean ± SD, n = 10).

Parameters Mean ± Standard deviation (µm)

Thickness of velamen 69.25 ± 9.23 Thickness of cortex region 741.30 ± 51.34 Diameter of central cylinder 318.58 ± 5.26 Diameter of an endodermal cell 18.44 ± 1.06 Diameter of a cortical cell 52.32 ± 1.71 Periclinal wall thickness of epidermal cells 0.86 ± 0.30 Wall thicknessmof cortical cell 1.41 ± 0.69 Wall thickness of endodermal cells 0.50 ± 0.16 Wall thickness of nacreous vessel members 2.89 ± 0.74

Sharaf Al-Tardeh Results

3.2 BIOMASS AND RESERVE ALLOCATION PATTERNS WITHIN THE BULB. 3.2.1 Bulb morphology Bulb is a rounded underground storage organ that contains the shoot of a new plant. The bulb consists of a short stem surrounded by fleshy scales (modified leaves) that store nutritive products for the new plant (Fig. 1). Bulb size correlates positively with bulb age and the number of bulb-scales, i.e., the older the bulb the bigger in size and the more bulb- scales. Thickens of bulb-scales varies according to their age and order within the bulb. However, the bulb-scales of the median axis are the thickest and the marginals are the thinnest ones (Fig. 2.3). Moreover, the bulb is covered by multiple-layered tunics, a physical barrier for plant protection from the environmental hazards. Tunics contain stomata which are clearlly seen under SEM (Fig. 36).

3.2.2 Bulb anatomy The bulb-scales are covered with a smooth cuticle of 12 ± 4 µm thick (Figs 39 & 40). The epidermis possesses anomocytic stomata (Fig. 37). The bulb scales possess different types of cells of storage character (Fig. 38). The epidermal cells are very similar, compactly arranged, global to oval shaped and exhibit rounded margins. The epidermal cell walls are evenly thickened. Nevertheless, there is no difference in the wall thickness of the outer and inner epidermal cell walls. Moreover, the abaxial (lower) epidermis is slightly thicker (periclinal) than the adaxial (upper) epidermis (Figs 39 & 40). Under electron microscope (TEM), the epidermal cells are more discernible. The periclinal epidermal cell walls are thicker than the anticlinal ones (Figs 41 & 42). The anticlinal epidermal cell walls possess plasmodesmata (Fig. 43). Dictyosomes and other organelles are also present in the epidermal cells (Fig. 44).

Bulb-scales in cross section show various cells of storage character in its ground tissues, e.g., idioblastic cells containing bundles of raphides, cells containing oils, other exhibiting lipids, some possessing mucilage and cells of water-storage character (Fig. 38). The intercellular spaces of the ground tissues are mostly triangular (Fig. 38). The ground tissue of the bulb-scales is composed of larger cells that are thin-walled, empty, irregularly shaped reflecting a water-storing character (Figs 38 & 45). Thick-walled assimilatory idioblastic

Sharaf Al-Tardeh Results cells containing bundles of raphides (profiles) within the central vacuoles are present among ordinary ground tissue cells (Figs 39 & 45). Under the TEM, the vacuolar content appears to be foamy and each crystal needle is embedded in a translucent homogeneous substance (Figs 46 & 47). The cells containing raphides are circular, oval, polygonal, irregularly shaped and 2-6 times longer than the neighboring ones as viewed in longitudinal section (Fig. 48), while in cross section they are more or less isodiametric (Figs 38, 39 & 45). Moreover, observations of hand-cut sections under polarized light reveal open bundles of calcium oxalate needles 432 ± 54 µm long (n=250) (Fig. 49).

The ground tissue of the bulb-scales possesses circular to oval assimilatory mucilage cells which are bitter and acrid in taste, owing to the presence of bitter glycosides (Figs 38, 39, 45 & 50). Abundant mucilage cells are observed close to the upper and lower epidermis. The presence of plasmodesmata between the cells of the ground tissues, particularly the oil cells, is evident (Fig. 51). Osmiophilic droplets also occur inside the mesophyll cells close to their cell walls (Fig. 52). The presence of mitochondria and straight endoplasmic reticulum (Fig. 53) is conspicuous in the cells of the bulb-scales in the central region.

The vascular bundles are collateral, scattered and numerous (Figs 38 & 39). Both xylem and phloem of large vascular bundles are associated with sclenechyma fibers. However, in the smaller bundles, only the phloem is associated with sclerenchyma fibers, while the xylem is subtended by parenchyma. The phloem sieve elements are supported by the companion cells in order to maintain the osmotic pressure of the cell during transportation of the photosynthetic products (Fig. 54). The metaphloem sieve elements (SE) become thicker, harder and are almost devoid of contents (Fig. 55). The presence of sieve pores between the phloem sieve elements is evident (Fig. 56). Sive pores are provided with callose to control the movements of the substances (Fig. 56). Plastids of the metaphloem sieve elements (SE) are characterized by peculiar protein crystalloid inclusions (Fig. 57). During the active phase, the parenchyma cells of the vascular bundle possess abundant organelles such as mitochondria, Golgi body and straight cisternae of endoplasmic reticulum (Figs 58 &59).

Sharaf Al-Tardeh Results

Morphologically, the bulb has a shortened stem called the basal plate that has one or more apical meristems and is ensheathed by several fleshy bulb-scales. During sectioning of the basal plate it is obvious that basal plate tissues are the hardest (woody) and possess adventitious root initials (Fig 2.3). The basal plate exhibit storage cells containing bundles of raphides within their central vacuoles. These cells are oriented in different directions. There are different shaped parenchymatous cells, i.e., circular and elongated and (Fig. 60).

3.2.3 Bulb histochemistry After employing the Schiff’s reagent the presence of cells with a polysaccharidic content in the bulb-scales becomes evident (Fig. 61). The red color intensity in the mesophyll cells of the bulb scales is variable in free-hand sections stained with the Schiff’s reagent (Fig. 62). These cells contain soluble polysaccharides in their large vacuole, which in some cases is more clearly seen after cells have been plasmolysed. The idioblastic cells containing bundles of raphides also react positively with Schiff’s reagent (Fig. 63). Moreover, large vessels, especially these that are close to the area where the xylem is more concentrated, are also red stained. As in the case of the bulb scales, some cells of the basal plate also react positively with Schiff’s reagent which is a convincing evidence of the presence of polysaccharidic content. In addition, all cells containing bundles of raphides react positively with this stain (Fig. 64).

It is worth mentioning that the red-color intensity of the positively reacted cells increases gradually from the innermost of the bulb-scales (Fig. 65) toward those present in the middle region (Fig. 63). The intensity decreases gradually toward the outermost bulb-scales (Fig. 66), up to the tunic which has no polysaccharidic contents. Thus, the most densely stained red-color cells occur in the third bulb-scale as illustrated in (Fig. 63). Moreover, all of the bulb cells that contain bundles of raphides react positively with Schiff’s reagent.

When semi-thin or hand-cut sections of bulb scales are treated with Sudan Black B, numerous cells appear to be intensely stained brown to black (Fig. 67). Oil cells occur sporadically as solitary idioblasts, but they do not differ greatly in size from their neighboring cells (Fig. 68). These oil cells accumulate abundant oil insoluble in water. The

Sharaf Al-Tardeh Results oil idioblasts are rounded and almost devoid of cytoplasmic content. Finally, oil droplets found in the cytoplasm, react positively with this stain.

Among the vascular cells of the vascular bundle there are parenchyma cells containing lipophilic substances (Fig. 69). The color intensity of these cells decreases gradually from the innermost bulb-scales toward the outermost ones. The basal plate tissues are negatively stain with Sudan Black B.

3.2.4 Bulb morphometry and reserves allocation patterns A morphometric analysis, listed in Table 3.3, was carried out in order to explain the relation between the bulb-scale reserve contents and their contribution to the total bulb-scale volume. Bulb-scales were arranged from the innermost to the outermost and analyses were carried out. The middle region of the bulb (almost, the 3rd bulb-scale) contains a considerable relative volume of the cells with polysaccharides (12.91 ± 2.43 %) and the highest percentage of cells containing lipids (21.92 ± 2.43 %) and mucilage (9.34 ± 1.03 %). The middle area of the bulb is the most efficient part in storing reserves, since the total relative volume of its storage cells is 50.66 %. The basal plate contains the highest relative volume of polysaccharidic contents (15.00 ± 1.62 %) and raphide inclusions (13.09 ± 1.67 %). Finally, lipids are almost absent from the outermost part of the bulb and the basal plate (approximately 0%). Allocation patterns (relative volumes %) of nutrient (resources) in the bulb parts of U. maritima at the onset of dormancy phase were illustrated in Fig. 3.1.

Sharaf Al-Tardeh Results

Table 3.3. Relative volumes (%) of the histological components with storage character in the bulb- scales and the basal plate of U. maritima. (Mean ± SD, n=10).

Relative volumes of the storage cells %

Storage cells First bulb- Third bulb- Seventh scale scale bulb-scale Basal plate Cells containing polysaccharides 5.56 ± 1.72 12.91 ± 2.43 6.59 ± 1.76 15 ± 1.62 Cells containing lipids 14.75 ± 2.54 21.92 ± 2.43 6.34 ± 2.96 Zero Idioblastic cells containing Raphides 5.85 ± 3.01 6.49 ± 1.47 5.38 ± 1.98 13.09 ± 1.67 Cells containing mucilagenous material No* 9.34 ± 1.03 No* No* Total 25.664 50.66 18.31 28.09 *Not calculated because the relative volumes were very low and in some cases could be zero.

Fig. 3.1. Allocation patterns (relative volumes %) of nutrient (resources) in the constituent bulb parts of U. maritima at the onset of dormancy phase.

3.2.5 Biomass allocation patterns and water status The water status and biomass allocation patterns within the bulb-scales and basal plate are differentially accumulated throughout the seasonal sequence (Table 3.4). The dormant bulb has the highest mean percentage of biomass allocation (32.50 ± 5.5%). During the stage of mature reproductive state (active phase) the bulb accumulates the lowest mean percentage of

Sharaf Al-Tardeh Results biomass (20.76 ± 5.5%), while at the stage of leaf emergence it accumulates a moderate mean percentage of biomass (26.09 ± 6.4%).

The water status within the bulb parts shows minor fluctuation through the stages of the plant’s annual life cycle (Table 3.4). The highest water content is found in the central bulb- scale (1st bulb-scale) (91 %) during the active phase. The lowest water content is recorded for the same bulb-scale zone (57%) during the dormancy phase. The bulb-scales show high capacity for water storage in their tissues, especially those of the middle region of the bulb. This is obvious when the water content, during the time of reproduction, is compared to that of the dormant bulb-scales (Table 3.4; Fig. 3.2). It is evident that the water content of the basal plate is the most stable throughout the year.

Table 3.4. Biomass percentage (%) and water contents (WC %) of the bulb-scales and the basal plate during the life cycle at the stage of leaf emergence (2 November 2006), at the mature leaf stage (2 January 2006), and at dormancy (dormant bulb) (12 July 2006) (Mean ± SD, n=5).

Biomass (%)† WC% Bioomass% WC% Biomass% WC% Bulb part At leaf emergence At leaf maturation At bulb dormancy

First bulb-scale* 26.67 73.33 9.02 90.98 39.09 60.91 Second bulb-scale 10.52 89.48 24.14 75.86 42.73 57.23 Third bulb-scale 21.52 78.48 25.37 74.63 24.21 75.79 Fourth bulb-scale 28.21 71.79 25.76 74.24 30.26 69.74 Fifth bulb-scale 29.73 70.27 24.53 75.47 30.16 69.84 Sixth bulb-scale 29.81 70.19 20.79 79.21 31.63 68.36 Seventh bulb-scale 30.08 69.92 18.48 81.52 30.25 69.75 Eighth bulb-scale 28.84 71.66 16.02 83.98 34.71 65.29 Basal plate 29.46 70.54 22.76 77.23 24.36 75.63 The bulb‡ 26 ± 6 73 ± 6 20 ± 5 79 ± 5 32 ± 5 67 ± 5 *The bulb scales are in order from one to eight followed by the tunics and the basal plate (Fig. 2.3). †Biomass is determined as dry mass percentage after drying to constant weight at 70 oC. ‡ Mean ± SD of the biomass of the whole bulb is calculated.

Sharaf Al-Tardeh Results

Fig. 3.2. Water status (%) is measured as total water contents in the constituent bulb parts of U. maritima. Measurements were done during the active phase and dormancy phase of the plant life cycle.

Sharaf Al-Tardeh Results

3.3 MORPHO-ANATOMICAL FEATURES OF THE LEAVES Individual plants of U. maritima, growing wild on a hill about 10 km NW of Chania, on the island of Crete, southern Greece, were used in this study. Leaves of the same age (3rd leaf from the rosette basis) and size (~20 cm length) were used in our investigations. Experiments and measurements were performed during the growing period (November- May) of 2004 and 2005. In all cases, main leaf drip was excluded.

3.3.1 Leaf anatomy U. maritima is a winter plant characterized by three phenological stages consisting, respectively, of inflorescence, leaves and no above-ground biomass. Leaves first appear after the flowers have wilted in response to winter onset rains during November or December in Crete-Greece, and may remain green until late spring (May), depending on rainfall and temperature.

The leaves are 714.2 ± 33.6 µm thick (Table 3.5 and Fig. 70). The main leaf drip is the thickest and the margins are the thinnest. The upper epidermis is slightly thicker (51.97 µm) than the lower epidermis (48.91 µm) (Table 3.5). Both the upper and lower epidermal cells are compactly arranged and covered with a relatively thick cuticle (~10 µm) (Table 3.5). The epidermal cells are very similar, polygonal elongated and possess rounded margins in longitudinal section (Fig. 71). In cross section, however, the epidermal cells are isodiametric and periclinally oriented with the outer walls slightly thicker than the other ones (Figs 70 & 72). The outer walls and the cuticle are more discernible in SEM micrographs in a tangential section (Fig. 73). The epidermal cells, especially those that are close to the guard cells, possess myelin-like structures (Fig. 74).

In the leaf of U. maritima, the stomata are arranged in rows parallel to the long axis of the leaf (Fig. 71). Their development begins at the tips of the leaves and progresses downwards. The leaves possess sunken stomata on both sides of the leaf (amphistomatic). The density of stomata in upper epidermis (736.83 ± 105.35 mm-2) is, somewhat, lower than that of the lower epidermis (867.06 ± 66.09 mm-2) (Table 3.5). The guard cell pairs have a mean length of 45.4 ± 2.3 µm and width of 22.8 ± 2.1 µm, as determined from SEM micrographs, and

Sharaf Al-Tardeh Results possess ledges (Fig. 72). The stomatal apparatuses are anomocytic, i.e., real subsidiary cells are absent (Figs 70 & 75). The stomatal configurations are almost 2-fold bigger in size than the adjacent mesophyll cells (Figs 70 & 72).

Table 3.5. Average thickness (µm) of leaf tissues in cross-sections [Mean ± Standard Deviation (SD), n = 15] and stomatal density (No. mm-2 ± SD) in longitudinal sections (n=15). Average leaf tissue Density of stomata Leaf tissue thickness (µm) (No. mm-2 ± SD) Adaxial cuticle 10.25 ± 1.16 Adaxial epidermis 51.97 ± 1.08 736.83 ± 105.35 Mesophyll 594.7 ± 33.3 Abaxial epidermis 48.91 ± 0.76 867.06 ± 66.09 Abaxial cuticle 8.41 ± 0.58 Total leaf thickness 714.2 ± 33.6

The leaves of U. maritima are equifacial, i.e., in terms of cuticle, epidermis and spongy cells, which, are found on both leaf sides. The mesophyll is 594.7 ± 33.3 µm thick. The spongy cells on the upper side of the leaves are elongated, while those of the lower side are oval to round in cross section (Fig. 70). In longitudinal section, however, both the upper and the lower side spongy cells are oval to rounded (Fig. 76).

The lower side (lower spongy mesophyll) of the leaf possesses idioblastic cells containing bundles of raphides which reflect the light under light microscope (Fig. 70). Raphides occur within the central vacuole. Under the electron microscope, the vacuolar content appears foamy and each crystal needle is embedded in a translucent homogeneous substance (Figs 77 & 78). The idioblastic cells contain also crystalloid inclusions of different shapes (Figs 79 & 80) and myelin-like structures. The cell wall of the idioblastic cells contains polysaccharides in a solid form (Fig. 81) and osmiophilic droplets (Fig. 82). In addition, mitochondrion is sometimes found beside the cell wall of an idioblastic cell (Fig. 83).

Sharaf Al-Tardeh Results

The bundle sheath of the leaf of U. maritima is hardly seen and does not possess either chloroplasts or bundle sheath extensions (Fig. 84). The absence or the presence of very few chloroplasts inside the bundle sheath cells and the frequent occurrence of chloroplasts inside mesophyll cells are a pronounced C3 plant character. The vascular bundles (the main veins) occur on rows parallel to each other and are crossed by small (secondary) veinlets (Fig. 85). Both xylem and phloem of (the) larger bundles are associated with sclerenchyma. However, in (the) smaller bundles the phloem is associated with sclerenchyma while the xylem is subtended by parenchyma. The phloem sieve elements are supported by the companion cells in order to maintain the osmotic pressure of the cell during transportation of the photosynthate products (Fig. 86). The presence of plasmodemata between the phloem sieve elements and the companion cells is evident (Fig. 87). Plasmodesmata are provided with callose to control the movements of the substances (Fig. 88). Plastids of the phloem sieve elements are characterized by peculiar protein crystalloid inclusions (Fig. 89). The protophloem sieve elements lose most of their organelles and keep only few plastids (Fig. 90). Finally, the parenchyma cells close to the vascular tissues show activity and possess Golgi bodies and mitochondria (Fig. 91).

The phenology of the chloroplast is correlated with the cell that belongs to, i.e., the chloroplasts of the upper spongy mesophyll cells are elongated (Fig. 92) and those of the lower ones are ovulated (Fig. 93), while some other chloroplasts are rounded and/or irregularly shaped (Fig. 94). The chloroplasts exhibit abundant osmiophilic droplets, phenolic compounds (Fig. 95) and starch granules (Fig. 96). Myelin-like structures, osmiophilic droplets, mitochondria and Golgi apparatus, usually, occur close to the chloroplasts (Fig. 95). Moreover, the grana are compactly stacked while the thylakoids show dilation (Fig. 96).

3.3.2 Leaf morphology and morphometry Leaves first appear after the flowers have wilted in response to (the) winter rains onset during November or December in Crete-Greece, and may remain green until late spring (May), depending on rainfall and temperature. The morphological traits of a leaf of the middle region are shown in Table 3.6. The leaves of U. maritima are 2-5 cm wide, 20-30 cm

Sharaf Al-Tardeh Results long, lanceolated in shape, somewhat undulated at the margin, shiny and dark-green in texture. The leaf surface area is almost 87.26 ± 13.93 cm2. The SLM and LTD of the leaf are 8.564 ± 3.139 mg cm-2 and 136.5 ± 14.41 mg cm-3, respectively.

Table 3.6. Morphological traits of a leaf (3rd leaf from the rosette basis) of U. maritima in the Mediterranean-type of climate (Mean ± SD, n = 10).

Leaf parameter Mean ± SD

Leaf length (cm) 25 ± 5

Leaf width (cm) 2.4 1.5 Leaf surface area (LSA, cm2) 87.26 ± 13.93 Leaf volume (V, cm-3) 6.232 ± 0.995 Leaf dry mass (LDM, mg) 843.5 ± 111.0 2 -1 Specific leaf area (cm g ) 103.5 ± 11.19 -3 Leaf tissue density (LTD, mg cm ) 136.5 ± 14.41 Specific leaf mass (SLM, mg cm-2) 8.564 ± 3.139

Leaf water content % 84.59 ± 1.424

A morphometric analysis, listed in Table (3.7), was carried out in order to explain the relation between the leaf components and their contribution to the total leaf volume. The relative volume of the spongy mesophyll cells is the highest (52.96 ± 1.42%), and that of the abaxial cuticle the lowest (2.21 ± 0.15%), while the corresponding one of the spongy air spaces and intercellular spaces is intermediate (30.41 ± 1.13%).

Table 3.7. Relative volumes (%) of leaf adaxial cuticle, adaxial epidermis, spongy mesophyll, abaxial epidermis and abaxial cuticle (Mean ± SD, n = 10).

Leaf tissue Relative volume %

Adaxial cuticle 2.41 ± 0.20

Adaxial epidermis 6.01 ± 0.97

Spongy mesophyll cells 52.9 ± 1.42

Spongy air spaces and intercellular spaces 30.4 ± 1.13

Abaxial epidermis 6.00 ± 0.21 Abaxial cuticle 2.21 ± 0.15

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3.3.3 Leaf histochemistry The presence of cells with polysaccharidic content in the leaf is evident after employing the Schiff’s reagent. The idioblastic cells containing bundles of raphides are positively reacting with this stain (Fig. 97). All of the cell walls, especially these of the vascular tissue are, also, red stained with Schiff’s reagent (Fig. 98).

When semi-thin or hand-cut sections are treated with Sudan Black B, numerous cells and/or part of cells appear intensely stained brown to black. Osmiophilic (oil) droplets occurring in the chloroplasts, (Fig. 95) nucleus and vacuoles (Fig. 99) react positively with this stain. Moreover, the sieve cells are stained brown to black with Sudan Black B (Fig. 100).

3.3.4 Leaf chlorophyll assay In order to understand the chlorophyll function and efficiency in relation to the Mediterranean climate, a leaf chlorophyll assay was done for leaves of U. maritima plants that were exposed to three environmental conditions: (1) natural habitat, (2) fully exposed to the sun, and (3) fully exposed to the shade for six months. The measurements in natural habitat were taken after three and six months from leaf emergence. The chlorophyll contents (µg cm-2 f. w.) of the leaves of plants grown in different environmental conditions are listed in Table 3.8.

Table 3.8. Chlorophyll content (µg cm-2 f. w.) of the leaves of U. maritima in natural habitat, fully exposed to the sun and fully exposed to the shade. (Mean ± SD, n = 15)

Leaf chlorophyll content (µg cm-2 f. w.) of plant of:

Chlorophyll Natural habitat Natural habitat Fully exposed to the Fully exposed to the (3 months old) (6 months old) sun (6 months old) shadow (6 months old)

Chl a 19.50 ± 0.60 19.16 ± 0.16 18.77 ± 0.25 19.02 ± 0.25 Chl b 14.06 ± 1.50 19.41 ± 2.92 13.47 ± 0.77 20.68 ± 2.07

Chl (a+b) 33.57 ± 2.12 38.57 ± 3.08 32.25 ± 0.98 39.70 ± 2.10 Chl b/chl a 0.72 ± 2.51 1.01 ± 0.15 1.39 ± 0.06 0.92 ± 0.08

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In general, chlorophyll a and b, and chlorophyll a/b ratio are affected slightly due to solar radiation. The highest value of chlorophyll a content was recorded for the plant of natural habitat after three months form leaf emergence (19.50 ± 0.60 µg cm-2 f. w.) followed by that one of six months. Meanwhile, the lowest value was recorded for plants that were fully exposed to the sun (18.77 ± 0.25 µg cm-2 f. w.). The highest value of chlorophyll b content was recorded for plants grown in habitats fully exposed to the shade (20.68 ± 2.07 µg cm-2 f. w). Meanwhile, the lowest value was recorded for plants grown in habitats fully exposed to the sun (13.47 ± 0.77 µg cm-2 f. w.). It is obvious from Table 3.8 that chlorophyll b was more sensitive to solar radiation than chlorophyll a. However, both chlorophyll a and b contents decreased in the plants that were fully exposed to the sun light.

Morphological variations were also recorded between plants grown fully exposed to the sun and those fully exposed to the shade. The shade plants possessed more elastic, weak and thin leaves. Moreover, the leaves of the shadow plants were longer than those of the sun plants (Figs 101 & 102). The specific leaf mass (SLM) for the plants fully exposed to the sun (8.664 ± 0.447 mg cm-2) was higher than that of those fully exposed to shade (5 ± 0.1 mg cm-2).

3.3.5 Chlorophyll a fluorescence Two groups of U. maritima were grown from leaf emergence till just before their senescence (approximately six months). The first group was fully exposed to the sun (light leaf), while the second group was fully exposed to shade (shade leaf). Chlorophyll fluorescence was then measured after six months from leaf emergence. The PSII activity reflected by the maximal efficiency of PSII photochemistry measured as Fv/Fm, was decreased in the light leaf (0.824 ± 0.008) versus that of the shade one (0.836 ± 0.030).

In order to understand the efficiency of PSII, the total quantum yield of PSII photochemistry was measured (Fig. 3.3). Both shade and light leaves show higher quantum yield at lower photon flux intensities. While, the photon flux intensity or the photosynthetic active radiation (PAR) increased more than 536 μmol quantum photons m-2 s-1, the quantum yield

Sharaf Al-Tardeh Results decreased. Moreover, the shade leaf shows higher activity than the light leaf at the PAR range from 66 to 536 μmol photons m-2 s-1, after which the light leaf is more active.

Fig. 3.3. Light curve of the total quantum yield of PSII photochemistry of the light and shade leaves of U. maritima.

Electron transport rate (ETR) was also measured to interpret the apparent rhythm of electron transport through PSII. As it shown in Fig. 3.4, the photon flux intensity of 536 μmol photons m-2 s-1 seems to be the point of altering the photochemical efficiency of the PSII in the shade and light leaves.

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Fig. 3.4. Light response curve of electron transport rate of the light and shade leaves of U. maritima.

Non-photochemical quenching (qN) of the light and shade leaves of U. maritima was measured in order to predict the amount of energy released in the form of heat (Fig. 3.5). The light leaf shows higher qN than that of the shade leaf when exposed to the photon flux density of 66 to 536 μmol photons m-2 s-1. However, at higher photon flux intensity, both qN of both leaves are overlapped.

Fig. 3.5. Light response curve of non-photochemical quenching parameter showing the amount of energy released in the form of heat by U. maritima light and shade leaves.

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3.4 MORPHO-ANATOMY OF THE INFLORESCENCE The position of the nectary of U. maritima was examined using an Olympus SZX12 stereo- microscope. Fresh hand-cut sections of the nectary were obtained and tested for their polysaccharide and lipid contents by using Schiff’s reagent and Sudan Black B, respectively. These examinations and the fixation process were applied to flowers during the flowering anthesis (August- September 2006).

3.4.1 Flower morphology The flowering shoot of U. maritima is characterized by an interesting one-dimensional growth system. Stalk emergence from the bulb is completed in a few days and is followed by a characteristic stoically rapid pattern of subapical elongation. Shoot length is considered to be the peduncle plus the rachis or flower-bearing portion. Elongation of rachis is accompanied by flowering opening in the manner shown in Fig. 103. Their appearance is most likely to be an open-flower zone continually moving toward the apex.

The inflorescence is eremurus-like and contains more than 250 florets. Bracts are subulate, often caduceus and shorter than pedicels. The pedicels are more or less erect. The flower of U. maritima consists of a heavy sclerefied calyx of six elongated white petals, six conspicuously long stamens with green to deep brown anthers and a central gynoecium with superior ovary (hypogynous flower) (Fig. 104). The light green trilocular ovary has isomerous carpels. The lateral faces of the carpels are united by fusion with one another (syncarpous gynoecium) and coalesce to form a cavity where nectar accumulates (Fig. 105). The nectar drains from the region around the inferior two-thirds of the ovary to its base and accumulates there into mucilaginous droplets, one for each nectary. Moreover, the presence of modified stomata inside that groove (Fig. 106) and in the outer surface of the ovary is evident (Fig. 107). The guard cell pairs have an average length of 38.95 ± 2.10 µm and 22.75 ± 2.02 µm width, as determined from SEM micrographs.

The androecial members of the six whitish perianth parts are organized into one whorl and are free from each other (Fig. 104). The stamen consists of the peduncle-like filament, upon which is borne a two-lobed anther containing four microsporangia (ca. ~5 mm) or pollen

Sharaf Al-Tardeh Results sacs (Figs 108 & 109). The anthers contain few peculiar stomata, which occur on its groove (Fig. 110). The guard cell pairs have an average length of 32.67 ± 3.29 µm and width of 21.97 ± 2.46 µm, as determined from SEM micrographs. Petals and stamens are attached to the receptacle below the ovary. The flower is actinomorphic in which the corolla is made up of similarly shaped petals that radiate from the centre of the flower and are equidistant from each other. Moreover, when the flower senesces (old stage), the petals do not fall but close up again and resume a cylindrical structure similar to the buds (zone 3; Fig. 103). The closed petals protect the growing fruit capsule. When the fruit is ripe, it splits open to release its black winged seeds.

3.4.2 Anatomy and development of the gynopleural (septal) nectary 3.4.2.1 Young stage nectary The nectary of U. maritima was investigated at stages of young, intermediate and old flower. Serial sections beginning from the top of the stigma till the receptacle were obtained and examined by LM and TEM. The stigma receives the pollen and is connected to the ovary by an elongated structure called style (ca. ~3 mm). The first (from the top) third of the trilocular ovary is composed of empty canals through which the pollen grains follow their way to the ovary in order to fertilize the ovules (Figs 111 & 112). At the end of the first third of the ovary, the nectary tissues start to appear (Fig. 112). Idioblast cells containing bundles of raphides are scatterly present among the parenchyma cells of the first third of the ovary (Figs 111 & 112).

At the middle of the ovary, the syncarpous gynoecium consists of three carpels that are separated from each other by distinct septal slits. Between these slits, the floral nectary is developed (Fig. 113). The three septal slits proceed downwards entering the ascidiate zone of the carples (Fig. 103). The cavities are lined by one layer of compactly arranged secretory epithelium (Ne) (Fig. 114). Beneath the epithelium there is a subsidiary tissue (St) composed of vacuolated cells, which are smaller than those in other parts of the ovary parenchyma (Fig. 114). The subsidiary tissue is approximately 2-5 cells wide. Idioblastic cells (Id) containing bundles of raphides are located among the subnectary (Sn) and ovary parenchyma (Fig. 114). A very thin and irregular cuticle is present on the surface of the

Sharaf Al-Tardeh Results secretory epithelium (Fig. 114 & 116). The epithelium is made up of small cells with dense cytoplasm and large nuclei (Fig. 115). The nectary cavity is very narrow and not well developed (Fig. 115). Cisternal profiles of endoplasmic reticulum (ER) dominate the cytoplasm of the secretory epithelium (Fig. 116). The cytoplasm is densely packed with ribosomes, many of which are attached to the endoplasmic reticulum.

Development of the starch granules starts (commences) in the secretory epithelium cells, which contain mostly elongated plastids. These are mainly devoid of starch granules at the beginning of flower development (Fig. 117). At the end of the young stage, the secretory epithelium cells become active and possess many dictyosomes and fully developed large starch grains (Fig. 118). However, starch grains are more conspicuous and larger in the subsidiary tissue (Fig. 119).

The ovarian epidermal cells are elongated, compactly constructed, with concaved margins and are anticlinally oriented (Figs 111-113). In addition, they are covered by a smooth cuticle (Fig. 120), which is more evident under higher magnification (Fig. 121). The cuticle is 0.52 ± 0.25 µm thick.

Histochemical analyses were applied to transverse sections of the ovary tissues in order to examine the polysaccharidic and lipophilic substances. The results suggest that there is a low polysaccharidic content since there are few idioblastic cells containing bundles of raphides that are red stained after employing the Schiff’s reagent (Fig. 122). However, the absence of lipophilic content is evident, because all the experiments yield negative results with Sudan Black B reagent.

3.4.2.2 Nectary at the intermediate stage Each nectary cavity has a nectar outlet located at one-third of the distance of the ovary from the base of the style (Fig. 105). Nectar drains from the region of around the inferior two- thirds of the ovary to its base. Nectar accumulates there into mucilaginous droplets, one for each nectary, as a secondary presentation to be exposed to nectar-feeding animals. The nectar outlet derives from merging of an invagination of the cutinized epidermal surface, in

Sharaf Al-Tardeh Results continuity with the carpellary suture with the apical part of the nectary (Fig. 123). Small cells are present in the vicinity of the merging point (Fig. 123). The carpellary suture is wide at the base of the ovary, but it becomes deeper and narrower towards the top of the ovary. At the distance of the two-thirds from the top of the ovary, the invagination of the epidermal surface has tightly connivent margins, except for the inner part where a tubular structure is formed (Fig. 123). The tubular structure becomes deeper towards the base of the ovary, where it merges with the apical part of the nectary. Although the tubular structure is in continuity with the outside, this communication is obstructed by the presence of the cuticle that occludes the narrow space between the connivent margins of the epidermis.

The intermediate (active stage) stage of the nectary and/or ovary is referred to the time of flower’s anthesis and nectar secretion. The active ER comprises a prominent feature of the epithelial cells during the active stage of nectar secretion. In the dense granular cytoplasm fully developed ER cisternae with attached ribosomes forming characteristic profile is located in the vicinity of the outer cell walls (Fig. 124). Rough endoplasmic reticulum with dilated cisternae becomes predominant (Fig. 125). Moreover, the secretory epithelial cells contain many dictyosomes and mitoconderia (Fig. 126), many, narrow and circular mitochondria (Fig. 127), numerous secretory vesicles, myelin-like figures, and vacuoles with electron dense materials (Fig. 128). In addition, various shaped plastids containing osmiophilic granules in the stroma, peripheral thylakoids and different sized starch grainules (Fig. 125).

The nectary is supported by the parenchymatic septal tissue (subnectary parenchyma), which consists of approximately 2-6 layers of relatively large isodiametric parenchyma cells. The ovary wall is comprised of approximately 3-10 layers of parenchyma cells (Fig. 134). The subnectary parenchyma is located below the nectary parenchyma (subsidiary tissue), and consists of larger cells, with bigger vacuoles, less dense cytoplasm, and larger intercellular spaces (Fig. 114). Amyloplasts are variously shaped and almost devoid of stroma and packed with starch. They also contain many starch grains per amyloplast. This increases starch surface area, thus facilitating and speeding up hydrolysis during nectar production (Fig. 129). Clusters of amuloplasts (Fig. 130) with starch grains are more

Sharaf Al-Tardeh Results concentrated in the peripheral parenchyma cells (small micrograph in the corner of Fig. 123). During nectar secretion, starch content drastically diminishes from the nectary epithelium toward the peripheral parenchyma cells.

The vascular system is simple and looks like that of the leaf, bulb and root of the plant. The vascular tissue occurs in the subnectary parenchyma near to the placenta. More specifically, it is located at the interior side of the nectary in conjunction with the obturator tissues (Fig. 114). It might serve both the nectary and the obturator glands. The vascular bundles consist of xylem vessels and sieve elements (Fig. 131). However, vascular tissue that occurs close to the interior nectary gland consists of xylem vessels only (Fig 311). Whereas, the vascular tissue, which occurs in pairs, one on each side of the nectary, consist of phloem strands only (Fig. 132). The sieve elements are connected by the sieve pores (Fig. 132). Plastids with crystalloid inclusions are also present in the mature sieve elements (Fig. 132).These bundles without xylem do not extend so far up to the ovary, and hence have disappeared. The bundles supply the nectary with carbohydrates. Therefore, they are absent from the region where the nectary tissues are reduced as in the top of the ovary.

The nectary at the one-third distance from the bottom of the ovary becomes shorter and almost wider than the other two zones (Fig. 133). From the hystochemical point of view, the highest concentration of polysaccharides occurs in the first-third from the base of ovary because of the strong reaction showed with Schiff’s reagent (Fig. 134). The intensity of the red colour of the idioblastic cells containing bundles of raphides is higher than that of the young stage (Fig.134 & 122). ). Moreover, the starch granules in the ovary wall react positively with this stain (Fig 134). In addition, the nectar remnants inside the nectary cavity and the starch granules in the subnectary parenchyma react positively with Schiff’s reagent (Fig. 135). However, there are negative reactions with Sudan Black B due to the absence of lipophilic substances.

3.4.2.3 Nectary at old stage The old (overmature) stage starts 19 hours from the flower opening. The epithelial and subsidiary tissue cells undergo cytological modification during development, being more

Sharaf Al-Tardeh Results vacuolated and with an irregular nuclear shape in the old flower stage (Fig. 136). Some cells in the subsidiary tissue seem to degenerate (Fig. 136). The thickness of the nectary decreases in the old stage, being approximately six cells wide from the epithelium to the ovary wall (Fig. 136).

The pores (microchannels), through which the nectar is discharged into the nectary cavity, are more obvious in the cell wall of the epithelial cells (Fig. 137). Microchannels are narrow tubular in continuity with the cell wall; some of them seem to have direct communication with the outside (Fig. 138). The old stage is characterized by crystallized form of nectar inside the epithelial cells (Fig. 139), collapse of the cortical parenchyma cells (Fig. 140), completely hydrolysed starch, as well as disappearance of the amyloplasts and ER. The relation between nectar production and starch hydrolysis is dynamic in the manner that almost all of the starch content is hydrolyzed at the old flower stage.

3.4.3 The ovary and ovules The ovary of U. maritima is partitioned into three locules formed from the fusion of three carpels (Fig. 113). The carpels contain several ovules, almost four per each one at the mature stage (Fig. 141). However, it could be less than four ovules per each carpel at the initial developmental stage (Fig. 113). After fertilization, the embryo is developed into seed. At the stage of overmature ovary, the seed coat appears to be well developed and the embryo is ready to germinate (Fig. 142). There are about 12 seeds per capsule.

The presence of cells of polysaccharidic content is evident in the tissues of the body of the ovary. These cells are red stained with Scheff’s reagent (Fig. 143). However, they give negative results when it reacts with Sudan Black B. The polysaccharide concentration seems to be the highest at the old stage (Fig. 143) and the lowest at the young stage (Fig. 122), while the intermediate one is at the intermediate stage (Fig. 134).

The relative volumes (%) of the idioblastic cells containing bundles of raphide were calculated from 10 light micrographs for each stage and are shown in Fig. (3.7). The relative volume of these cells was calculated in proportion to the ovarian tissues (excluded from the

Sharaf Al-Tardeh Results ovules). The highest value is recorded for the young stage (7.41 ± 1.8 %). The relative volume decreases at the intermediate stage (4.33 ± 1.8 %) and increases again at the old stage (6.45 ± 1.55 %).

Fig. 3.7. Relative volume (%) of the idioblastic cells containing bundles of raphides distributed in the ovary tissues (excluded from the ovules) of U. maritima (L.) Baker at young, intermediate and old stage. (Mean ± SD; n = 30).

3.4.4 Obturator gland Placentation is axile, in which the ovules are borne on a central column of tissue in the partitioned ovary of U. maritima. The obturator gland is a prominent ovary wall outgrowth of placental origin, which lies in close contact with the micropyle of each ovule (Figs 141). The secretory cells of the obturator are epithelial and possess cisternal elements of the endoplasmic reticulum that are in parallel arrangement with dilated cisternae (Fig. 144). Among the endoplasmic reticulum cisternae, large vesicles with granular content occur (Fig. 145). Plastids occur sporadically in the secretory cells. Most of these plastids contain huge starch granules during the initial stages (Fig. 146). However, the plastids become devoid from their contents at the stages of mature and overmature ovary and/or nectary.

Dictyosomes are prominent inside the secretory cells of the obturator gland. Both dictyosomes and endoplasmic reticulum are involved in the process of mucilage production and its subsequent secretion. The mucilage is secreted by the dictyosomes in the form of

Sharaf Al-Tardeh Results mucoprotein, while the protein units were provided by the endoplasmic reticulum. Moreover, the glandular cells are internally surrounded by an extraplasmic space filled with mucilage (Fig.146). The presence of plasmodesmata between the epithelial cells of the obturator is also evident (Fig. 147). The obturator gland diminishes to its minimum size and becomes inactive at the old stage.

Fig. 1. An overview photo of U. maritima with a bulb diameter of 10 cm sampled from a natural population in Souda (Greece). The photo illustrates the typical view of the plant and its root system which consists of about 10 adventitious roots attached to the basal plate disk.

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Fig. 2. Photo of U. maritima with a bulb diameter of 10 cm sampled from a natural population in Souda (Greece). The root system consists of adventious roots that have unlimited growth downward and could have up to five branches.

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Fig. 3. LM micrograph of a transverse section of the adventitious root (overview) of U. maritima revealing the velamen and the hypodermis. Scale bar = 50 µm.

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Fig. 4. LM micrograph of a transverse section of the root of U. maritima revealing the multiple-layered velamen (Ve) with root hairs (Rh). (Ep: epidermis). Scale bar = 100 µm.

Figs 5-8. TEM micrographs of transverse sections of the velamen of a root of U. maritima.

Fig. 5. Cell walls of velamen cells reveals densely packed zig-zag-like folding (arrows) during the summer period. Scale bar = 1 µm.

Fig. 6. A velamen epidermal cell contains myelin-like structures (Ms). Scale bar = 10 µm.

Fig. 7. Different myelin-like structures (Ms) occur inside the velamen epidermal cells. Scale bar = 1 µm.

Fig. 8. Electron dense materials (Edm) occur inside the intercellular space and the central vacuole of the velamen cells. Scale bar = 10 µm

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Figs 9-12. TEM micrograph of a transverse section of the velamen of a root of U. maritima. Scale bar = 1 µm. Fig. 9. Putative bacteria (Ba) and electron-dense materials occur inside the velamen epidermis.

Fig. 10. Intercellular space of the velamen epidermis contains electron-dense material (Edm).

Fig. 11. Extracellular space of the velamen epidermis possesses amorphous electron- dense remnants (Edr).

Fig. 12. Intercellular space of the velamen cells has peculiar electron-dense material (Edm).

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Fig. 13. LM micrograph of a transverse section of the root of U. maritima showing the cortex region with cells of water-storing character, idioblastic cells with bundles of raphides (Id) and cells with mucilage in their central vacuole (Mc). (Ed: endodermis, Ph: phloem, X: xylem). Scale bar = 50 µm.

Fig. 14. LM micrograph of a transverse section of the root of U. maritima at the cortex region revealing needles of raphides (Rb) stored in central vacuole of a cortical idioblastic cell beside the mucilage ones (Mc). Scale bar = 25 µm.

Fig. 15. TEM micrograph of a transverse section of the root of U. maritima at the cortex region showing cortical idioblastic cell with densely packed raphides (Rb) within the central vacuole. Scale bar = 1 µm.

Fig. 16. LM micrograph of a longitudinal section of the root of U. maritima at the cortex region showing rows of cells with bundles of raphides. Scale bar = 50 µm.

Fig. 17. LM micrograph of fresh, hand-cut longitudinal section of the root of U. maritima at the cortex region showing rows of cells with bundles of raphides. Scale bar = 50 µm.

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Figs 18-22. TEM micrographs of a transverse section of the root of U. maritima showing the fine structure of the cortex region. The sections were made at the region of 3 cm above the root tip. Scale bar = 1µm except for Fig. 18, scale bar = 10 µm

Fig. 18. Idioblastic cortical cell possesses central vacuole filled with homogenous substances. Primary raphide needles (arrows) are embedded in the homogenous substances.

Fig. 19. Details of Fig. 18. A connection between the central vacuole and the cell wall through which translocation of the homogenous substances might occur.

Fig. 20. Cortical intercellular space (Is) containing homogenous substances. Moreover, many organelles accumulate close to the intercellular space. (ER: endoplasmic reticulum; Mt: mitochondria; G: Golgi body; Ms: myelin-like structure; Pd: plasmodesmata)

Fig. 21. Cortical cells containing myelin-like structure (Ms), while possessing a lobed nucleus.

Fig. 22. Cortical cell possessing a straight endoplasmic reticulum (ER).

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Figs. 23-25. Micrographs of a transverse section of the root of U. maritima showing the fine structure of the central cylinder. Scale bar = 25µm except for Fig. 23, scale bar = 50 µm.

Fig. 23. LM micrograph of a transverse section of the root of U. maritima showing the central cylinder with 10-arched xylem alternating with a cluster of phloem cells (Px: protoxylem; Mx: metaxylem; Ed: endodermis).

Fig. 24. TEM micrograph of a transverse section of the root of U. maritima showing the uniseriate endodermis (Ed). The cells are periclinally oriented.

Fig. 25. TEM micrograph of a transverse section of the root of U. maritima showing the phloem cells (Ph) and the active parenchyma cells (Pc).

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Fig. 26. LM micrograph showing the typical vessel member of the root of U. maritima as a model for the angiosperm. The vessel members were separated from other cells by the maceration technique. Scale bar = 500 µm.

Fig. 27. Continuation of Fig. 26 showing the other end of the vessel member of the root of U. maritima after maceration. Scale bar = 500 µm.

Fig. 28. TEM micrograph of a transverse section of the root of U. maritima showing the central cylinder parenchyma cell with multi-lobed nucleus. Scale bar = 10 µm.

Fig. 29. TEM micrograph of a transverse section of the root of U. maritima showing a lobed nucleus in the parenchyma cell of the central cylinder. Scale bar = 10 µm.

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Figs. 30-33. TEM micrographs of a transverse section of the root of U. maritima revealing the ultrastructure of the central cylinder. Scale bar = 10 µm, except for Fig. 33, scale bar = 1 µm.

Fig. 30. Plasmodesmata (Pd) are located between the parenchyma cells of the central cylinder.

Fig. 31. Parenchyma cells of the central cylinder show an intens shrinkage (arrows) due to the loss of water during the dry and hot summer. It could be also the result of solution loss during the fixation process.

Fig. 32. Long and uniseriate endoplasmic reticulum (arrows) present in the companion cell of the central cylinder.

Fig. 33. Myelin-like structure (Ms) is located inside the parenchyma cell of the central cylinder.

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Fig. 34. LM micrograph of a transverse section of the root of U. maritima showing scattered cortical cells (arrows) with polysaccharide contents, red-stained with Schiff’s reagent. The idioblastic cells, containing bundles of raphides, which react positively with this stain, are shown in the inset. Mucilage cells (Mc) do not react positively with this stain. Scale bar = 50 µm.

Fig. 35. LM micrograph of a transverse section of the root of U. maritima showing scattered cortical cells (arrows) with lipid contents stained brown to black after employing Sudan Black B. Idioblast cells (Id) containing bundles of raphide do not react positively with this stain. Scale bar = 50 µm.

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Fig. 36. SEM micrograph of the bulb of U. maritima revealing the tunics with anomocytic stomata. Scale bar = 20 µm.

Fig. 37. LM micrograph of the bulb-scales of freshly peeled epidermis of U. maritima revealing anomocytic stomata after employing the tolidine blue. Scale bar = 500 µm.

Fig. 38. LM micrograph (overview) of a transverse section of the bulb of U. maritima revealing the third bulb-scale with cells of storage character after employing the tolidine blue. (Ab: abaxial epidermis; Ad: adaxial epidermis; Id: idioblastic cell containing bundles of raphides; Mc: mucilage cell: Vb: vascular bundle; Oc: oil cell) Scale bar = 100 µm.

100

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101

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Fig. 39. LM micrograph of a transverse section of the bulb of U. maritima revealing the abaxial epidermis (Ab) of the third bulb-scale. (Cu: cuticle; Mc: mucilage cell; Id: idioblastic cell containing bundles of raphides). Scale bar = 50 µm.

Fig. 40. LM micrograph of a transverse section of the bulb of U. maritima revealing the adaxial epidermis (Ad) of the third bulb-scale. (Cu: cuticle; Id: idioblastic cell containing bundles of raphide; Mc: mucilage cell). Scale bar = 50 µm.

102

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103

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Fig. 41-44. TEM micrographs of a transverse section of the bulb of U. maritima showing the fine structure of the epidermis: Fig. 41. Abaxial epidermis has periclinal walls thicker than the anticlinal ones. Scale bar = 10 µm.

Fig. 42. Adaxial epidermis has anticlinally oriented cell wall. Scale bar = 1 µm.

Fig. 43. The anticlinally oriented cell wall of the adaxial epidermis possesses plasmodesmata (arrows). Scale bar = 1 µm.

Fig. 44. Golgi body (arrow) occurs inside epidermal cells. Scale bar = 1 µm.

104

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105

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Fig. 45. LM micrograph of a transverse section of the bulb of U. maritima revealing the vascular bundle and the cells of storage character. (Id: idioblastic cell containing bundles of raphides; Mc: mucilage cell; X: xylem; Ph: phloem). Scale bar = 50 µm.

Fig. 46. TEM micrograph of a transverse section of the bulb of U. maritima revealing a mesophyll idioblastic cell with raphide needles (Rn) embedded in polysaccharidic content. Scale bar = 10 µm.

Fig. 47. TEM micrograph of a transverse section of the bulb of U. maritima revealing an idioblastic cell in mesophyll with homogenous material of polysaccharides. Scale bar = 10 µm.

106

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107

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Fig. 48. LM micrograph of a fresh hand-cut longitudinal section of the bulb of U. maritima revealing an idioblastic cell with bundle of raphides. The idioblastic cell is seven times longer than the normal parenchymatous ones. Scale bar = 500 µm.

Fig. 49. LM micrograph of a fresh hand-cut transverse section of the bulb of U. maritima revealing open bundle of raphide needles which are about 500 µm long. Scale bar = 500 µm.

108

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109

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Figs. 50-53. TEM micrographs of transverse sections of the bulb of U. maritima, illustrating the fine structure of the ground tissue (mesophyll) of the third bulb-scale. Scale bar = 1 µm except for Fig. 53, scale bar = 0.5 µm.

Fig. 50. Idioblastic cell containing a mucilaginous material.

Fig. 51. Plasmodesmata (arrows) are located between the active mesophyll cells.

Fig. 52. Osmiophilic droplets (Os) are found inside the ground tissue cells near the lower epidermis.

Fig. 53. Straight endoplasmic reticulum (ER) and mitochondria (Mt) occurring in the mesophyll cells near the lower epidermis.

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Figs. 54-59. TEM micrographs of transverse sections of the bulb of U. maritima illustrating the fine structure of the vascular system of the third bulb-scale. The vascular system consists of scatterly vascular bundles of phloem and xylem.

Fig. 54. The phloem cells consisted of sieve-tube elements (SE) in conjunction to the companion cell (Cc). Scale bar = 10 µm.

Fig. 55. A sieve element (SE) is harder, thicker and devoid of most of its contents in the mature stage of the phloem development (metaphloem sive element). Scale bar = 10 µm.

Fig. 56. TEM micrograph showing sive pores between the sive elements. The the sieve pores are provided by callose. Scale bar = 1 µm.

Fig. 57. TEM micrograph reveals plastid with proteinous inclusions inside a metaphloem sieve element. Scale bar = 1 µm.

Fig. 58. Large mitochondria (Mt) are located in the parenchyma cells beside the vascular tissues. Scale bar = 1 µm.

Fig. 59. Straight cisternae of endoplasmic reticulum (ER), Golgi body (G) and mitochondria (Mt) are located in the parenchyma cells near the vascular tissues. Scale bar = 10 µm.

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Fig. 60. LM micrograph of a transverse section of the basal plate of an U. maritima bulb showing idioblast cells containing raphide bundles. These cells (arrows) are oriented in different directions. Scale bar = 50 µm.

Fig. 61. LM micrograph of a transverse section of the third bulb-scale of an U. maritima bulb showing the red stained cells (arrows) after employing the Schiff’s reagent. Scale bar = 100 µm.

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Fig. 62. LM micrograph of fresh hand-cut transverse section of an U. maritima bulb scale showing cells containing soluble polysaccharides stained red with Schiff’s reagent (arrows). Scale bar = 100 µm.

Fig. 63. Idioblast cells (arrows) of an U. maritima bulb scale containing a bundle of raphides embedded in polysaccharide that has stained red with Schiff’s reagent. Scale bar = 50 µm.

Fig. 64. LM micrograph of a transverse section of the basal plate of an U. maritima bulb with idioblast cells (arrows) containing bundles of raphides embedded in polysaccharide that stained red with Schiff’s reagent. Scale bar = 100 µm.

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Fig. 65. LM micrograph of a transverse section of the bulb of an U. maritima showing the color intensity of the first bulb-scale after employing the Schiff’s reagent. Scale bar = 100 µm.

Fig. 66. LM micrograph of a transverse section of the bulb of an U. maritima showing the color intensity of the seventh bulb-scale after employing the Schiff’s reagent. Scale bar = 50 µm.

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Figs 67-69. LM micrographs of a transverse section of the third bulb-scale of an U. maritima after employing Sudan Black B.

Fig. 67. Numerous cells (arrows) containing lipids and oils in their central vacuoles stained brown to black with Sudan Black B. (Ad: adaxial epidermis; Ab: abaxial epidermis) Scale bar = 100 µm.

Fig. 68. Numerous cells (arrows) stained brown to black with Sudan Black B due to the presence of oil and lipids content. Scale bar = 10 µm.

Fig. 69. Cells within a vascular bundle that have reacted positively with Sudan Black B. (Sc: sclerenchyma fibers, Oc: oil cell, X: xylem, Ph: phloem). Scale bar = 10 µm.

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Fig. 70. LM micrograph of a transverse section (overview) of the leaf of U. maritima revealing idioblastic cells containing bundles of raphides (Id). (Ade: adaxial epidermis; Abe: abaxial epidermis; Abc: abaxial cuticle; Adc: adaxial cuticle; As: air space; X: xylem, Ph: phloem). Scale bar = 50 µm.

Fig. 71. LM micrograph of hand-cut paradermal section the leaf of U. maritima revealing stomata occurring in rows parallel to the main axis (drip). Scale bar = 50 µm.

Fig. 72. LM micrograph of a transverse section of the leaf of U. maritima revealing stomata complexes in adaxial epidermis. (Ade: adaxial epidermis; Lg: ledges; Gc: guard cells; Sch: stomatal chamber). Scale bar = 25 µm.

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Fig. 73. SEM micrograph of a tangential view of a transverse section of the leaf of U. maritima showing the mesophyll, epidermis and the cuticle, and stoma. Scale bar = 100 µm.

Fig. 74. TEM micrograph of a transverse section of the leaf of U. maritima showing the epidermal cell beside the guard cell with myelin-like structure (Ms). Scale bar = 1 µm.

Fig. 75. SEM micrograph of a paradermal view of the leaf of U. maritima showing anomocytic stomata. Scale bar = 50 µm.

Fig. 76. LM micrograph of a longitudinal section of the leaf of U. maritima showing the lower mesophyll region. Scale bar = 50 µm.

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Fig. 77. TEM micrograph of a transverse section of the leaf of U. maritima showing idioblast cells with crystal needles (Cn) embedded in polysaccharidic content. Scale bar = 5 µm.

Fig. 78. TEM micrograph of a transverse section of the leaf of U. maritima at the top of the idioblastic cell showing the homogenious translucent material. Scale bar = 10 µm.

Fig. 79-80. TEM micrographs illustrare the areas marked by big and small sequres in Fig. 78, repectively. The idioblastic cell possesses crystalloid inclusions (Ci) in different shapes. Scale bar = 1 µm.

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Fig. 81-83. TEM micrographs of a transverse section of the leaf of U. maritima shows parts of an idioblast cell. Scale bar = 1 µm.

Fig. 81. Polysaccharide content that might be in a solid form (arrows).

Fig. 82. Idioblast cell contains osmiophilic droplets (arrows).

Fig. 83. Idioblast cell possesses a mitochondrion (arrows).

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Fig. 84. LM micrograph of a transverse section of the leaf of U. maritima revealing the bundle sheath (Bs) and the vascular system. (X: xylem; Ph: ploem). Scale bar = 50 µm.

Fig. 85. LM micrograph of a longitudinal section of the leaf of U. maritima showing the vascular system with main vein (Pv) crossed by the veinule (Sv). (X: xylem; Ph: phloem). Scale bar = 50 µm.

Fig. 86. TEM micrograph of a transverse section of the leaf of U. maritima showing the sieve-tube elements (SE) and companion cell (Cc) in the vascular bundle. (Os: osmiophilic droplet; Pd: plasmodesmata; Pl: plastids). Scale bar = 1 µm.

Fig. 87. TEM micrograph of a transverse section of the leaf of U. maritima showing plasmodesmata (Pd) between SE and Cc. Scale bar = 1 µm.

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Figs 88-91. TEM micrographs of a transverse section of the leaf of U. maritima revealing phloem cells. Scale bar = 1 µm.

Fig. 88. Sieve pores (Sp) occur between SE and Cc provided by callose (Ca).

Fig. 89. Plastids containing peculiar protein inclusions inside the phloem sieve elements.

Fig. 90. Plastids (arrows) occuring inside the phloem sieve elements during its old stage.

Fig. 91. Golgi bodies (G) and a mitochondrion (Mt) are located in a parenchyma cell close to the vascular tissue.

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Figs 92-96. TEM micrographs of a transverse section of the leaf of U. maritima showing chloroplasts. Scale bar = 1 µm, except for Fig. 95, scale bar = 0.5 µm.

Fig. 92. Elongated chloroplasts occur in the upper mesophyll.

Fig. 93. Oval chloroplasts occur in the middle mesophyll.

Fig. 94. Spherical to oval chloroplasts occur in the lower mesophyll.

Fig. 95. Chloroplast possesses osmiophilic droplets (Os), while other structures such as mitochondria (Mt) and myelin-like structures (Ms) occur close to chloroplast.

Fig. 96. Part of a chloroplast has starch granules (Sg) and possessing a stacked grana (Gr) and delated stroma (arrows).

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Figs 97-100. LM micrographs of a transverse section of the leaf of U. maritima after employing the histochemals. Scale bar = 50 µm, except for Fig. 98, scale bar = 100 µm.

Fig. 97. Lower mesophyll possesses idioblastic cell containing a bundle of raphides embedded in polysaccharidic content stained red with Schiff’s reagent (arrows).

Fig. 98. Vascular tissue possesses cell walls stained red with Schiff’s reagent.

Fig. 99. Leaf tissues reveal cells and oil droplets stained brown to black with Sudan Black B.

Fig. 100. Vascular system contains numerous cells containing lipids stained brown to black with Sudan Black B.

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Fig. 101. A digital camera photo (overview) of U. maritima plant grown fully exposed to shade for six months from leaf emergence showing morphological differences with the plant grown fully exposed to the sun light (Fig.102).

Fig. 102. A digital camera photo (overview) of U. maritima plant grown fully exposed to the sun light for six months from leaf emergence showing morphological differences with the plant grown fully exposed to shade (Fig.101).

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Fig. 103. A digital camera photo (overview) of inflorescensce stalk of U. maritima containing more than 250 florets. Flower opening is in a manner of an open-flower zone continually moving toward the apex and differentiated into three flower stages: 1. Young (pre-anthesis) 2. Intermediate (at anthesis) 3. Old (post-anthesis). Scale bar = 3 cm.

Fig. 104. A digital camera photo (overview) of open flowers (the intermediate stage) showing the arrangement of the floral parts. The flower consists of six elongated white petals, six conspicuously long stamens with deep brown anthers and a central gynoecium with light green superior ovary. Scale bar = 0.5 cm.

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Figs 105-107. SEM micrographs of U. maritima ovary. Fig. 105. The outer groove of the septal slits (arrows) from where nectar drains. Scale bar = 1 mm.

Fig. 106. The septal slit is located between the fused carpels with stomata (arrows). Scale bar = 100 µm.

Fig. 107. Stomata occur on the outer surface of the ovary for aeration purpose. Scale bar = 50 µm.

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Figs 108-110. SEM micrographs of U. maritima perianth (stamen).

Fig. 108. Four microsporangia are attached to the tip of a peduncle filament. Scale bar = 1 mm

Fig. 109. Peduncle-like filament carries an anther. Scale bar = 1 mm

Fig. 110. Peculiar stomata present on the surface of the anther. Scale bar = 20 µm

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Figs. 111. LM micrograph of a transverse section of the first third from the top of U. maritima ovary showing the pathway (asters) through which pollen grains follow their way to the ovules. (Ep: epidermis; Id: idioblastic cell containg bundles of raphides; Cu: cuticle). Scale bar = 500 µm.

Fig. 112. LM micrograph of a transverse section at the end of the first third from the top of U. maritima ovary showing the upper side nectary tissues. The pathway (asters) through which pollen grains follow their way to the ovules is more obvious at this level. (Ny: nectary tissue; Id: idioblastic cells containing bundles of raphides). Scale bar = 500 µm.

Fig. 113. SEM micrograph of a transverse section of the ovary of U. maritima showing the location of the three gynopleural nectaries (arrows) alternating with the ovary locules. (O =ovules). Scale bar = 1mm.

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Figs 114-116. LM and TEM micrographs of a transverse section of young ovary of U. maritima.

Fig. 114. LM micrograph revealing the structure of the nectary after staining with TBO. The nectary consists of a compactly arranged epithelium (Ne), a subsidiary tissue (St) and subnectary parenchyma (Sn). Idioblastic cells containing bundles of raphides (Id) and the vascular bundles (Vb) are present in the subnectary parenchyma. Ep = epidermis. Scale bar = 50 µm.

Fig. 115. TEM micrograph showing the narrow nectary cavity (arrows) and the epithelium made up of cells with dense cytoplasm and relatively large nuclei. Scale bar = 10 µm.

Fig. 116. TEM micrograph showing an epithelium cell with ER (arrows) and discontinuing cuticle (arrowheads). Scale bar = 10 µm.

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Figs 117-119. TEM micrographs of a transverse section of the young ovary of U. maritima. Scale bar = 1 µm. Fig. 117. Epithelium cell with initial starch granules growing in the peripheral side of elongated plastid (Pl).

Fig. 118. Epithelium cell has well-developed starch granules (Sg) and Golgi bodies (G) at the end of the young stage.

Fig. 119. Subsidiary tissue possesses huge starch granules (Sg) at the end of the young stage. Scale bar = 1 µm.

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Fig. 120. TEM micrograph of a transverse section of the young ovary of U. maritima showing the epidermal cell covered with smooth cuticle (Cu). (Cw: cell wall) Scale bar = 50 µm.

Fig. 121. Detals of Fig. 120. The cuticle is more obvious at higher magnification. (G: Golgi body). Scale bar = 1 µm.

Fig. 122. LM micrograph of a transverse section of the young ovary of U. maritima showing the subsidiary tissue with red stained cells (arrows) containing polysaccharide content after employing the Schiff’s reagent. Scale bar = 500 µm.

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Figs 123-126. LM and TEM micrographs of a transverse section of ovary of U. maritima at the intermediate stage. Scale bar = 1 µm, except for Fig. 123, scale bar = 25 µm.

Fig. 123. LM micrograph showing a tubular structure (Ts) and the carpellary suture in the middle of the ovary.

Fig. 124. TEM micrograph showing the nectary epithelium with well-developed endoplasmic reticulum (ER) that occurs as long strands of parallel cisternae facing the outer walls. Plastids may contain plastoglobuli (arrows).

Fig. 125. TEM micrograph showing dense granular cytoplasm of the epithelial cells with plastids (Pl) which are almost devoid of starch granules at the stage of maximal development. The active endoplasmic reticulum (ER) with dilated cisternae (arrows) is also monenate the cytoplasm.

Fig. 126. TEM micrograph showing dense glandular cytoplasm of the epithelial cells with Golgi bodies (G) and mitochondria (Mt) at the stage of maximal development.

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Fig. 127. TEM micrograph of a transverse section of an ovary of U. maritima at the stage of maximal development showing the dense glandular cytoplasm of an epithelial cell with many, narrow and circular mitochondria. Scale bar = 10 µm.

Fig. 128. TEM micrograph of a transverse section of an ovary of U. maritima at the stage of maximal development showing the dense glandular cytoplasm of the epithelial cells with many secretory vesicles (arrowheads) and vacuoles containing electron dense material (arrows). Scale bar = 10 µm.

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Figs 129-132. LM and TEM micrographs of a transverse section of ovary of U. maritima at the intermediate stage. Scale bar = 10 µm, except for Fig. 131, scale bar = 50 µm.

Fig. 129. Abundant starch granules occurring in parenchyma cells near the nectary tissues during the secretion time (arrows).

Fig. 130. Abundant starch granules occurring in the peripheral parenchyma cells (arrows).

Fig. 131. LM micrograph showing vascular tissue consisting of xylem vessels (Xv) and sieve elements (SE) are occurring close to the placenta. Whereas, that occurs at the interior region of the nectary gland is consisting of xylem vessels (Xv) only.

Fig. 132. Sieve pores (Sp) occurring between the sieve elements, which contain plastids (Pl) with crystalloids.

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Fig. 133. LM micrograph of a transverse section of the first third of nectary from the bottom of the ovary of U. maritima at the intermediate stage showing a shorter and wider nectary than that of the two-third. Scale bar = 50 µm.

Fig. 134. LM micrograph of a transverse section of the middle of the mature ovary of U. maritima at the intermediate stage showing the more densely red stained idioblastic cells containing raphide bundles after employing the Schiff’s reagent (arrows). Starch granules have also reacted positively with this stain (arrowheads). (Ow: ovary wall). Scale bar = 50 µm.

Fig. 135. LM micrograph of a transverse section of the middle of the mature ovary of U. maritima at the intermediate stage showing the red stained nectar (nectar remnants) in the nectary cavity after employing the Schiff’s reagent (arrows). Starch granules have also reacted positively with this stain (arrowheads). Scale bar = 50 µm.

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Figs 136-140. LM and TEM micrographs of a transverse section of the old ovary of U. maritima.

Fig. 136. LM micrograph revealing epithelial (Ne) and subsidiary cells (St) more vacuolated than in the young stage. Some cells in the subsidiary tissue seem to degenerate (asterisks). Scale bar = 50 µm.

Fig. 137. Cell wall of a secretory cell possesses pores through which the nectar translocated into the cavity of the nectar (arrows). Scale bar = 10 µm.

Fig. 138. Pores located in the cell wall (arrows) of the secretory epithelium cell at the end of the secretion. Scale bar = 1 µm.

Fig. 139. Crystallized nectar (arrows) occurs inside the epithelial cell at the end of secretion. Scale bar = 10 µm.

Fig. 140. Shrinkage of the parenchyma cells and an idioblastic cell containing bundles of raphides at the end of secretion is obvious. Scale bar = 10 µm.

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Fig. 141. LM micrograph of a transverse section of the ovary of U. maritima at the intermediate stage showing four ovules (arrows) inside a carpel and the obturator gland (Ob). Scale bar = 100 µm.

Fig. 142. LM micrograph of a transverse section of the ovary of U. maritima at the late intermediate stage showing the well developed seed (Em: embryo) covered with the seed coat. Scale bar = 100 µm.

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Fig. 143. LM micrograph of a transverse section of the ovary of U. maritima at the old stage showing a well developed seed (embryo) covered with the seed coat. Idioblast cells react positively with Scheff’s reagent. Scale bar = 100 µm.

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Figs 144-147. TEM micrographs of a transverse section at the intermediate stage of the U. maritima ovary. Scale bar = 10 µm

Fig. 144. Epithelial cells of the obturator gland with many organelles, e.g. Golgi bodies, endoplasmic reticulum, ribosomes and plastids, during the active stage of mucilage secretion.

Fig. 145. The epithelial cells of the obturator gland have dilated endoplasmic reticulum (ER) and abundant vesicles (Vs) involved in mucilage production.

Fig. 146. Huge starch granules occur beside a mucilage cell (Mc) in the obturator gland.

Fig. 147. The presence of plasmodesmata (Pd) between the secretory cells of the obturator tissues is evident. (G: Golgi bodies) Scale bar = 10 µm.

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CHAPTER IV DISCUSSION 4.1 ADVENTITOUS ROOTS 4.1.1 Root system The nature of the root system varies in different kinds of plants. Plants, in which the primary root of the seedling eventually gives rise to most or all of the whole root system, are said to have a primary root system. Plants, in which most or all of the roots develop adventitiously from underground stems or from the base of aerial stems, are said to have an adventitious root system. Among plants with primary root systems, there are all variations from taproot system, with a central taproot that is larger than any of its branches, to a fibrous root system, in which the primary root is quickly deliquescent into several or many roots all about of the same size. Adventitious root systems are, usually, also fibrous, but both adventitious roots and taproots are sometimes thickened and fleshy, serving as storage organs. Most vascular cryptogams and all monocots, as well as a considerable number of dicots (especially those with creeping rhizomes) have an adventitious root system. Many dicots and most gymnosperms, on the other hand, have a primary root system of either the taproot or the fibrous type (Cronquist, 1982).

U. maritima has an adventitious root system in which all the roots develop adventitiously from the underground short stem called basal plate. A bulb of 10 cm diameter possesses 5- 10 adventitous roots. These could have approximately 5 branches. Moreover, the root system of U. maritima is, also, of the fibrous type (Carpenter, 1938). However, the adventitious roots are thickened and fleshy, serving as storage organs (Cronquist, 1982).

4.1.2 Special adaptation of roots The two principal functions of typical roots are anchorage and the absorption of water and minerals from the soil. A third important function of many, but not all, roots is the storage of food. Various types of specialized roots may have other functions. Roots require oxygen for respiration. That is why most plants cannot live in soil without an adequate drainage and consequently lacking air spaces. Some trees that grow in swampy habitats develop roots that grow out of the water and serve not only to anchor, but also to aerate the plant. Such an

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Sharaf Al-Tardeh Discussion adaptation was reported in the root system of the black mangrove Avicennia germinans which develops negatively geotropic extensions called pneumatophores (air roots). They grow upward out of the mud, thus providing adequate aeration (Raven et al., 1981). In the case of U. maritima, the root system develops a multi-layered epidermal structure, the velamen, which may, also, provide adequate aeration.

Aerial roots are adventitious roots produced from aboveground structures. Some aerial roots, such as those of the English ivy, cling to vertical surfaces, thus providing support for the climbing stem. In some plants, as in the case of corn, aerial roots serve as prop roots. Most roots are storage organs and in some species the roots are highly specialized for this function. Beets and carrots are examples of roots with an abundance of storage parenchyma (Curtis, 1983).

4.1.3 Velamen A characteristic feature of the U. maritima roots is the presence of the velamen. The latter provides osmotic and mechanical protection to the cortex and reduces water loss as well. It may, also, function in the absorption of water and minerals (Hew et al., 1993; Pridgeon, 1987; Benzing, 1989; Guttenberg, 1968; Fahn, 1990; Raven et al., 1981). During the dry period, the velamen cells are filled with air, while during the raining season they become filled with water acting as an absorptive tissue. Since the roots of U. maritima explore shallow soil horizons (10-30 cm in depth), they are likely to be vulnerable to dehydration in the upper soil profile. This is witnessed by a visible shrinkage of the older portions of the roots. This shrinkage probably reflects a hydraulic effect resulting from the water moving from the non-growing to the growing plant regions (Matyssek et al., 1991; Sawidis et al., 2005). The presence of a velamen depicts an adaptive strategy of the plants in the arid Mediterranean region, aiming to protect them from the loss of water which is valuable on account of the short seasonal rainfalls.

The velamen structure of some taxa is associated with mycorrhizae (Pridgeon and Chase, 1995). Some fungi play a crucial role in the mineral nutrition of the higher plants. Mycorrhizae (fungus-roots) complexes occur in most groups of vascular plants. Only a few

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Sharaf Al-Tardeh Discussion families of flowering plants characteristically lack mycorrhizae or form them very rarely; these include the mustard family (Brassicaceae) and the sedge family (Cyperaceae). There are two major types of mycorrhizae: a) endomycorrhizae, in which the fungal hyphae penetrate the cortical cells of the plant root, where they form coils, swellings, or minute branches, and b) ectomycorrhizae, in which the fungus surrounds but does not penetrate the living cells in the roots (Raven et al., 1981). In the case of U. maritima, nitrogen uptake by the roots might result from a mutual plant-fungus interaction. This hypothesis is supported by the presence of dead fungi in the velamen cells. Such an interaction of myccorhizae was, also, reported in other geophytes (Kauff et al., 2000; Tsavkelova et al., 2003 a & b).

There is close anatomical resemblance between the roots of U. maritima, Orchidaceae (Stern and Carlswad, 2006; Stern et al., 2004; Stern and Judd, 2001 & 2002) and those of Asphodelaceae (Asparagales) (Sawidis et al., 2005) in terms of the presence of velamen. A well-known example of a multi-layered root epidermis is the velamen radicum of perennating storage organs (air roots) of some Orchidaceae (terrestrial, epiphytic or mycoparasitic) and epiphytic Araceae species (Burr and Barthlott, 1991; Dahlgren and Clifford, 1982; Porembski and Barthlott, 1988; Pridgeon and Chase, 1995). The root velamen has also been reported in other families such as Cyperaceae and Velloziaceae (Porembski and Barthlott, 1995), as well as in some other plants from China (Yang-Tong et al., 2006).

4.1.4 Dimorphic hypodermis A characteristic feature of the U. maritima adventitious roots is the presence of a dimorphic hypodermis with regularly alternating long and short cells. Hypodermis is a layer of cells beneath the epidermis, which are distinct from the underlying cortical cells. In the mature roots the hypodermis would be a relatively distinct layer, but its developmental origin is still unknown. It could be interpreted as the outermost cortical (exodermal) layer, since in roots with a multiseriate exodermis the cells of the hypodermal layer more closely resemble the exodermal than the velamen cells. In cross sections of older roots the shorter cells are often recognizable by a more or less obvious thickening of their outer tangential wall. The

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The dimorphic hypodermis could provide an extra mechanical protection for the epidermis. It is present in the roots of almost all taxa of Asparagales and Araceae (Kauff et al., 2000). However, it is absent from most of the non-asparagoid monocotyledons and the Liliales in which U. maritima belongs to. Rudall et al. (1999) noted that in Hanguana malayana the rhizodermis (live cells producing root hairs) rather than the hypodermis has long and short cells (a dimorphic rhizodermis). This is also true of many other monocot taxa.

4.1.5 Cortex The cortical parenchyma of geophytes such as the Hyacinthaceae and Amaryllidaceae could be subdivided into two or three clearly distinct concentric regions, distinguishable by cell size (Wilson and Honey, 1966; Reynke and Van Der Schijff 1974; Wilson and Anderson, 1979; Ruzin, 1979; Jernstedt, 1984). However, in the case of U. maritima, the size of the cortical cells decreases gradually from the area close to velamen towards the endodermis. It has been reported that in some other plants (not geophytes), the cortex is subdivided into three layers as in Osmoglossum pulchellum (Stern and Carlsward, 2006).

Like other geophytes, the root cortex of U. maritima is mostly parenchymatous. It proved to be the basic storage region of the roots since it contributes to the total root thickness by 75%, while the central cylinder by 16.12%. Correspondingly, the cortical storage cells occupy about 34.11% of the total root volume. The storing character of the cortex in nutrients is the result of an adaptive strategy of U. maritima to survive the long summer drought of the Mediterranean climate.

4.1.6 The storage tissue According to the literature (Fernandez et al., 1972; Vega et al., 1972; Meletiou-Christou et al., 1992; Praznik and Spies, 1993; Kauff et al., 2000; Stern et al., 2004; Kamenetsky, et al., 1997; Sawidis et al., 2005; Stern and Carlsward, 2006), it seems that most of the storage tissues of the root of the plants occur in the cortex region. Specialized cells of storage

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The presence of specialized storage cells for polysaccharides, lipids and bundles of raphides in the roots of U. maritima is evident within the present study. Moreover, the presence of carbohydrates (Praznik and Spies, 1993), lipids, flavonoids (Fernandez et al., 1972) and anthocyanins (Vega et al., 1972) has also been reported. As it is mentioned above (see section 4.1.5 of this chapter), the primary storage tissue in the roots of U. maritima is the cortical cells. The root content of starch, lipids, proteins and soluble sugars varies considerably over the year. The amount of polysaccharides as well as total sugar contents, are always higher in the root than in the leaves. The highest values of soluble sugars appear in late spring-early summer like in other geophytes (Kamenetsky et al., 2005; Meletiou- Christou et al., 1992).

This is obviously an adaptive strategy for the plant to survive the dormancy stage in the hot and dry summer and to resist the environmental hazards and enemies. Furthermore, the plant exhibits not only quantitative but also qualitative differences in the bufadienolide (glycosides) composition according to its nativity (Krenn et al., 1994). Our morphometric evaluations (Tables 3.1, 3.2) revealed correspondences with the morphological and histochemical results.

4.1.7 Water economy The adventitious roots of U. maritima show a narrow fluctuation of total water content between the photosynthetic period (91.2 %) and the dormancy period (87.27%), which is an indication that these roots have developed efficient water-storage features. When values of water content in the upper part of the soil profile vary around zero, the roots remain hydrated and turgid with a total water content of about 87.27%. They are less susceptible to climatic stress and constitute a rather energetically stable system. This adaptation synchronizes the plant’s phenotypical development with the seasonal changes of the Mediterranean climate. This phenomenon has also been reported for other geophytes such as Asphodelus aestivus (Pantis, 1993; Sawidis et al., 2005).

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It is well-known that vascular organization can affect and/or limit the patterns of water translocation (Marshall, 1996; Marshall and Price, 1997; Price et al., 1996; Zhang et al., 2003). Moreover, vascular constraints are usually absent in many monocotyledons (Marshall, 1990; Pitelka and Ashmun, 1985; Liu et al., 2007) like U. maritima. The apoplastic pathway is the most probably followed by the water in roots and seems to be related to water economy in order for the plant to survive the very dry summer. At the endodermis, however, the water is forced to traverse the plasma membranes, through the plasmadesmata and protoplasts of the tightly packed endodermal cells. Therefore, the endodermis forms an osmotic barrier between the cortex and the vascular cylinder of the roots (Raven et al., 1981).

The water content is stored in water-storage cortical cells. Moreover, the velamen seems to be a feature depicting an adaptive strategy for the plant in order to reduce water loss during the summer period (dormancy). It may also function in the absorption of water and minerals available during the short raining season (Hew et al., 1993; Pridgeon, 1987; Benzing, 1989;

Guttenberg, 1968; Fahn, 1990; Raven et al., 1981).

4.1.8 Raphides Crystalline inclusions of different chemical composition and shape are found in many plants. Needle-shaped crystals are termed raphides (from the Greek word raphis = needle). Bundles of raphides, or needle-like crystal of calcium oxalate that occurring in adventitious root cells of U. maritima are typical of those found in many monocots, especially in geophytes (Esau, 1965; Kauff et al., 2000; Sawidis et al., 2005). Accumulation of oxalic acid in the tissues, which is not readily metabolized, may cause osmotic problems. Therefore, precipitation of calcium oxalate in the form of crystals, as a metabolic waste or by-product, seems to be an appropriate way for the plant to avoid these undesirable situations. The relationship between calcium-ion absorption and oxalic acid synthesis in plants is most probably established in order for the ionic balance in tissues to be maintained (Bosabalidis, 1987). On the other hand, calcium content in dormant roots may be viewed as an osmoregulatory adaptation to drought during the dry-warm summer period (Levitt, 1980;

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El Chonemy et al., 1978; Evans et al., 1992). Moreover, these cortical cells, which contain bundles of raphides are vertically oriented and located in rows around the central cylinder.

Raphides take part in both mechanical and chemical irritation when they come in contact with tender tissues of soil living worms and herbivores. Hence, raphides serve as a major defense mechanism against them (Ruiz et al., 2002). The mechanical irritation could be vital for the parenchymatous tissues of the roots of U. maritima. Therefore, defense strategies may also be viewed for other stored compounds (see section 4.1.6 of this chapter). Morphologically, the aciculate shape of these crystals is a critical component of the proposed defense mechanisms. In addition, twinning is an important factor in allowing plant cells to produce the raphide morphology (Arnott and Webb, 2000; Al-Tardeh et al., 2008a).

4.1.9 Vascular cylinder The vascular cylinder of the root consists of vascular tissues and one or more layers of cells, the pericycle, which completely surrounds the vascular tissues. The pericycle plays an important role in the formation of lateral roots (Raven et al., 1981). In U. maritima, a uniseriate pericycle is observed at the region of 3 cm above the root tip.

The vascular cylinder of U. maritima is typical of monocots regarding to xylem and phloem arrangement. It is polyarch (10-arch) with parenchymatous pith in young roots. The vessels differentiate into metaxylem which is more or less thick walled and lignified. Thickenings on the lateral walls of the vessels are helical or ring-shaped. Moreover, reticulated and scalariform or pitted thickenings have been reported in other geophytes (Wagner, 1977; Cheadle, 1969; Cheadle and Kosakai, 1971). Like other geophytes of Asparagales (Kauff et al., 2000), U. maritima bears anatomical features such as sclerenchymatous cylinder in the older roots, which are probably xeromorphic and developmentally convergent.

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4.2 BIOMASS AND RESERVE ALLOCATION PATTERNS WITHIN THE BULB 4.2.1 Morpho-anatomy of the bulb Perennials are plants in which the vegetative structures live year after year. The vegetative perennials pass unfavourable seasons as dormant underground roots, rhizomes, bulbs or tubers. However, the woody perennials, which include vines, shrubs and trees, survive above ground, but usually stop growing during the unfavourable seasons. Woody perennials flower only when they become adults, which may take many years. For example, Aesculus hippocastanum does not flower until it is about 25 years old, while, Puya raimondii, a large relative of the pineapple, takes about 150 years to flower. Many woody plants are deciduous, losing all their leaves at the same time and developing new leaves from buds, when the season again becomes favourable for growth (Raven et al., 1981). U. maritima is one of those belonging to this widely distributed group of plants, the perennials. It develops a perennial type in order to survive the unfavorable seasons. Moreover, U. maritima flowers after almost six years from germination (McCorohan 1990), when the bulb reaches a critical mass (Pascual-Villalobos and Fernandez 1999; Al-Tardeh et al., 2008a).

Plants of the Mediterranean ecosystems, as well as those from semiarid and arid regions are typically limited by water and/or nutrients rather than carbon (Pate and Dixon, 1982; Dixon et al., 1983; Pate and Dell, 1984; Bloom et al., 1985; Stock et al., 1987; Gutterman and Boeken, 1988; Boeken, 1990; Chapin et al., 1990; Witkowski, 1990; Boeken and Gutterman, 1991). Plants with underground storage organs most commonly occur in dry coastal areas of the Mediterranean and in the Irano-Turanian climate, e.g. Iran, Afghanistan, Central Asia, where a strong winter rainfall pattern is combined with periodic drought and soils of low nutrient status. Under these conditions a plant, e.g. U. maritima, is likely to have subterranean life forms to primarily avoid drought while it carries substantial fractions of its nutrient resources from one growing season to the next (Raunkiaer, 1907 & 1934; Pate and Dixon, 1982; Rees, 1981). Moreover, the development of a geophytic life form is an adaptive strategy for some plants to survive extreme cold environments (Wang, 2002 & 2005)

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The bulb of U. maritima represents a typical model of geophytes. Plant organs respond to environmental conditions and develop morpho-anatomical variations. These could be interpreted as adaptations to specific environments (Fahn and Cutler, 1992; Kamenetsky and Japarova, 1997). U. maritima is well adapted and synchronized to the Mediterranean climate by developing both underground storage organs (bulb) and subterranean life forms. Moreover, the results of the present study, also, indicate a higher underground biomass than that of the aboveground one. Anatomically, the storage cells occupy 18.31% to 50.66 % of the total volume of the bulb. Therefore, one could be led to the assumption (conclusion) that most of the bulb occurs as a storage tissue which, in turn, accounts for the meaning of the term ‘geophyte’.

4.2.2 Tunics The tunic of U. maritima is the bulb scale (leaf-base) that has been totally modified when the bulb matures. The tunic provides protection against disease infection and mechanical damage to the outer scales and the basal plate that contain the adventitious roots initials (De Hertogh and Le Nard, 1993). Moreover, a study on nitrate reductase activity of geophytes from the Mediterranean environment showed that the highest nitrate reductase activity occurs in the tunics of Allium scorodoprasum (Arslan and Güleryüz, 2005).

It is well known that when the tunic is extremely hard and unbroken, such as in the tulip, it can restrict root growth. Moreover, it could have a chemical effect as well. The removal of tunic enhances respiration and/or aeration of the unplanted bulb of Iris hollandica (Kamerbeek, 1962). In the case of U. maritima the stomata in tunics might play a synergic role in bulb aeration, especially during the storage and transportation of the unplanted bulbs. In addition, the presence of stomata supports the idea that tunic originates from the leaf bases.

4.2.3 Biomass, resources allocation and plant growth There is no relationship between the size of storage organs and species diversity. However, reliable winter rainfall makes large storage organs unnecessary and depresses extinction rates, thus leading to the accumulation of species (Proches et al., 2005). Like other perennial

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Sharaf Al-Tardeh Discussion geophytes (Ruiters et al., 1993 a & b), the bulb’s size and age are determining factors for the ability of reproductive development. In such geophytes with hysteranthous leaves, accumulation of storage reserves is a prerequisite for flowering (Rees, 1972; Dafni et al., 1981 a & b; Ruiters et al., 1993 a & b). The species possesses large underground biomass and/or reserve resources as required by the “reserve-pulse” model (Westoby, 1972; Noy- Meir, 1973). Changes of below and above-ground biomass allocation synchronize the plant’s phenotypical development with the seasonality of the Mediterranean climate (Pantis, 1993; Sawidis et al., 2005). Analogous variations in these adaptations have been reported for other perennial geophytes from Japan (Kawano, 1975; Kawano et al., 1982) and South Africa (Ruiters, 1995; Ruiters et al., 1993 a & b; Stock et al., 1992).

Plant species such as Asphodelus aestivus (Brouilet and Simon, 1979; Hume and Cavers, 1983; Pantis, 1993) and U. maritima exhibit resource allocation patterns that are the result of both their genotype and environment (Pfosser and Speta, 2004). In U. maritima, the bulb content of stored materials such as glycosides (cardiac glycosides) of the bufadienolide type (Kopp et al., 1996), anthocyanins (Vega et al., 1972), flavonoids (Fernandez et al., 1972), fatty acids, polysaccharides (Spies et al., 1992) and calcium oxalate (Cogne et al., 2001) varies considerably over the year (Miller et al., 1997; Pantis, 1993): the highest values were recorded in late spring just after leaf senescence. At this time, the plant possesses the highest mean percentage value of biomass (32 ± 5%; Table 3.4). The plant keeps its nutrients in situ in the bulb components, minimizing energy cost associated with synthesis, breakdown and translocation of storage compounds. This adaptation has also been reported for other geophytes (Rhoades and Cates, 1976; Westman and Rogers, 1977; Bloom et al., 1985; Chapin et al., 1990; Stock et al., 1992).

Urginea maritima exhibits three distinct phenophases, consisting of inflorescence, leaves and no above-ground biomass, respectively, during the annual life cycle. These phenophases can be linked to the seasonality of the Mediterranean type climate. The reproductive currency is also the stored resources in the constituent bulb parts. At the stage of leaf emergence the bulb biomass decreases (26 ± 6%) and decreases more at the mature leaf stage (20 ± 5%; Table 3.4). Like other geophytes (Pantis, 1987, 1993 & 1994), the

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Sharaf Al-Tardeh Discussion synchronization of the timing of U. maritima phenophases with the periodicity of the Mediterranean type climate is to be accomplished by means of biomass and nutrients trade- offs which take place within the different plant parts.

In geophytes, bulbs serve both as sinks and sources, and change carbohydrate composition as they grow and develop (Theron and Jacobs, 1996). Reserves are used for the development of new leaves (foliage and bases) and roots. Once the foliage becomes the photosynthate source, resources are stored in old and new leaf bases. Inflorescence becomes the major sink when elongation of the flower stalk is initiated (Theron and Jacobs, 1994). Thus, geophytes, particularly the hysteranthous ones are expected to possess great efficiency and flexibility of utilization of the compounds stored in their bulb (Dafni et al., 1981b; Boeken, 1990; Ruiters et al., 1993b). Therefore, plants might change their allocation of resources according to herbivore pressure.

Generally, carbohydrates are the major reserves within perennating organs. The two major functions of these reserves for growth strategy are: (1) supplying carbon and energy for the initiation of re-growth following seasonal dormancy; and (2) enabling plants to have some independence from the climatic periodicity of their habitat. Underground storage organs contain a variety of carbohydrates. The most common are starch, fructans, sucrose and glucomannans (De Hertogh and Le Nard, 1993; Miller, 1992; Miller et al., 1997; Risser and Cottam, 1968; Meier and Reid, 1982; Meyer and Hellwig, 1997; Pollock, 1986; Brocklebank and Hendrt, 1989; Orthen, 2001). Seasonal variations suggest that the polysaccharides are utilized for carbon and energy supply for re-growth and flower development (Orthen and Wehrmeyer, 2004). In addition, there is a possibility that some storage proteins act as a temporary store for nitrogen in several perennial species (Bewley, 2002; Al-Tardeh et al., 2008 b & c), and are reutilized to support plant growth (Cyr and Bewley, 1989, 1990).

From the morphometry point of view, the innermost bulb scale contains lower contents of lipids and polysaccharides than that of the middle. Presumably, the contents of the innermost bulb scales are consumed in order to supply the carbon and energy for re-growth

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Sharaf Al-Tardeh Discussion and/or development of the inflorescence buds. A similar result has been observed for bulbs of Lachenalia minima (Orthen, 2001). Different bulb scales of Lachenalia minima revealed that the major changes within the carbohydrate fraction occurred only within the innermost scales. Within this compartment the starch pool was depleted by 75%, whereas within middle and outermost scales 29 and 25%, respectively, were mobilized.

4.2.4 Histochemistry and the storage tissue From the histochemical point of view, mucilage of the cells containing bundles of raphides is polysaccharide, as indicated by Schiff’s reagent staining. The mucilage idioblasts associated with the vascular bundles and the epidermis, generally, have been regarded as water storage cells and are, thus, adaptively significant in warm habitats (Sawidis, 1991 & 1998). The mucilage has a substantial water-binding capacity due to its many hydroxyl groups and especially the carboxy groups of galacturonic acid (Goldstein and Nobel, 1991). Hygroscopic polysaccharides such as mucilage can bind 51 times their weight of water when hydrated in vivo (Nobel et al., 1992). Moreover, cell walls of the sclerenchyma fibers and the vessels are positively reacted with Schiff’s reagent.

On the other hand, the positively reacting cells with Sudan Black B and the oil cells are located around the phloem cells presumably to protect them from herbivores and insects. The stored protein may serve as donors of nitrogen in addition to that one provided by the root system. Moreover, nitrate assimilation can occur in both shoots and roots of most of the plants and the geophytic ones (Pate, 1973; Arslan and Güleryüz, 2005).

4.2.5 Defense mechanisms The tunic of U. maritima provides protection against disease infection and mechanical damage to the outer scales and the basal plate that contain the adventitious roots initials (De Hertogh and Le Nard, 1993). It is more likely to be the first line of defense for the plant. In parallel to that, the formation of the subterranean life form is also a contribution to this function protecting the plant from herbivores and environmental hazards.

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Calcium oxalate needles, usually, occur in cells of U. maritima adventitious roots (Al- Tardeh et al, 2006), leaves (Al-Tardeh et al, 2008b) and constituent bulb parts (Al-Tardeh et al, 2008a). The basal plate seems to be the active centre and needs extra protection which could be viewed by physical protection of raphide inclusions (Table 3.3, see section 4.1.8), the leaf-bases and the tunics. The polysaccharides content is strongly correlated with the raphides content (R = 0.78), which might give a synergic defense. Moreover, the production of calcium oxalate defenses mechanism has been observed to increase in cut bulbs of Pancratium sickenbergeri (Ruiz et al., 2002).

The stored compounds are not only used for developmental purposes, but also for the plant defense against herbivores, environmental hazards and stresses. The defense mechanisms by means of stored compounds act against microbial agents (Sathiyamoorthy et al., 1999; Hoffmann et al., 1993; March et al., 1991), herbivores (Blood and Radostits, 1989; Makhlouf, 1978), rodents (Verbiscar et al., 1986a; Fitzpatrick, 1952; Heth et al., 2000; Crouch et al., 2005), fungi (Miyakado et al., 1975; Singh and Rai, 2000; Samuel et al., 2000) and insects (Pascual-Villalobs, 2002; Pascual-Villalobs and Fernandez, 1999; Pascual-Villalobos and Robledo, 1999; Civelek and Weintraub, 2004).

Some authors have described the physiological requirements and production systems of the common flower bulbs under temperate-climate conditions (De Hertogh and Le Nard, 1993; Halevy, 1989). Because of their physiological traits, these bulbs are injured by drought and high growth temperatures. In addition, higher growth temperatures facilitate bacterial and viral infections, thus making bulb growing in warm regions more challenging. Moreover, Ruiz et al. (2002) found that Pancratium sickenbergeri has a complex resource allocation pattern as the result of combining defense and tolerance on herbivore activity. Plants respond to high levels of this kind of activity with a high re-growth capacity as a tolerant mechanism to maintain fitness. Scilliroside, the major toxic glycoside, occurs in all plant parts including the flowers, stalk, leaves, scales, and especially the roots and core of the bulbous part (Verbiscar et al., 1986b). This is in agreement with our results that the middle of the bulb contains the highest content of resources, especially the mucilaginous material and/or glycosides (cardiac glycosides) of the bufadienolide type (Kopp et al., 1996). The

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Sharaf Al-Tardeh Discussion bulb is not completely protected from herbivores and high temperatures at the soil surfaces because it is half buried in the soil. Correspondingly, the presence of a large number of glycosides (scilliroside) is considered as an adaptive strategy to protect the plant from herbivores and environmental hazards.

4.2.6 Water contents Evolution of geophytes in climatic areas with remarkable seasonal changes has led to their adaptation to periods of high and low temperatures and/or drought. In order to survive extreme environments, geophytes have undergone adaptations that may include increased capacity for water binding, tolerance of and/or resistance to desiccation and drought, and development of subterranean organs that contain specialized storage compounds (De Hertogh and Le Nard, 1993; Kamenetsky et al., 2003 & 2005).

One of the prominent features of the Mediterranean climate is its periodicity, to which U. maritima responds by synchronizing the annual development of its biological cycle (Al- Tardeh et al., 2006 & 2008a). As stated by Evans et al. (1992), survival during drought is ultimately dependent on the maintenance of the cell turgor. This species is adapted to dehydration as witnessed by a visible shrinkage of the cells of the older bulb-scales, which eventually alter into tunics. These cells are filled with water during the active stage and the water is reabsorbed and/or reused at the end of the summer dry season (dormancy). Moreover, the development of tunics seems to be an adaptive strategy to avoid water loss by reducing both the transpiration and overheating. In addition, the species proves to be very efficient in storing water during the long summer drought. When the values of the water content in the upper part of soil profile vary around zero, the bulb remains hydrated and turgid with water content higher than 67.5%. The growing flower bud might be a sink for this water, which is supplied from the basal plate and bulb-scales (Zemah et al., 1999; Kamenestsky et al., 2003), since the leaf spouting depends on the external water supply (Al- Tardeh et al., 2008b; Dafni et al., 1981a; Boeken, 1990).

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4.3 MORPHO-ANATOMICAL FEATURES OF THE LEAVES 4.3.1 Leaf anatomy Leaves are the characteristic photosynthetic organs of most vascular plants. They are appendages of the stem to which are attached by a distinct stalk, the petiole. Large proportions of monocots have sessile leaves (without petiole) with a relatively broad, more or less sheathing base that extends most or all of the way around the stem. This sheathing base is developed into a well-defined leaf sheath that encloses the stem for some distance above the node (Cronguist, 1982).

4.3.2 Epidermis and cuticle The Mediterranean-type climate is characterized by hot and dry summers alternating with cold and wet winters (Daget, 1977; Nahal, 1981). Deciduous species in the dry regions possess leaves that undergo senescence and desiccation during the period of dryness (Orshan, 1963). The seasonal fluctuations in soil moisture are considered as one of the limiting factors for growth and productivity of the Mediterranean perennial species (Mitrakos, 1980; Specht, 1987).

Restriction of water loss to a minimum is of decisive importance under conditions of severe drought. This is mainly achieved by the coverage of leaf surfaces with the cuticle (Schönherr, 1982). Moreover, the thick cuticle increases leaf reflectance, thus reducing solar inception, heat load and therefore water deficit (Gausman and Quisenberry, 1990). In addition, the cuticle is considered as the outermost defensive barrier of the plant leaves against pathogens and environmental hazards (Stenglein et al., 2005). The cuticle is, usually, thicker on the upper epidermis than on the lower one, and thicker on plants grown under the sun light than those in the shade (Cronguist, 1982). This is in agreement with our results that the upper cuticle is slightly thicker than that of the lower one (Table 3.5).

4.3.3 Stomata as an adaptive strategy The only intercellular spaces in the epidermis of most leaves are the stomates (from Greek word stoma, a mouth). The term stoma applies only to the small pore (space) between the guard cells. The latter are the only epidermal cell containing chloroplasts. The stomata may

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Sharaf Al-Tardeh Discussion be equally abundant on both sides of the leaf, but usually they are more numerous on the lower surface. Stomata of xerophytes (plants adapted to dry conditions) are often individually sunken or grouped in sunken stomatal pockets. The stomatal opening is very small; commonly 7-40 µm long and 3-12 µm wide when it is fully-open. The fully open stomata, usually, cover about 0.5-2% of the lower epidermis. The role of stomata is to control transpiration (Cronguist, 1982). In the case of U. maritima, the guard cell pairs have a mean length of 45.4 ± 2.3 µm and 22.8 ± 2.1 µm width, as determined from SEM micrographs, and possess ledges.

The size and density of the stomata have been widely studied and related to many environmental factors (Salibury, 1927; Shields, 1950; Klich et al., 1996 a & b). Small stomata in large numbers seemed to be characteristics of xeromorphic leaves (Shields, 1950). However, Meidner and Mamsfield (1968) found a great stomatal frequency variation among mesophytes, so that the efficiency of stomata in regulating water loss could not be directly related to their size and frequency (Klich, 2000).

Amphistomaty and leaf compartmentalization have been repeatedly evaluated concerning leaf xeromorphy. Amphistomaty, which is more common in xeric habitats (Parkhurst, 1978;

Fahn and Cutler, 1992), shortens the distance of CO2 diffusion to mesophyll cells (Parkhurst et al., 1988; Terashima et al., 2005). Small but abundant stomata are also believed to lower

CO2 diffusion resistance toward the photosynthesizing tissue. Thus, non-succulent species show increased stomatal density (Sundberg, 1986) as is the case of U. maritima.

4.3.4 The mesophyll ground tissues From the anatomical point of view, the mesophyll of a leaf usually consists largely or completely of chlorenchyma, that is a chloroplast-bearing parenchyma. Typically it is rather distinctly differentiated into an upper layer, the palisade parenchyma (palisade chlorenchyma, palisade mesophyll) and a lower layer, the spongy parenchyma (spongy chlorenchyma, spongy mesophyll). The mesophyll is structurally well adapted to the primary function of leaves, which is photosynthesis. The more compact tissue and the greatest concentration of chloroplasts are toward the upper surface, which receives most of

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Sharaf Al-Tardeh Discussion the light. The intercellular spaces are greater close to the stomata, which are the gateways to the outer air, and diminish progressively toward the upper surface (Cronguist, 1982). U. maritima possesses large petiole-less, unbendable leaves with a permanent position in respect to solar radiation. This position is at 40-45º inclination from the horizontal one. This in turn allows the light to reach the leaf from both sides. Therefore, the presence of spongy parenchyma in both upper and lower mesophyll is ensconced as an adaptive strategy for optimum light energy exploitation and higher possible photosynthetic efficiency. However, the upper mesophyll cells are more elongated, but the intercellular spaces are the same on both sides of the mesophyll. Therefore, it could be concluded that the leaf of the studied species is equifacial.

Abundant stomata on both leaf surfaces and the absence of any kind of compartmentalization, as well as the fine structure of spongy parenchyma, i.e., cell arrangement and chloroplast distribution along the cell walls facing intercellular spaces, may decrease CO2 diffusion resistance (Parkhurst et al., 1988; Psaras et al., 1996; Evans and

Loretto, 2000). Abundant palisade parenchyma is also believed to increase the CO2 absorbing surface of the mesophyll (Rhizopoulou and Psaras, 2003; Terashima et al., 2005). Moreover, mesophyll intercellular spaces and air spaces occupy 30.4% of the total leaf volume of U. maritima. Indeed, one third of the leaf’s volume is reserved for gas exchange, which in turn increases the photosynthetic rate. Before leaf senescence the nutrients are translocated to the storage organs of the plant (Al-Tardeh et al., 2008b).

4.3.5 The vascular system In most monocots the leaf has several to many almost parallel veins, which run the length of the leaf, and very small inconspicuous branches interconnecting these veins. Monocots, as well as U. maritima, are therefore usually said to have parallel venation. A vein, typically, consists of a strand of xylem and a strand of phloem, collectively surrounded by a bundle sheath. The phloem is usually on the lower (abaxial) side and the xylem on the upper (adaxial). The bundle sheath is ontogenetically part of the mesophyll, but it is usually discussed with the vascular system because of its close morphological and physiological connection to that. The bundle sheath of U. maritima consists of a single layer of elongate,

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Sharaf Al-Tardeh Discussion thin-walled, living cells, completely surrounding the vein. The bundle sheath’s principal function is to facilitate the conduction of water, as well as of solutes from the vein to the mesophyll, and also of solutes from the mesophyll to the veins (Cronguist, 1982).

The bundle sheath of U. maritima is indiscernible and does not possess either chloroplasts or bundle sheath extensions. The absence or the presence of very few chloroplasts inside the bundle sheath cells and the frequent occurrence of chloroplasts inside mesophyll cells are a pronounced C3 plant feature. Wang (2005) found that the higher abundance of C3 plant species than that of C4 is consistent with moist climates (850 mm rainfall). Therefore, C3 species, such as U. maritima, tend to be distributed in moist and low temperature conditions, which are a characteristic of the Mediterranean climate in winter.

4.3.6 Leaf morphology Leaves are the plant organs mostly exposed to aerial conditions and changes in their characteristics have been interpreted as an adaptation to specific environments (Fahn and Cutler, 1992). Reduced leaf size, increased thickness, thick external walls of the epidermal cells, high stomatal density and palisade parenchyma developed at the expense of spongy parenchyma are common features of plants grown in xeric environments (Fahn and Cutler, 1992; Shields, 1950). The leaf of U. maritima is thicker compared to that of evergreen sclerophylls (714.2 ± 33.6 μm vs. 250–550 μm, (Christodoulakis and Mitrakos, 1987; Rotondi et al., 2003) and of semi-deciduous species (714.2 ± 33.6 μm vs. 100–150 μm). Thus, relatively thick leaves of U. maritima possess certain structural characteristics of xeromorphic leaves, considered to facilitate CO2 uptake by the mesophyll (Terashima et al., 2005). However, U. maritima leaf could account for mesomorphic leaves, since they possess flat lamina (Strasburger et al., 1982).

Many traits from leaf structure can be explained as adaptations to enhance CO2 diffusion within the leaf for photosynthesis (Parkhurst, 1986). A morphological trait of the leaf that correlates with CO2 assimilation is the specific leaf mass (SLM) (Mooney et al., 1978; Field and Mooney, 1983; Ellsworth and Reich, 1992). SLM is considered as a useful index of xeromorphism (Witkowski and Lamont, 1991). Species with low SLM tend to have a higher

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Sharaf Al-Tardeh Discussion photosynthetic capacity per unit leaf mass, resulting from a larger light-capture area per leaf mass (Wright et al., 2001), and a shorter diffusion path from stomata to chloroplasts (Parkhurst, 1994). SLM of the leaf of U. maritima (8.564 ± 3.139 mg cm-2) is lower than that of evergreen sclerophylls (14-20 mg cm-2) (Gratani and Bombelli, 2001) and of drought semi-deciduous species (8.5-14.7 mg cm-2) (Gratani and Bombelli, 2000), while, it is somewhat, higher than that of the non woody perennial species (6-9 mg cm-2) (Yadav et al., 2004) and of the geophyte Pulmonaria officinalis (4.4 mg cm-2) (Gaberščik et al., 2001). Moreover, Cowling and Campbell (1983) proposed to consider the specific leaf mass (SLM) value of 7.0 mg cm-2 as the borderline between malacophylls and sclerophulls. On the basis of this value, U. maritima with a value of 8.564 ± 3.139 mg cm-2 is very close to the borderline. Therefore, SLM of the leaf of U. maritima could account for considering the plant to be adapted both as a drought deciduous and mesophytic species (plant that prefers to live in a moderate supply of water. The word is derived from the Greek words meso = medium, and phyton = plant).

LTD of the leaf of U. maritima is lower than that of evergreen sclerophylls (416-669 mg cm- 3) (Gratani and Bombelli, 2001), semi-deciduous species (610-690 mg cm-3) (Gratani and Bombelli, 2000) and of that of evergreen shrub species (504-756 mg cm-3) (Gratani and

Varone, 2004). The lower LTD allows a better CO2 movement through the air spaces between the cells (Parkhurst, 1986), resulting in a higher photosynthetic rate during the favourable period. On the contrary, leaves with a higher amount of biomass per unit area may be more efficient in water use during drought. The leaf of U. maritima is adapted to such conditions by possessing low LTD and a moderate SLM for a better CO2 movement (Gratani and Varone, 2004). It also has thick cuticle, which increase leaf reflectance, thus reducing solar inception, heat load and therefore water deficit (Gausman and Quisenberry,

1990). However, the latter may disturb the CO2 movement through the stomata; this undesired condition may be compensated by possessing high density of stomata on both sides of the leaf which reduces the diffused distances of CO2.

Like other bulbous plants (Evenari and Gutterman, 1985; Gutterman and Boeken, 1988; Boeken, 1989; Kamenetsky, 1994), U. maritima lacks specific xerophytic adaptations such

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Sharaf Al-Tardeh Discussion as sclerophyllous or succulent leaves or special physiological mechanisms to cope with drought. As Fahn (1982) stated, xeromorphism is not confined to xerophytes, while not all xerophytes exhibit xeromorphic features. Therefore, elucidating and understanding the leaf structure of U. maritima under natural conditions will be useful for further studies dealing with the physiology of this species.

4.3.7 Leaf histochemistry The accumulation of water-soluble carbohydrate is evident in the leaf of U. maritima. Polysaccharides of the idioblastic cells could be fructans (sinistrin) and fructo- oligosaccharides (Spies et al., 1992). They resemble the branched structure of fructans isolated from excised leaves of Phormium tenax (Sims et al., 2001) and from other related species within the order Asparagales, such as Agave vera cruz (Dorland et al., 1977; Allium cepa (onion; Henry and Darbyshire, 1980), Asparagus officinalis (asparagus; Shiomi, 1993), and Cordyline australis (New Zealand cabbage tree; Brasch et al., 1988). These polysaccharides might have a function as water binding molecules as it has been discussed in section 2.4 of this chapter and/or contribute to the accumulated reserves in the belowground organs.

The lipid contents of the leaf were stained brown to black with Sudan Black B. These contents might be the phenolic compounds which usually occur in terrestrial plant tissues (Dakora, 1995). Among them particular attention has been given to their possible function as selective filters against ultraviolet-B (UV-B) radiation damage (Caldwell et al, 1983). Phenolics such as flavonoids are absorbed strongly in the UV, but not in the visible region of the spectrum, and their biosynthesis is accelerated by UV-B radiation (Beggs and Wellman, 1994). Indeed, as it has been repeatedly shown, that light availability and the whole leaf phenolic content are positively correlated (Stephanou and Manetas, 1997).

The presence of the UV-B-absorbing components (Grammatikopoulos et al., 1999) in the leaf of U. maritima is evident as flavonoids (Fernandez et al., 1972). Grammatikopoulos et al. (1999) stated that the concentrations of the UV-B-absorbing components of the whole leaf or its epidermis fluctuate according to the site-dependent radiation stress. Therefore,

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Sharaf Al-Tardeh Discussion high irradiance in U. maritima, apart from inducing an increase in UV-B-absorbing compounds on a whole leaf basis, also causes a change in the distribution of these compounds between epidermis and mesophyll. Similar results have been recorded for other geophytes from Europe (Gaberščik et al., 2001).

Phenolic compounds are suggested to protect cells from structural damages (Tevini, 1994) by absorbing UV-B radiation (Karabourniotis et al., 1992). These essential oils may be efficient in reducing both transpiration and overheating (Todorović and Stevanović, 1994). Moreover, they might act as allelochemicals (allelochemicals mediating plant-plant, plant- animal, plant-microbe interactions) in order to protect the plant form herbivores. Further work is still needed to examine this hypothesis.

4.3.8 Photosynthetic efficiency of the leaf of U. maritima. 4.3.8.1 Chlorophyll a content After leaf chlorophyll examination of plants grown in shade and others grown under the sun light, the results suggest that chlorophyll contents are affected slightly by solar radiation. However, in U. maritima and another plant (Gratani and Ghia, 2002), chlorophyll a is more concentrated than chlorophyll b. Chlorophyll and carotenoid contents in the leaves of the geophyte Pulmonaria officinalis do not vary significantly among different UV-B radiation (Gaberščik et al., 2001). However, it has been reported that in some cases UV-B radiation can stimulate chlorophyll and carotenoid synthesis (Rau and Schrott, 1987)

Popovič et al. (1996) found a strong positive correlation between a decrease in the chlorophyll a/b ratio and a decrease in irradiance (gradually from “light” to “dark” forest phase) in the geophyte Arum maculatum L., whereas the chlorophyll a/b ratio in the geophyte Scilla bifolia L. leaves did not significantly differ over the same period. This phenomenon of chlorophyll modification and decreased photosynthesis, accompanied by a large amount of leaf area developing in the shade, indicates greater photosynthetic plasticity of A. maculatum relative to S. bifolia (Popovič et al., 2006). This is, also, in agreement with our results that the chlorophyll a/b ratio decreases in the leaves of plants shifted from sun light to shade.

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The relationship between leaf photosynthesis and growth is more complex in perennials with below storage organs such as U. maritima. The considerably greater leaf area in A. maculatum does not lead to a higher growth rate at the whole plant level, but enables a longer photosynthetically active period and consequently, a longer accumulative period. This is the pattern whereby dry matter partitioned into leaves modifies leaf area index, with consequent strong positive feedback to total biomass production (Heuvelnik, 1999).

4.3.8.2 The basis of chlorophyll fluorescence measurements Chlorophyll fluorescence is used as a sensitive probe for studying the photosynthetic ability of plants. Therefore the Fv/Fm ratio is a measure of PSII photochemical efficiency, and it is a sensitive indicator of photoinhibition and other stresses (Araus and Hogan, 1994). It is known that in healthy plants its value is equal to 0.83 in most plant species (Björkman and Demming, 1987; Johnson et al., 1993). Values lower than 0.83 are observed for plants under distress factors and in association with changes in the value of Fo parameter, which become reliable indices of the photo-inhibition phenomenon (He et al., 1996; Maxwell and Johnson, 2000). In the case of U. maritima, the maximal efficiency of PSII photochemistry measured as Fv/Fm was decreased in light leaf (0.824 ± 0.008) versus that in the shade (0.836 ± 0.030). More specifically, plants grown in shadow showed more photosynthetic ability than those grown under sun light. Moreover, U. maritima seems to be adapted to high solar radiation by inducing photo-inhibition factors (He et al., 1996; Maxwell and Johnson, 2000) and/or an increase in UV-B-absorbing compounds (Grammatikopoulos et al., 1999). On the contrary, plants grown in shade seem to be adapted to low solar radiation by increasing the efficiency of PSII photochemistry (Fv/Fm; 0.836 ± 0.030).

The total quantum yield of PSII photochemistry was measured (Genty et al., 1989). It informs about the light analogy absorbed by chlorophyll molecules in relation to PSII which is used in photochemical processes. Both of the shade and light leaves show high activity and/or absorbance of photons at the low photon flux intensity. When the photon flux intensity increases more than 536 μmol photons m-2 s-1, the yield decreases. Moreover, the shade leaf shows higher quantum yield than the lighte leaf at the PAR range from 66 to 536 μmol photons m-2 s-1 and lower quantum yield at the higher PAR intensities. Thus, U.

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Sharaf Al-Tardeh Discussion maritima leaf seems to show adaptation to fluctuating environmental conditions (light intensities)

Electron transport rate (ETR) represents also the apparent rhythm of electron transport through PSII measured in μmol electron m-s s-1. In the case of U. maritima, the photon flux intensity of 536 μmol photons m-2 s-1 seems to be the point of altering the efficiency of the PSII between the shaded and lighted leaves. This also corresponds to the parameter yield.

Non-photochemical quenching (qN) parameter reflects the energy released in the form of heat and its values range between 0 to 1 (Krause and Weis, 1991). U. maritima light leaf shows higher qN than that of the shade leaf when exposed to photon flux intensities of 66 to 536 μmol photons m-2 s-1. However, at higher photon flux intensity, qN of both leaves are overlapped. All the values of qN at various photon flux intensities are in the range of 0 to 1 except for those at 1911 and 3111 μmol photons m-2 s-1, where the qN values are higher than 1. In conclusion, the leaf of U. maritima shows a very “induced fit” to the Mediterranean- type climate.

Morphological variations were also recorded between plants grown fully exposed to sun light and those fully exposed to the shade. The shaded plants possess more elastic, weak and thin leaves. Moreover, the leaves of the shade plants are longer than those of the sun lighted plants (Figs 101 & 102). Thick leaves with a greater area of chloroplast surfaces facing the intercellular spaces would be advantageous because chloroplasts can be thin and the amount of Rubisco per unit area of chloroplast surface can be small. On the other hand, with an increase in leaf thickness, the bulk resistance to CO2 diffusion in the gas phase increases, which causes a decrease in bulk intercellular spaces. In this respect, thick leaves are disadvantageous (Terashima et al., 2001).

As it has been mentioned (in section 3.2 of this chapter), species with low SLM tend to have a higher photosynthetic capacity per unit leaf mass, resulting from a larger light-capture area per leaf mass (Wright et al., 2001), and a shorter diffusion path from stomata to chloroplasts (Parkhurst, 1994). This explains the thin lamina and the higher Fv/Fm ratio for the shadow

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Sharaf Al-Tardeh Discussion plants. Moreover, Cowling and Campbell (1983) proposed to consider the specific leaf mass (SLM) value of 7.0 mg cm-2 as the borderline between malacophylls and sclerophylls. On the basis of this value, U. maritima grown under sun light with a value of 8.664 ± 0.447 mg cm-2 is very close to the borderline. While those grown in the shade with a value of 5 ± 0.1 mg cm-2 are lower than the borderline. The values of the SLM of the leaf grown in the sun and the natural habibta are close to each other because the chosen plants in natural habitat were almost exposure to the sun. SLM of the leaf of U. maritima in all cases is lower than and/or close to the borderline. Therefore, considering the plant to be adapted as both a drought deciduous and mesophytic species is strongly recommended.

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4.4 MORPHO-ANATOMY OF THE INFLORESCENCE 4.4.1 Gynopleural (septal) nectary 4.4.1.1 Nectary epithelial cells A topographical classification of floral nectaries indicates nine different types (Fahn, 1979). Among them, the “ovarial nectary” type includes nectaries that are placed in the septal region between adjacent carpels, the so-called septal nectaries or gynopleural nectaries as they have been more recently defined by Smets and Cresens (1988). Therefore, the gynopleural nectary, being a cavity inside the ovary, is not directly exposed to nectar- feeding animals and the site of nectar emission is often different from the site of nectar production (Smets et al., 2000). Gynopleural nectaries are restricted to monocotyledons, where they represent the most common type of floral nectary (Smets et al., 2000). In the flower of U. maritima a typical gynopleural (septal) nectary was found.

In U. maritima nectary, the nectary cavity is lined by a layer of secretory epithelial cells that are covered by a cuticle, as well as other plants (Fahn and Benouaiche, 1979; Nepi et al., 2005). A continuous cuticle is generally present on the surface of the nectary epidermis, although it may be thinner than that on the areas adjacent to the nectary (Gaffal et al., 1998) or discontinuous as in septal nectaries. Complex cuticle organization with a lamellar-type outer layer and a reticulate-type inner one has been described in the floral nectary of Aptenia cordifolia (Aizoaceae) and Limodorum abortivum (Orchidaceae) (Meyberg and Kristen, 1981; Figueiredo and Pais, 1992).

The patterning, thickness and permeability of the nectary cuticle vary widely. In the case of nectary trichomes, as in Hibiscus rosa-sinensis (Malvaceae) the cuticle covering the secreting cell seems to be completely impermeable and the nectar accumulates in a subcuticular space formed by the separation of the cuticle from the epidermis (Sawidis et al., 1987b). The continuing build up of pressure within this compartment ultimately reaches the level where pores in the cuticle over the tip cell become patent (Spreading open or expanded) and release a pulse of nectar to the exterior (Robards and Stark, 1988). As secretion proceeds, the cuticle stretches and becomes very thin. It has not been determined whether the nectar is released when the cuticle breaks or whether thin areas of the stretched

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Sharaf Al-Tardeh Discussion cuticle become permeable to nectar. However, a continuous thick cuticle covers the epidermal cells of the extrafloral nectaries and nectar release generally takes place through cuticle rupture.

The cell wall and the cuticle may have microchannels from which the nectar exudes (Davis et al., 1988; Stpiczyńska, 2003). In Platanthera chlorantha (Orchidaceae), the microchannels appear as fibrillar outgrowths of the outer epidermal cell wall, as also observed in Abutilon sp. (Kronestedt-Robards et al., 1986). In the case of U. maritima and Helleborus foetidus (Ranunculaceae) (Koteyeva, 2005), microchannels are narrow tubular in continuity with the cell wall; some of them seem to have direct communication with the outside. Very similar microchannels are described in the cuticle of the epidermal cells of the Echinacea purpurea (Asteraceae) nectary, although they have no direct communication with the outside (Wist and Davis, 2006).

4.4.1.2 Subsidiary tissue Different extents of the subsidiary tissue were observed in various species of Tillandsia (Cecchi-Fiordi and Palandri, 1982) and were related to nectar production rates (Nepi et al., 2006). The nectary is supplied by a prominent vascular bundle comprising phloem and xylem elements. Most studies concur that both sugar and water components of the nectar are, generally, provided by phloem sap and this is transported from the sieve elements to the nectary cells (Fahn, 2000; Pacini et al., 2003; de la Barrera and Nobel, 2004; Stpiczyňska et al., 2005). Pre-nectar unloading is favoured by phloem companion cells that often have wall ingrowths of the transfer cell type as observed in Vicia faba, Pisum sativum (Fabaceae) (Davis et al., 1988; Razem and Davis, 1999), and Eccremocarpus scaber (Bignoniaceae) (Belmonte et al., 1994).

In the case of U. maritima, water and sugars are most probably transported from the bulb via the vascular system and are stored in the nectary cells (epithelial, subsidiary and parenchyma cells) as starch during the young stage. A symplastic route dependent on intercellular plasmodesmatal connection is evident in the gynopleural nectary of U. maritima, from the sieve elements to the epithelium nectary. Plasmodesmata connect sieve 195

Sharaf Al-Tardeh Discussion elements to companion cells and sieve element-parenchyma cells. Moreover, plasmodesmata connect also the subsidiary cells to the epithelial ones of the nectary. However, the apoplastic transfer of the nectar into the nectary cavity may be facilitated by microchannels in the cuticle (Fig. 138) lining the outer nectary surface, similar to other nectaries (Radice and Galati, 2003).

4.4.1.3 Amyloplasts and nectar secretion The nectary parenchyma may have amyloplasts or chloroplasts containing only a few stacks of grana and small starch granules. Nectaries with amyloplasts in the nectary and subnectary parenchyma cells seem less common than those with chloroplasts. In addition, all extrafloral nectaries have chloroplasts in their parenchyma (Pacini et al., 2003). Chloroplasts with grana stacks and plastoglobules may contain small or large starch grains (Razem and Davis, 1999; Baum et al., 2001; Horner et al., 2003) or may even be empty (Stpiczyńska and Matusiewicz, 2001; Stpiczyńska et al., 2003). Nevertheless, the precise approach of either the nectar sucrose or other carbohydrates derived from chloroplast photosynthesis is still undefined.

The environment may affect nectar production irrespectively of the plastid type in the nectary parenchyma (chloroplasts without starch, chloroplasts with starch grains, amyloplasts), but the effects of environmental conditions act at different times. If nectary parenchyma cells have chloroplasts, nectar production is affected by immediate environmental conditions, whereas it is affected by the environmental conditions of previous days when nectary parenchyma cells contain amyloplasts.

On the other hand, an advantage of nectaries with amyloplasts in the nectary parenchyma cells is that nectar may be available for consumers, and in large quantities, at any time of the day or night. The nectar may also have a high sugar concentration and can be produced in a short time. In the case of Cucurbita pepo L. (Nepi et al., 1996 a & b) for example, nectar becomes available from 6 a.m. and has a high sugar concentration (30–40%). This high rate

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Sharaf Al-Tardeh Discussion of nectar production is impossible with nectary parenchyma cells containing chloroplasts. In C. pepo, nectar is secreted for only 3-4 h but the nectary does not immediately degenerate. Empty amyloplasts may be involved in temporary storage of reabsorbed carbohydrates if the nectar is not totally consumed by flower visitors (Nepi et al., 1996 a & b). Unconsumed nectar carbohydrates are temporarily polymerized to spherical electron dense bodies which react positively to the PAS test for total insoluble polysaccharides (Nepi et al., 1996a). Afterwards these plastids empty and the carbohydrates are probably totally reabsorbed by contiguous parts of the flowers. The nectary then abscisses. Similar behavior has been recorded for U. maritima; a high and wide range of sugar concentrated nectar (10-74%) is secreted only for 3-4 hours. Nectar ruminants inside the nectary cavity react positively with PAS reaction.

This pattern of nectary plastid development (from proplastids to amyloplasts) could be more common in the tropics, where more animals are active at night, than in temperate zones. Degeneration of nectaries with amyloplasts at the end of secretion or after nectar reabsorption seems to be a general feature. In Aloe and Gasteria, which have septal nectaries, dedifferentiation of amyloplasts to chloroplasts is recorded (Schnepf and Pross, 1976; Nepi et al., 2006). This dedifferentiation enables transformation of nectary parenchyma into fruit parenchyma. Floral nectaries differ widely according to the dynamics of nectar production and plastid differentiation patterns, because they are visited by a wider spectrum of consumers and may reabsorb unconsumed nectar

It is generally accepted that the nectar originates from the phloem sap (Fahn, 2000; Gaffal et al., 2007). The sugar-secreting nectaries are undisputedly sink organs dependent on source tissue assimilates. Most of them, as well as U. maritima, are vascularized exclusively by phloem strands (Frei, 1955). The efflux of carbohydrates may occur across the plasma membranes of the sieve elements-companion cell complexes (apoplastic pathway) or through plasmodesmata interlinking these complexes with adjacent cells (symplastic pathway; Patrick et al., 2001).

Undifferentiated plastids (proplastids) are present in the very early stages of nectary 197

Sharaf Al-Tardeh Discussion development (Nepi et al., 1996a). Close to flower anthesis, they may differentiate to chloro- amyloplasts (Pacini et al., 1992; Nepi et al., 1996a). In some cases, chloro-amyloplasts lose their thylakoid structure and starch grains increase in size a few days before anthesis (Zer and Fahn, 1992; Fahn and Shimony, 2001). In other cases, proplastids differentiate into amyloplasts and store great amounts of starch in many large grains per plastid before nectar secretion begins (Durkee et al., 1981; Pais and Figueiredo, 1994; Nepi et al., 1996a). Amyloplasts in the nectar-producing parenchyma are generally almost devoid of stroma and packed with starch (Nepi et al. 1996a). They also contain many starch grains per amyloplast; this increases starch surface area, facilitating and speeding up hydrolysis during nectar production. Thus many authors infer that hydrolysis of starch in the parenchyma contributes directly to nectar carbohydrate content (Durkee et al., 1981; Nepi et al., 1996b; Pacini et al., 2003; Peng et al., 2004). Like other plants with septal nectaries (Schnepf and Pross, 1976; Nepi et al., 2006a), dedifferentiation of amyloplasts to chloroplasts mostly occurs after nectar secretion in U. maritima nectary.

Starch stored within amyloplasts at the young (pre-secretory) stage can be utilized both as a source of energy for highly metabolic processes and as a source of sugars for nectar synthesis (Stpiczyňska et al., 2003). During successive stages of secretory activity, plastids generally become elongated or develop an irregular profile and this is usually associated with depletion in starch content (Sawidis, 1988; Sawidis et al., 1989; Sawidis et al., 1998; Horner et al., 2007). Degradation of plastids (amyloplasts) starts with hydrolysis of starch grains (Gaffal et al., 2007). The engulfing pattern of starch hydrolysis occurs in the floral nectaries of Eccremocarpus saber (Belmonte et al. 1994), Cucumis sativus (Peng et al., 2004), senescing French bean leaves (Minamikawa et al., 2001) and U. maritima. The engulfing hydrolysis depends on the open access of vacuolar sap to the interior of the amyloplast, engulfing of the plastids by vacuole and subsequent hydrolysis of starch in the vacuole is processed.

The correlation between nectar production and starch hydrolysis is dynamic in the manner that almost all the starch is hydrolyzed at the old flower stage, as it has been observed in other species (Durkee et al., 1981; Pacini et al., 2003; Nepi et al., 2006). In the case of U.

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Sharaf Al-Tardeh Discussion maritima, the flower opens at about 1:00 am and closes the following evening at 19:00. The maximal nectar volume is about 7 µl which is produced at 5:00 am and drops to 1 µl at 9:00 pm. The nectar initially has a low sugar concentration (shortly after secretion ca. 10-15%), but this increases rapidly (to 75%) toward noon into almost crystalline form. Nectar concentration is affected directly by both temperature and relative humidity, being higher in dry conditions (Dafni and Dukas, 1986). These facts are in agreement with our results according to which nectar production rate increases from young to intermediate stages and decreases in old flowers. More specifically, the concentration of starch during the initial stages of the flower development is higher than that at the old stage.

4.4.1.4 Fate of the nectary The fate of the nectary after the secreting phase could be either a breakdown of the nectary epithelium as in the female flower of Musa paradisiaca (Fahn and Kotler, 1972) or the transformation of the nectary tissue into parenchyma as in Aloe, Gasteria and Tillandsia (Schnepf and Pross, 1976; Cecchi-Firordi and Palandri, 1982). The transformation occurs by means of an elongation of the epithelium cells and occlusion of the nectary cavity. The differentiation of transfer cells in septal nectaries is supposed to be an anatomical mechanism to increase nectar output (Schmid, 1985).

In U. maritima, differentiation of thickened outer walls in the epithelium cells is already evident at the young stage, which is an aspect that can be related to the differentiation of transfer cells to parenchyma fruit (Saunders, 1890; Schmid, 1985). However, it lacks the wall ingrowths. On the contrary, at the end of secretion (3-4 h from the flower opening) the epithelial cells maintain their initial shape, even at the old stage with cytological modifications. The latter depend on the increasing of color intensity, presence of electron dense material and large vacuoles, and absence (degradation) of the endoplasmic reticulum and amyloplasts. These modifications support the hypothesis of the breakdown of the nectary. The thickness of the nectary decreases at the old stage, being approximately six cells wide from the epithelium to the ovary wall. In addition, the nectary cavity might be occluded. Moreover, the deposition of callose in the thickened outer walls signals the end of

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Sharaf Al-Tardeh Discussion secretion activity (Schnepf and Pross, 1976). Finally, the fate of gynopleural nectary of U. maritima could be the transformation of the epithelial cells to parenchyma.

4.4.1.5 Nectar presentation and pollinators Flower morphology and site of nectar presentation, combined with nectar quantity and composition, are the main factors determining potential pollinators among nectar-feeding animals (Faegri and Van der Pijl, 1979; Baker and Baker, 1983; Proctor et al., 1996). An appropriate positioning of the nectar inside the flower ensures the efficiency of pollination while exploiting the nectar, thus, the visitor should inevitably contact the reproductive organs. Nectar presentation is primary when the site of nectar production and the site of nectar emission are the same. However, when these sites are different, the term secondary presentation is used. As in all plants with gynopleural nectaries, U. maritima has secondary nectar presentation. This kind of presentation and ducts were reported for species of Asphodelaceae (Weryszko-Chmielewaska et al., 2006; Nepi et al., 2006) and Alliaceae (Vogel, 1998).

Competition for pollinators is regarded as a possible selective pressure leading to the shift of flowering season from spring (the main flowering season under Mediterranean climatic conditions) to autumn, as it occurs in some geophytes with hysteranthous leaves, as is the case of U. maritima (Dafni et al., 1981; Al-Tardeh et al., 2008c). The latter exposes to the pollinators both its pollen and its nectar with sugar concentration varying from 10-75% (Dafni and Dukas, 1986) in order to attract different pollinators (Corbet, 1978). Besides that, alternative nectar resources are very scarce. The horizontal orientation of the flowers and the drainage of the nectar droplets to the base of the ovary in close contact to the base of the anthers could be an adaptive strategy aiming to enable nectar to be consumed easily by the visitors which in turn enhance fertilization. Moreover, the flowering occurrence in autumn, after a very dry season (summer), can be interpreted as a preference of the plant to produce nectar with high viscosity. High viscosity of nectar is due to the shortage in the available water within the bulb after a very dry summer and some water loss from nectar by evaporation (Al-Tardeh et al., 2008c). This phenomenon was, also, reported in other species of Asteraceae (Wist and Davis, 2006).

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Up to now, there have been no reports on flower initiation. However, flowering could be enhanced by applying a temperature of 28 °C for one month combined with more than 48% relative humidity (Scaramuzzi and Bianco, 1962). Flowering can also be hastened by producing a larger than normal temperature difference between day and night (McCorohan, 1990). The flowering shoot of U. maritima shows patterns of very rapid elongation. For instance, it takes only 25 days to reach a length of about 140 cm and flowering, while florets continue to open for 19 days (Mitrakos et al., 1974). This pattern can be interpreted as an adaptive strategy aiming to reduce the plant exposure to the environmental hazards of this harsh season (autumn).

4.4.2 Contents of raphides Raphides, or needle-shaped crystals of calcium oxalate, usually, occur in cells of U. maritima adventitious roots (Al-Tardeh et al., 2006), leaves (Al-Tardeh et al., 2008b), bulb parts (Al-Tardeh et al., 2008a), subsidiary tissue and the ovarian wall of the flower (Al- Tardeh et al., 2008c). Morphologically, their aciculate shape is a critical component of the proposed defense mechanisms (Davies et al., 2000; Cogne et al., 2001; Salinas et al., 2001; Ruiz et al., 2002). In addition, twinning is an important factor in allowing plant cells to produce the raphide morphology (Arnott and Webb, 2000). Moreover, raphides may be involved in phloem metabolism and the active transport of sucrose (Elias and Gelband, 1977).

The presence of cells with polysaccharidic content is evident in the tissues of the ovarian body. Polysaccharide concentration seems to be the highest at the old stage and the lowest at the young stage, while the intermediate one occurs at the intermediate stage. Moreover, the relative volume of the idioblast cells containing bundles of raphides reaches the highest value at the young stage (7.41 ± 1.8 %). The relative volume decreases at the intermediate stage (4.33 ± 1.8 %) and increases again at the old stage (6.45 ± 1.55 %). These products may provide beside the ovarian epidermal cuticle, an extra defense protection to the flower, particularly for its reproductive parts.

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4.4.3 The obturator gland The obturator is a placental protuberance at the ovary entrance connecting the transmitting tissue with the ovarian cavity in Rosaceae (Arbeloa and Herrero, 1987; Herrero, 1992; Cousin and El Mattaoui, 1998), Liliaceae (Tilton and Horner, 1980), and Fabaceae (Tilton et al., 1984). The obturator may have a role in fertilization process by secreting different components involved in the heterotrophic growth of the pollen tubes in the final stage just before they penetrate the micropyle (Arbeloa and Herrero, 1987) or to control of the direction of pollen tube growth (Tilton et al., 1984)

Axile placentation, in which the ovules are borne on a central column of tissue in the partitioned ovary of U. maritima is evident. The secretory cells of the obturator are epithelial and resemble those of the nectary ones. Most of the plastids contain huge starch granules during the initial stages. However, the plastids become devoid of their contents at the stages of mature and overmature ovary and/or nectary. The mucilage is secreted by the dictyosomes in the form of mucoprotein, while the protein units wree provided by the endoplasmic reticulum. Therefore, the secreted products of the obturator gland are of sugary contents which might be involved in the heterotrophic growth of the pollen tubes or to in controlling the direction of pollen tube growth. The obturator gland diminishes to its minimum size and becomes inactive at the old stage.

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CHAPTER V

CONCLUSIONS

The geophyte Urginea maritima (L.) Baker is well adapted to the Mediterranean- type climate by means of synchronization with its seasonality and periodicity. The plant is characterized by exhibiting seasonal and micro-environmental adaptation, i. e. each plant organ is well adapted to the environmental conditions of its growing season.

The roots of U. maritima play an important role in storing and utilizing water and nutrients, thus protecting the plant from drought stress and environmental hazards.

The presence of velamen is an adaptive strategy of the plant in the Mediterranean region, aiming to use up short seasonal rainfalls. The velamen provides mechanical protection to the cortex and reduces water loss. It may also play a role in aeration and absorption of water, minerals and the nitrogen.

The cortex proves to be the basic storage region of the roots since it contributes to the total root thickness by 75%. Correspondingly, the cortical storage cells occupy 34.11% of the total root volume. The cortical storage cells are specialized for storage of polysaccharides, lipids, mucilage and water.

The stored materials of polysaccharides, lipids and water are the raw materials for the development of new shoots. Moreover, as part of these functions, the stored materials play a role in the defense of the plant against herbivores and other environmental hazards, as well as they seem to synchronize the plant with the seasonality and periodicity of the Mediterranean climate.

The bulb of U. maritima is well adapted and synchronized to the Mediterranean climate. The different bulb parts serve different ecological functions in terms of their resources and their importance for these functions. The basal plate is the active centre, giving rise to one or two apical meristems and roots in autumn, as well as the

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flowering bud in late summer. The middle region of the bulb (almost, the 3rd bulb- scale) stores the resources and the tunics give mechanical defense.

The main adaptive strategy is the development of a semi-subterranean life form and a perennial type in order to survive the unfavourable seasons, especially as a primarily avoidance of drought and preservation of nutrients from herbivores and environmental hazards.

The tunic provides protection against disease infection and mechanical damage to the outer scales and the basal plate that contain the adventitious roots initials. Beside that, the tightly packed epidermis and smooth cuticle provide an extra mechanical protection for the scales and the basal plate.

Anatomically, the storage cells occupy 18.31-50.66% of the total volume of the bulb. The stored water, lipids, oils, polysaccharides, mucilage and/or glycosides are a prerequisite for flowering.

The defense strategies depend mainly on the presence of raphides, polysaccharides, mucilage and/or glycosides. The toxic mucilage is the main defense principle against herbivores, insects and rodents. Moreover, the mucilage is associated with water storage while the bundles of raphides support the central cylinder.

Depending on the morpho-anatomical studies, one could be led to the conclusion that most of the bulb occurs as a storage tissue, which, in turn accounts for the meaning of the term ‘geophyte’.

Leaves first appear after the flowers have wilted in response to winter rains onset during November or December, and may remain green until late spring (May), depending on rainfall and temperature.

U. maritima possesses mesomorphic leaves. These are equifacial in order to achieve the highest photosyntheic rate. This could be fulfilled by shortening the distances of

CO2 diffusion, the high density of sunken stomata on both sides, the fine structure

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of the mesophyll cells, the distribution of the chloroplasts parallel to the cell wall facing the intercellular spaces, the large air spaces in the mesophyll area and the low LTD and SLM.

It has been suggested that U. maritima has evolved two different forms, a more mesophytic, and another more xerophytic, to optimize adaptation to the seasonal fluctuation of environmental conditions throughout the year.

The defense strategies of the leaves depend on the presence of the thick cuticle, well packed epidermis, the presence of the crystalloid inclusions and other stored compounds. Before leaf senescence, the photosynthetized compounds, especially polysaccharides, are translocated to the storage underground organs.

The inflorescence stalk of U. maritima appears during August or September. Competition for pollinators is regarded as a possible selective pressure leading to the shift of flowering season from spring to autumn.

The flowering shoot is characterized by rapid and one-dimensional growth. The opened flowers constitute an open-flower zone continually moving toward the apex. This rapid pattern is an adaptive strategy in order to reduce exposure of the plant to the environmental hazards of this harsh season (autumn).

The flowering occurrence in autumn, after a very dry season (summer) is due to the plant preference to produce nectar with high viscosity.

U. maritima possesses a typical gynopleural (septal) nectary with secondary presentation. The nectary consists of one layer of epithelial cells and 1-4 layers of subsidiary tissue followed by 2-6 layers of parenchyma cells.

The outlets of the nectar occur at a distance of the two-thirds from the top of the ovary by means of the carpellary suture. Nectar secretion mechanism depends largely on the hydrolysis of the starch grains stored in amyloplasts.

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The young stage of the nectary is characterized by the abundance of endoplasmic reticulum with ribosomes and the small starch granules inside the mostly elongated amyloplasts.

The intermediate stage of the nectary is characterized by the presence of dense cytoplasm, active and abundant rough endoplasmic reticulum, mitochondria, dictyosomes and variously shaped plastids. Numerous secretory vesicles, myelin-like figures and vacuoles with electron dense materials are also dominant.

The old stage of the nectary is characterized by crystallization of nectar and large vacuoles inside the epithelial cells, collapse of the cortical parenchyma cells, completely hydrolysed starch, as well as disappearance of the amyloplasts and ER.

Some subsidiary cells degenerate and/or transform into parenchyma cells. However, the epithelial cells maintain their initial shape even in the old stage.

The ovary of U. maritima is partitioned into three locules formed from the fusion of

three carpels. After fertilization, however, about 12 seeds per capsule are produced.

The obturator is a placental protuberance secreting mucilage of polysaccharidic

nature, which may be involved in the heterotrophic growth of the pollen tubes or in

controlling its direction.

206

Sharaf Al-Tardeh Acknowledgements

ACKNOWLEDGEMENTS

I would like to gratefully acknowledge the State Scholarship Foundation (IKY),

Greece, for the Ph.D. scholarship that offered to me during the years 2004-2007.

I would like also to acknowledge the Arab Student Aid International (ASAI) Dublin,

USA, for a small loan that they provided me during the year 2007.

I would like to express my deep acknowledgement to the secritarate of the

Department of Botany, Mrs Arete Dimopoulou, for the linguistic revision of this

doctoral dissertation.

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CHAPTER VI

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Yahel, H. 1990. Growth retardants for flowering pot plant production of Narcissus ‘Grand Soleil d’Or’. Acta Hortic., 266: 205-210.

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248

Sharaf Al-Tardeh Original papers

ORIGINAL PUBLISHED ARTICLES

1194 Nectary structure and nectar presentation in the Mediterranean geophyte, Urginea maritima (Hyacinthaceae)

Sharaf Al-Tardeh, Thomas Sawidis, Barbara-Evelin Diannelidis, and Stylianos Delivopoulos

Abstract: The morphology, anatomy, and ultrastructure of the floral nectary of Urginea maritima (L.) Baker were investigated at three stages of nectary development. The plant possesses a typical gynopleural (septal) nectary with secondary presentation. The nectary consists of one layer of epithelium secretory cells and one to four layers of subsidiary cells subtended by two to six layers of parenchyma (subnectary) cells. The nectary releases the nectar at a point two-thirds towards the summit of the ovary by means of carpellary sutures. Nectar secretion appears to depend largely on the hydrolysis of starch grains stored in amyloplasts at the intermediate stage. The hydrolysis process most likely commences in the epithelium layer followed by the subsidiary tissue and then the parenchyma cells of the ovary wall. A symplastic transfer of the secreted nectar occurs by plasmodesmata connecting the subsidiary cells to the parenchyma and the epithelial secretory cells. However, microchannels in the cell wall of the epithelial cells may facilitate the apoplastic transfer of the nectar into the nectary cavity. The old stage of nectary development is characterized by a crystallized form of nectar, collapse of the parenchyma cells, complete starch hydrolysis, and disappearance of the amyloplasts and endoplasmic reticulum. Key words: amyloplast, geophytes, nectar secretion, septal nectary, starch, Urginea maritima. Re´sume´ : Les auteurs ont e´tudie´ la morphologie, l’anatomie et l’ultrastructure des nectaires floraux de l’Urginea maritima (L.) Baker, a` trois stades du de´veloppement des nectaires. La plante posse`de un nectaire (septe´) gynopleural typique avec pre´sentation secondaire. Le nectaire comporte une couche e´pithe´liale de cellules sećre´trices et 1–4 couches de cellules subsidiaires sous-tendues par 2–6 couches de cellules de parenchyme (sous-nectaire). Les nectaires relaˆchent le nectar a` un point situe´ aux deux tiers vers le sommet de l’ovaire, au moyen de structures carpellaires. La sećre´tion du nectar semble de´pendre de l’hydrolyse de grains d’amidon en re´serve dans des amyloplastes au stade interme´diaire. Le processus d’hydrolyse commence vraisemblablement dans la couche e´pithe´liale, suivi du tissu subsidiaire et par apre`s des cellules de parenchyme de la paroi ovarienne. On observe un transfert symplastique du nectar sećre´te´ par des plasmodesmes reliant les cellules subsidiaires au parenchyme et aux cellules e´pithe´liales. Cependant, des microcanaux dans la paroi cellulaire des cellules e´pithe´liales pourraient faciliter le transfert apoplastique du nectar dans la cavite´ a` nectar. Le dernier stade du de´veloppement des nectaires se caracte´rise par une forme cristalline de nectar, un affaissement des cellules de parenchyme, une hydrolyse comple`te de l’amidon et une disparition des amyloplastes et du re´ticulum endoplasmique. Mots-cle´s:amyloplastes, geóphytes, sećre´tion du nectar, amidon, Urginea maritima. [Traduit par la Re´daction]

Introduction nectar production (Smets et al. 2000). Flower morphology and site of nectar presentation, combined with nectar quan- A topographical classification of floral nectaries indicates tity and composition, are the main factors determining po- nine different types (Fahn 1979). Among them, the ‘‘ovarial tential pollinators among nectar-feeding animals (Faegri and nectary’’ type includes nectaries that are located in the septal Van der Pijl 1979; Baker and Baker 1983; Proctor et al. region between adjacent carpels, the so-called septal necta- 1996). An appropriate positioning of the nectar inside the ries or gynopleural nectaries as they have been more re- flower ensures the efficiency of pollination: while exploiting cently defined by Smets and Cresens (1988). Gynopleural the nectar, the visitor would likely contact the reproductive nectaries are restricted to monocotyledons, where they repre- organs. sent the most common type of floral nectary (Smets et al. It is generally accepted that nectar originates from phloem 2000). The gynopleural nectary, being a cavity inside the sap (Fahn 2000; Gaffal et al. 2007). The sugar-secreting ovary, is not directly exposed to nectar-feeding animals, and nectaries are undisputedly sink organs dependent on source the site of nectar emission is often different from that of tissue assimilates. Most nectaries are vascularized exclu-

Received 22 January 2008. Published on the NRC Research Press Web site at botany.nrc.ca on 2 October 2008. S. Al-Tardeh, T. Sawidis, B.-E. Diannelidis, and S. Delivopoulos.1 Department of Botany, School of Biology, Aristotle University, Thessaloniki 541 24, Greece. 1Corresponding author (e-mail: [email protected]).

Botany 86: 1194–1204 (2008) doi:10.1139/B08-075 # 2008 NRC Canada Al-Tardeh et al. 1195 sively by phloem strands (Frei 1955). The efflux of carbohy- Heaps and Northcote 1969), they were photographed using drates may occur across the plasma membranes of the sieve an inverted photomicroscope ECLIPSE TE2000-S (Nikon). element – companion cell complexes and into adjacent cell Ultra-thin sections (0.08 mm thick) were collected on copper walls (apoplastic pathway) or through the plasmodesmata in- grids and stained with uranyl acetate and lead citrate (Rey- terlinking these complexes with adjacent cells (symplastic nolds 1963). The sections were examined and photographed pathway; Patrick et al. 2001). On the other hand, starch in a Zeiss 9 S-2 transmission electron microscope (TEM). stored within amyloplasts at the young (presecretory) stage For scanning electron microscopy (SEM), cross and longi- can be utilized both as a source of energy for highly meta- tudinal sections were made of the ovaries from the three bolic processes and as a source of sugars for nectar synthesis flower stages. To be able to distinguish between top and (Stpiczynˇska et al. 2003). bottom, we left the upper part of each ovary with 1 mm of Urginea maritima (L.) Baker is a perennial bulbous geo- the style still attached. The materials were fixed for 3 h at phyte of the family Liliaceae (Bruneton 1996). However, ac- room temperature in the same buffered fixative as described cording to the new taxonomy of monocots, it belongs to the above (Karnovsky 1965). After postfixation in 1% osmium order Asparagales of the family Hyacinthaceae (Mabberley tetroxide for 3 h at room temperature and dehydration in an 1997). It generally occurs on the slopes of hills and sandy increasing acetone series (30%, 50%, 70%, 90%, and grounds near the Mediterranean Sea (Gentry et al. 1987), 100%), the sections were critical-point dried in a Balzers and in certain regions of Northern Africa (Bellakhdar CPD 030 device. Sections were mounted on SEM stubs, 1997), the Middle East, and Europe (Al-Tardeh et al. 2006). then sputter coated with carbon in a JEE-4X vacuum evapo- In this study, we examine the morphology, anatomy, and rator. Observations were made with a JSM 840-A SEM ultrastructure of the floral nectary in relation with nectar se- (Tokyo, Japan). cretion, from its commencement to cessation, at three stages of flower development. This study provides some possible Histochemistry explanations for the abundance of the plant in the Mediterra- To stain lipophilic substances, semi-thin sections of fixed nean region. material or hand-cut sections of fresh ovaries were stained with 1% sudan black B and 2% osmium tetroxide (Bronner Materials and methods 1975), respectively. For polysaccharide staining, sections of fixed or fresh material were treated with periodic acid – Study site and plant material Schiff’s reagent (PAS) according to Nevalainen et al. The study site was located at Souda (35833’N, 24807’E), a (1972) and examined with LM. town about 10 km south of Chania, on the island of Crete, southern Greece. Souda’s climate is of the Mediterranean Results type, and most of its annual rainfall occurs in autumn– winter, whereas the summer is very hot and dry. The Flower morphology and nectar presentation mean annual air temperature is 18.1 8C. January (10.8 8C) The flowering shoot of U. maritima emerges as a stalk and February (10.8 8C) are the coldest months, while July from the bulb and is followed by a characteristic very rapid (26.3 8C) and August (26 8C) are the warmest. It is also pattern of subapical elongation over a few days. Elongation characterized by high humidity and low barometric pres- of the inflorescence axis is accompanied by opening of the sure because of its close location to the sea and the moun- flowers in an acropetal direction shown in Fig. 1. tains. Each flower consists of six elongated white petals, six Flowers of U. maritima were collected from plants grow- conspicuously long stamens with green to deep brown ing wild in Souda. Serial hand-cut sections of the flower anthers, and a central gynoecium with a light green superior were obtained and examined by using an Olympus SZX12 ovary (Fig. 2). The trilocular ovary results from a fusion of stereo-microscope to determine the position of the nectary. three isomerous carpels forming a cavity, where nectar accu- Three different flower stages were examined: (i) young mulates (Fig. 3). The presence of stomata on the outer sur- flowers with the corolla starting to open but not all anthers face of the ovary is evident (Fig. 4). In addition, when the dehisced; (ii) intermediate flowers with all anthers dehisced flower senesces (old stage), the petals do not fall but close and the corolla completely open; (iii) old flowers in which up again and resume a cylindrical structure similar to that the corolla had started to wilt (Fig. 1). of the buds (Fig. 1). The closed petals protect the growing fruit (capsule). When the fruit is ripe, it splits open to re- Microscopy (light, transmission, and scanning electron lease its black winged seeds. microscopy) Floral parts of young, intermediate, and old flowers were Nectary structure and development fixed for 3 h at room temperature in a mixture of 2.5% glutaraldehyde and 2% paraformaldehyde fixative (Karnovsky Young stage 1965) in 0.1 mol/L sodium cacodylate buffer, pH 7.1. After The gynopleural nectaries consist of three clefts located in postfixation in 1% osmium tetroxide for 3 h at room temper- the septal region between adjacent carpels (Fig. 5). The nec- ature and dehydration in an ethanol series, the samples were tary cavity is very narrow and not fully developed (Figs. 6 embedded in low viscosity resin (LV). For light microscopy and 7). The single-layered epithelium is subtended by a sub- (LM), semi-thin sections (1 mm thick) were cut on a sidiary tissue of 2–5 layers of vacuolated cells that are Reichert Om U2 ultramicrotome. After staining with 0.5% smaller than the parenchyma cells in the other parts of the toluidine blue O (TBO) in 5% borax solutions (Pickett- ovary (Fig. 6). Idioblastic cells containing bundles of ra-

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Figs. 1–5. Habit of the inflorescence, flower, and gynopleural nectary of Urginea maritima. Fig. 1. Photo of inflorescence stalk of Urginea maritima containing more than 250 florets. Flower opening along the inflorescence occurs acropetally; three flower stages were designated: young (1), intermediate (2), and old (3). Scale bar = 3 cm. Fig. 2. Intermediate stage showing arrangement of floral parts. Scale bar = 0.5 cm. Figs. 3–5. Scanning electron micrographs. Fig. 3. Ovary showing the outer grooves of the septal slits (arrows) from where nectar drains. Scale bar = 2 mm. Fig. 4. Stomata in the ovary’s outer surface (arrows). Scale bar = 50 mm. Fig. 5. Transverse section of the ovary showing localization of the three gynopleural nectaries (arrows) alternating with the ovary locules. O, ovules. Scale bar = 1 mm.

# 2008 NRC Canada Al-Tardeh et al. 1197 phides are located among the ovary parenchyma (Fig. 6). A tion, numerous organelles such as amyloplasts, Golgi bodies, very thin and irregular cuticle is present on the surface of and mitochondria are also present (Figs. 18 and 19). Vari- the secretory epithelium (Figs. 6 and 8) that consists of ously shaped plastids containing osmiophilic granules in the small and long cells with dense cytoplasm and large nuclei stroma, peripheral thylakoids, and starch granules of various (Fig. 7). Cisternal profiles of endoplasmic reticulum (ER) sizes also characterize the cytoplasm (Fig. 19). and ribosomes dominate the cytoplasm of the secretory epi- The nectary is subtended by the parenchymatic septal tis- thelium (Fig. 8). sue (subnectary parenchyma), which consists of approxi- The floral nectary is situated at a distance of two-thirds mately 2–6 layers of relatively large isodiametric the height of the ovary (Fig. 5). The three septal slits pro- parenchyma cells. The ovary wall is made up of approxi- ceed downwards entering the ascidiate zone of carpels mately 3–10 layers of parenchyma cells (Fig. 23). The sub- (Fig. 3). Development of starch granules begins in the secre- nectary parenchyma is located below the nectary tory epithelium cells, which contain mostly elongated pro- parenchyma (subsidiary tissue), and consists of larger cells plastids that are, usually, devoid of starch grains (Fig. 9). with larger vacuoles, less dense cytoplasm, and pronounced Golgi bodies and a few plastids with large starch grains oc- intercellular spaces (Fig. 6). Amyloplasts are variously cur in the secretory epithelium cells (Fig. 10). However, shaped, almost devoid of stroma, and contain many starch more abundant and larger starch grains occur in cells of the grains (Fig. 20). Amyloplasts with starch grains are more subsidiary tissue (Fig. 11). The epidermal cells of the ovary concentrated in the peripheral parenchyma cells (small mi- are elongated, compactly arranged, with concaved margins crograph in the corner of Fig. 14). During nectar secretion, and are anticlinally oriented (Figs. 6 and 13). A smooth cu- starch content drastically diminishes from the nectary epi- ticle of 0.52 ± 0.25 mm thick covers the epidermal cells, thelium toward the peripheral parenchyma cells. which is more obvious under higher magnification (Fig. 12). The vascular tissue occurs in the subnectary parenchyma Histochemical analyses showed that the content of poly- near to the placenta. More specifically, it is located at the saccharide is very low, since only a few idioblastic cells interior side of the nectary in conjunction with the obturator containing bundles of raphides are red following staining tissues (Fig. 6). It might serve both the nectary and the ob- with Schiff’s reagent (Fig. 13). Sudan black B staining was turator. The vascular bundles consist of xylem tracheary ele- negative for all tissues. ments and sieve elements (Fig. 21). However, vascular tissue that occurs close to the nectary consists of xylem tra- Nectar outlet and presentation cheary elements only (Fig. 21). Whereas, the vascular tissue, Each nectary cavity has a nectar outlet located at one- which occurs in pairs, one on each side of the nectary, con- third of the distance of the ovary from the base of the style sist of phloem strands only (Fig. 22). The sieve elements are (Fig. 3). Nectar drains from the region around the inferior connected by sieve pores (Fig. 22). Plastids with crystalloid two-thirds of the ovary to its base and accumulates as muci- inclusions are present in the mature sieve elements (Fig. 22). laginous droplets, one for each nectary, as a secondary pre- The bundles supply the nectary with carbohydrates. There- sentation to be exposed to nectar-feeding animals. The fore, they are absent from the region where the nectary tis- nectar outlet is derived from the merging of an invagination sues are reduced as in the top of the ovary. of the cutinized epidermal surface, in continuity with the From the histochemical point of view, the highest concen- carpellary suture with the apical part of the nectary tration of polysaccharides occurs in the first third from the (Fig. 14). Small cells are present in the vicinity of the merg- base of ovary because of the strong reaction showed with ing point (Fig. 14). The carpellary suture is wide at the base Schiff’s reagent (Fig. 23). The intensity of the red colour of of the ovary, but it becomes deeper and narrower towards the idioblastic cells containing bundles of raphides is higher the top of the ovary. Two-thirds from the top of the ovary, than that of the young stage (compare Figs. 13 and 23). the invagination of the epidermal surface has tightly conni- Starch grains in the ovary wall react positively with this vent margins, except for the inner part where a tubular struc- stain (Fig. 23). In addition, the nectar remnants inside the ture is formed (Fig. 14). The tubular structure becomes nectary cavity and the starch grains in the subnectary paren- deeper towards the base of the ovary, where it merges with chyma react positively with Schiff’s reagent (Fig. 24). Lip- the apical part of the nectary. Although the tubular structure ophilic substances are absent as shown by the negative is in continuity with the outside, this communication is staining with Sudan black B. obstructed by the presence of the cuticle that occludes the narrow space between the connivent margins of the epider- Old stage mis. The epithelial and subsidiary tissue cells undergo cytological modification during development, being more vacuo- Intermediate stage lated and with an irregular nuclear shape in the old flower The intermediate (active) stage of the nectary is desig- stage (Fig. 25). Some cells in the subsidiary tissue seem to nated as the period of the flower’s anthesis and nectar secre- degenerate (Fig. 25). The thickness of the nectary decreases tion. Fully developed ER cisternae with attached ribosomes in the old stage, being approximately six cells wide from the forming characteristic profiles are located in the vicinity of epithelium to the ovary wall (Fig. 25). The pores (micro- the outer cell walls of secretory epithelial cells (Fig. 15). Se- channels), through which the nectar is discharged into the cretory epithelial cells also contain many narrow and curled nectary cavity, are more obvious in the cell wall of the epi- plastids (Fig. 16), numerous secretory vesicles (Fig. 17), thelial cells (Fig. 26). The old stage is characterized by a myelin-like figures, vacuoles with electron-dense materials crystallized form of nectar inside the epithelial cells (Fig. 18), and ER with dilated cisternae (Fig. 19). In addi- (Fig. 27), collapse of the parenchyma cells (Fig. 28), com-

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Figs. 6–11. Young stage of nectary of Urginea maritima. Fig. 6. Light micrograph of a transverse section of young ovary stained with toluidine blue O (TBO). The nectary consists of a compactly arranged epithelium (Ne), a subsidiary tissue (St), and subnectary parenchyma (Sn). Idioblastic cells containing bundles of raphides (Id) and vascular bundles (Vb) are present in the subnectary parenchyma. Ep, epidermis. Scale bar = 50 mm. Figs. 7–11. Transmission electron micrograph of transverse sections of the ovary. Fig. 7. The narrow nectary cavity (arrows) and the epithelium made up of cells with dense cytoplasm and relatively large nuclei. Scale bar = 10 mm. Fig. 8. Epithelium cell with endoplasmic reticulum (ER, arrows) and uneven cuticle (arrowheads). Scale bar = 10 mm. Fig. 9. Epithelial cell containing starch grains at the peripheral side of an elongated proplastid (Ps). Scale bar = 1 mm. Fig. 10. Well-developed starch grain (Sg) and Golgi bodies (Gi) in an epithelial cell. Scale bar = 1 mm. Fig. 11. Large starch grains (Sg) in the subsidiary tissue. Scale bar = 1 mm.

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Figs. 12–19. Figs. 12–13. Young stage of nectary of Urginea maritima. Fig. 12. Epidermal cell covered by a thin layer of cuticle (Cu). Cw, cell wall. Scale bar = 1 mm. Fig. 13. Light micrograph of a transverse section of the ovary stained with Schiff’s reagent. A cell containing red-stained (web version only) mucilage (arrow) is indicated. Id, idioblast cells containing bundles of raphide crystals. Scale bar = 50 mm. Figs. 14–19. Intermediate stage of nectary of Urginea maritima. Fig. 14. Light micrograph of a transverse section at the middle of ovary illustrating the tubular structure (Ts). Starch grains in the subnectary parenchyma are shown in the inset. Scale bar = 25 mm. Figs. 15–19. Transmission electron micrograph showing transverse sections of an ovary. Fig. 15. Epithelial cell with well-developed endoplasmic reticulum (ER) that occurs as long strands of parallel cisternae near the outer wall. Scale bar = 10 mm. Fig. 16. Granular cytoplasm of epithelial cells with many narrow, curled plastids. Scale bar = 1 mm. Fig. 17. Secretory vesicles in epithelial cells. Scale bar = 10 mm. Fig. 18. Dense granular cytoplasm of the epithelial cells with vacuoles containing electron-dense materials (arrows) and secretory vesicles (arrowheads). Mi, mitochondria. Scale bar = 1 mm. Fig. 19. ER with dilated cisternae (arrows) and plastids (Pl) in an epithelial cell. Scale bar = 1 mm.

# 2008 NRC Canada 1200 Botany Vol. 86, 2008 pletely hydrolysed starch, as well as disappearance of the cases, proplastids differentiate into amyloplasts and store amyloplasts and ER. The relation between nectar production large amounts of starch in many large grains per plastid be- and starch hydrolysis is dynamic in the manner that almost fore nectar secretion begins (Durkee et al. 1981; Pais and all of the starch content is hydrolyzed at the old flower Figueiredo 1994; Nepi et al. 1996). Amyloplasts in the stage. nectar-producing parenchyma are generally almost devoid of stroma and packed with starch (Nepi et al. 1996). They Discussion also contain many starch grains per amyloplast, which increases starch surface area, thus facilitating and speeding up There are two main nectary types in monocotyledons: hydrolysis during nectar production. Therefore, many septal (i.e., persistent) and perigonal (i.e., caducous) (Smets authors infer that hydrolysis of starch in the parenchyma et al. 2000). In U. maritima, the typical septal (gynopleural) contributes directly to nectar carbohydrate content (Durkee nectary is found. Nectar presentation is characterized as et al. 1981; Pais and Figueiredo 1994; Nepi et al. 1996; primary when the site of nectar production and the site of Pacini et al. 2003; Peng et al. 2004). nectar emission are the same. However, when these sites Starch stored within amyloplasts at the young (presecre- are at different points, the term secondary presentation is tory) stage can be utilized both as a source of energy for used (Pacini et al. 2003; Nepi et al. 2006). As in all plants highly metabolic processes and as a source of sugars for having septal nectaries, U. maritima has secondary nectar nectar synthesis (Stpiczynˇska et al. 2003). During successive presentation. Secondary presentation in U. maritima seems stages of secretory activity, where predominantly granulo- to be an adaptive strategy for protecting the nectary tissues crine secretion seems likely, plastids generally become elon- from nectar-feeding animals and environmental hazards. gated or develop an irregular profile, and this is usually This kind of presentation and ducts have also been reported associated with depletion in starch content (Sawidis et al. for species of Asphodelaceae (Nepi et al. 2006; Weryszko- 1989; Sawidis. 1998; Horner et al. 2007). Degradation of Chmielewska et al. 2006) and Alliaceae (Vogel 1998). plastids (amyloplasts) starts with hydrolysis of starch grains The fate of the nectary after the secreting phase could be (Gaffal et al. 2007). The engulfing pattern of starch hydrol- either a breakdown of the nectary epithelium as in the fe- ysis occurs in the floral nectaries of Eccremocarpus scaber male flower of Musa paradisiaca (Fahn and Kotler 1972) (Belmonte et al. 1994), Cucumis sativus (Peng et al. 2004), or the transformation of the nectary tissue into parenchyma and senescing French bean leaves (Minamikawa et al. 2001). as in Aloe, Gasteria, and Tillandsia (Schnepf and Pross This mechanism of starch hydrolysis involves engulfing of 1976; Cecchi-Fiordi and Palandri 1982). During the break- the plastids by vacuoles and subsequent hydrolysis of starch down of the nectary tissue, the cytoplasm becomes very in the vacuole (Belmonte et al. 1994; Minamikawa et al. electron-dense, plastids and mitochondria degenerate, and 2001; Peng et al. 2004). the vacuole increases gradually in volume until it occupies In U. maritima, the flower opens at about 0100 hours and most of the cell (Fahn and Kotler 1972). On the contrary, closes the following night at 1900 hours. The maximum transformation of nectary tissue into parenchyma occurs by nectar volume of about 7 mL takes place at 0500 hours and means of elongation of the epithelial cells and occlusion of drops to 1 mL at 0900 hours (Dafni and Dukas 1986). The the nectary cavity by acidic polysaccharides (Schnepf and nectar initially has a low sugar concentration (shortly after Pross 1976). In the case of U. maritima, vacuolation and secretion ca. 10%–15%), but this increases rapidly (to 75%) elongation of the epithelial cells is evident in the old flow- toward noon into almost a crystalline form (Dafni and ers, but the nectary cavity is still present. In addition, the ep- Dukas 1986). Nectar concentration is affected directly by ithelial cells’ outer walls are thicker in the old flower both temperature and relative humidity, being higher in dry (Fig. 25), in comparison with those of the young (Fig. 6) conditions (Dafni and Dukas 1986). These facts are in and intermediate-staged ones (Figs. 14 and 21). agreement with our results, according to which, nectar pro- The fate of the nectary parenchyma after secretion may duction increases from young to intermediate stages (nectar have different patterns when nectar secretion does not cause peak) and decreases in the old flower stage. Thus, the in- cell death. The nectary tissue may be involved in nectar re- creased starch hydrolysis elevates the nectar production dur- absorption (Nepi et al. 1996) or may differentiate into paren- ing the active stage and vice versa in the old stage. This chyma tissue, as in the case of septal nectaries (Schnepf and dynamic manner has also been reported for other species Pross 1976; Cecchi-Fiordi and Palandri 1982; Nepi et al. (Durkee et al. 1981; Pacini et al. 2003; Nepi et al. 2006). 2006) and degenerate. In the case of U. maritima, both de- Competition for pollinators is regarded as a possible se- generation and transformation of some subsidiary and sub- lective pressure leading to the shift of flowering season nectary cells occur. Moreover, the development of these from spring (the main flowering season under Mediterranean tissues seems to be an adaptive strategy to increase the nec- conditions) to autumn, as it occurs in some geophytes with tar output, as well as to support and protect the nectary epi- hysteranthous leaves, which include U. maritima (Dafni et thelium. al. 1981; Al-Tardeh et al. 2008b). The latter exposes to the Undifferentiated plastids (proplastids) are present in the pollinators both its pollen and nectar with sugar concentra- very early stages of nectary development (Nepi et al. 1996). tions ranging from 10% to 75% (Dafni and Dukas 1986) to Close to flower anthesis, they may differentiate to chloro- attract different pollinators (Corbet 1978). Beside that, alter- amyloplasts (Pacini et al. 1992; Nepi et al. 1996). In some native nectar resources are very scarce. The horizontal ori- cases, chloro-amyloplasts lose their thylakoid structure, and entation of the flowers and the drainage of the nectar starch grains increase in size a few days before anthesis droplets to the base of the ovary in close contact with the (Zer and Fahn 1992; Fahn and Shimony 2001). In other base of the anthers could be an adaptive strategy enabling

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Figs. 20–28. Figs. 20– 24. Intermediate stage of nectary of Urginea maritima. Transmission electron micrographs (TEM; Figs. 20 and 22) and light micrographs (LM; Figs. 21, 23, and 24) of transverse sections of ovary. Fig. 20. Abundant starch grains (arrows) of amyloplasts in peripheral parenchyma cell. Scale bar = 10 mm. Fig. 21. Vascular tissue consists of xylem vessels (Xv) and sieve elements (Se) at the conjunction with the obturator, whereas the vascular tissue occurring at the interior region of the nectary consists of xylem vessels (Xv) only. Scale bar = 50 mm. Fig. 22. Sieve pores (Sp) between sieve elements that contain crystalloid occlusions (arrows), in the isolated phloem located on each side of the nectary. Scale bar = 10 mm. Fig. 23. Starch grains (arrowheads) and idioblastic cells (arrows) intensively red stained (web version only) with Schiff’s reagent. Ow, ovary wall. Scale bar = 50 mm. Fig. 24. Starch grains (arrowheads) and nectar remnants in the nectary cavity (arrows) intensively red stained (web version only) with Schiff’s reagent. Scale bar = 50 mm. Figs. 25–28. Old stage of nectary of Urginea maritima. Fig. 25. LM micrograph of a transverse section of ovary showing epithelial (Ne) and subsidiary cells (St) more vacuolated than in the young stage. Some cells in the subsidiary tissue seem to degenerate (asterisks). Scale bar = 50 mm. Figs. 26–28. TEM micrographs of transverse sections of ovary. Fig. 26. Channels in the cell wall (arrows) of an epithelial cell. Scale bar = 1 mm. Fig. 27. Crystallized nectar (arrows) on the surface of an epithelial cell. Scale bar = 10 mm. Fig. 28. Collapse of the parenchyma cells and idioblastic cell containing bundles of raphides. Scale bar = 10 mm.

# 2008 NRC Canada 1202 Botany Vol. 86, 2008 nectar to be consumed easily by the visitors that in turn en- wide range of sugar concentration. In addition, the gyno- hance pollination and eventual fertilization. Moreover, the pleural nectary seems to play an important role in protecting flowering occurrence in autumn after a very dry season the plant’s sexual reproduction. Structurally, these functions (summer) can be interpreted as a preference of the plant to are based on: (i) development of a gynopleural nectary with produce nectar with high viscosity. The latter is due to the secondary presentation, (ii) presence of subsidiary and sub- shortage in the available water within the bulb after a very nectary parenchyma that elevate the nectar output, (iii) pres- dry summer and some water loss from nectar by evapora- ence of idioblastic cells containing bundles of raphides, and tion. This phenomenon has been reported in the Asteraceae (iv) transformation of the nectary tissue into parenchyma (Wist and Davis 2006). that assists fruit development. Different extents of the subsidiary tissue have also been observed in various species and have been related to differ- Acknowledgements ences in nectar production (Cecchi-Fiordi and Palandri Sharaf Al-Tardeh gratefully acknowledges the provision 1982; Nepi et al. 2006). Most studies concur that generally of a scholarship by the State Scholarship Foundation (IKY), both sugar and water components of the nectar are provided Greece. by phloem sap, and this is transported from sieve element to nectary cells (Fahn 2000; Pacini et al. 2003; de la Barrera References and Nobel 2004; Stpiczynˇska et al. 2005). In U. maritima, Al-Tardeh, S., Sawidis, T., Diannelidis, B.-E., and Delivopoulos, S. however, sugars are most probably transported from the 2006. Anatomical studies on the adventitious roots of the geo- bulb via phloem elements and are stored in the nectary cells phyte Urginea maritima (L.) Baker. J. Biol. Res. 5: 61–70. as starch during the young stage. A symplastic route de- [Available from www.jbr.gr/]. pendent on intercellular plasmodesmatal connection is avail- Al-Tardeh, S., Sawidis, T., Diannelidis, B.-E., and Delivopoulos, S. able in the floral nectary of U. maritima, from the sieve 2008a. Water content and reserve allocation patterns within the elements to the nectary epithelium. An apoplastic route is bulb of the perennial geophyte Urginea maritima (Liliaceae) in also hypothesized between the sieve elements, as witnessed relation to the Mediterranean climate. Can. J. Bot. 86: 291–299. by the sieve pores in their cell walls. Moreover, microchan- Al-Tardeh, S., Sawidis, T., Diannelidis, B.-E., and Delivopoulos, S. nels in the cell wall of the epithelial cells, similar to those 2008b. Morpho-anatomical features of the leaves of the Mediter- observed in other nectaries (Radice and Galati 2003), may ranean geophyte Urginea maritima (L.) Baker (Liliaceae). J. facilitate the apoplastic transfer of the nectar into the nectary Plant Biol. 51: 150–158. [Available from submission.bosk.or.kr/]. cavity. Baker, H.G., and Baker, I. 1983. Chemical constituents of nectar in Bundles of raphides usually occur in cells of U. maritima relation to pollination mechanisms and phylogeny. In Biochem- roots (Al-Tardeh et al. 2006), leaves (Al-Tardeh et al. ical aspects of evolutionary biology. Edited by M.H. Nitecki. 2008b), bulb parts (Al-Tardeh et al. 2008a), subsidiary tis- University of Chicago Press, Chicago, Ill. pp. 131–171. sue (Figs. 6, 13, and 28), and the ovary wall (Fig. 23) of Bellakhdar, J. 1997. La pharmacopee traditionnelle marocaine. Ibis the flower. Morphologically, their aciculate shape is a crit- Press, Paris, France. pp. 376–377. ical component in proposed mechanisms for defence (Davies Belmonte, E., Cardemil, I., and Kalin-Arroyo, M.J. 1994. Floral nectary structure and nectar composition in Eccremocarpus sca- et al. 2000; Cogne et al. 2001; Salinas et al. 2001; Ruiz et ber (Bignoniaceae), a hummingbird-pollinated plant of central al. 2002). Moreover, raphides may be involved in phloem Chile. Am. J. Bot. 81: 493–503. doi:10.2307/2445499. metabolism and the active transport of sucrose (Elias and Bronner, R. 1975. Simultaneous demonstration of lipids and starch Gelband 1977). It is worthy to notice that entire bundles of in plant tissues. Stain Technol. 50: 1–4. PMID:46630. raphides are immersed in a mucilaginous material. Mucilage Bruneton, J. 1996. Plantes toxiques. Ve´ge´taux pour l’homme et les cells have been reported in the subnectary parenchyma of animaux. Lavoisier, Paris, France. pp. 48–49. Hibiscus rosa-sinensis (Sawidis 1998). Because of the Cecchi-Fiordi, A., and Palandri, M.R. 1982. Anatomic and ultra- water-binding capacity of mucilage, with rapid water uptake structural study of the septal nectary in some Tillandsia (Brome- and slow release, it was hypothesized that, in this species, liaceae) species. Caryologia, 35: 477–489. mucilage cells offer an ideal regulation mechanism for water Cogne, A.L., Marston, A., Mavi, S., and Hostettmann, K. 2001. balance during nectar secretion and efficient protection of Study of two plants used in traditional medicine in Zimbabwe nectary tissue against water stress damage (Sawidis 1998). for skin problems and rheumatism: Dioscorea sylvatica and Ur- The flowering shoot of U. maritima shows very rapid pat- ginea altissima. J. Ethnopharmacol. 75: 51–53. doi:10.1016/ terns of elongation; for instance, it takes only 25 d to reach S0378-8741(00)00347-0. PMID:11282443. a length of about 140 cm and flowering, while florets con- Corbet, S.A. 1978. Bees and nectar of Echium vulgare. In The pol- tinue to open for 19 d (Mitrakos et al. 1974). This rapid pat- lination of flowers by insects. Edited by A. J. Richards. Aca- tern can be interpreted as an adaptive strategy to reduce demic Press, London, UK. pp. 89–96. exposure of the plant to the environmental hazards of this Dafni, A., and Dukas, R. 1986. Insect and wind pollination in Ur- ginea maritima (Liliaceae). Plant Syst. Evol. 154: 1–10. doi:10. harsh season (autumn). 1007/BF00984864. In conclusion, no sign of degeneration was observed in Dafni, A., Shmida, A., and Avishai, M. 1981. Leaf-less autumnal- the nectary of U. maritima. However, the plant possessing a flowering geophytes in the Mediterranean region: phytogeogra- gynopleural nectary is well adapted and synchronized with phical, ecological and evolutionary aspects. Plant Syst. Evol. the seasonality of the Mediterranean-type climate. The adap- 137: 181–193. doi:10.1007/BF00989872. tive strategies are the following: (i) development of a hyster- Davies, K.L., Winters, C., and Turner, M.P. 2000. Pseudopollen: its anthous type of flowering, (ii) having a very rapid growth structure and development in Maxillaria (Orchidaceae). Ann. pattern of the inflorescence, (iii) production of nectar with a Bot. (Lond.), 85: 887–895. doi:10.1006/anbo.2000.1154.

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