Absorption and Translocation of Zn Foliar Fertilisers

Cui Li Master of Science

A thesis submitted for the degree of Doctor of Philosophy at The University of Queensland in 2018 School of Agriculture and Food Sciences Abstract Crop Zn deficiency is associated with low Zn availability in soils and is a widespread problem, also resulting in human Zn deficiency. Foliar Zn fertilisation has been shown to be an efficient approach for overcoming this problem. However, the mechanisms by which Zn moves across the leaf surface (penetration across the cuticle and epidermal cells and into the underlying tissues) and is subsequently translocated remain unclear, with this being the main focus of the present research. Specifically, using ZnSO4 and nano-ZnO, the roles of leaf cuticle, trichomes, and stomata in foliar Zn absorption were examined, with the distribution and the speciation of the absorbed Zn in the leaves determined. In addition, the influence of plant Zn status on foliar Zn absorption was also examined.

The first experiment aimed to obtain a general understanding of the effects of leaf properties on the foliar absorption of Zn. Methyl jasmonate (MeJA) (0, 0.1, 0.5, 1, and 2.5 mM) was applied to leaves of soybean (Glycine max), sunflower (Helianthus annuus), and tomato (Solanum lycopersicum) to alter leaf properties. It was found for all three plant species that treating of MeJA caused substantial increases (1.3–3.5-times higher) in leaf trichome density, and the hydrophobicity of the adaxial leaf surface. The changes in stomatal density and leaf cuticle thickness varied with plant species and MeJA concentration. Based upon these results, the second experiment used 0 (control) and 1 mM MeJA treatments to examine the effects of changes in leaf properties on the absorption of foliar-applied Zn, Mn, and Fe, using synchrotron-based X-ray fluorescence microscopy (µ-XRF) to examine the distribution of elements in situ within hydrated leaves. Interestingly, foliar absorption of Zn, Mn, and Fe increased up to 3- to 5-fold in sunflower leaves treated with 1 mM MeJA, but decreased by 0.5- to 0.9-fold in leaves of tomato treated with 1 mM MeJA. It was found that these changes were related to the thickness of the cuticle and epidermal cell wall, suggesting that the cuticle is important for absorption of foliar-applied nutrients. However, subsequent translocation of the absorbed Zn, Mn, and Fe within the leaf tissues was limited (moving < 1.3 mm in 6 h) irrespective of the plant species and treatment. Of the three nutrients, Zn was translocated a greater distance within the vein than in the interveinal tissues (being translocated ca. two-fold further).

The third experiment gave specific consideration to the role of trichomes in foliar Zn absorption. Using µ-XRF for the in situ analyses of nutrient distribution in leaves of soybean and tomato, it was found that upon the foliar-application of ZnSO4, Zn accumulated in some

1 non-glandular trichomes of soybean, but not in any trichomes of tomato. However, examining foliar Zn absorption in soybean near-isogenic lines (NILs) differing 10-fold in trichome density found that foliar Zn absorption was not related to leaf trichome density. Therefore, it is suggested that, for soybean and tomato, trichomes are not an important pathway for the movement of foliar-applied Zn across the leaf surface. However, as trichomes are highly diverse, depending upon the plant species and their properties, trichomes could potentially be important for foliar Zn absorption.

Next, the importance of the cuticle, stomata, and trichomes in foliar Zn absorption was compared in sunflower – a species in which trichomes are known to play an important role in metal sequestration when supplied in the rooting environment. Analyses of intact hydrated leaves using µ-XRF showed that foliar-applied Zn accumulated rapidly (≤ 15 min) within non-glandular trichomes (NGTs), with the NGTs having the highest Zn concentrations within the leaf tissue. Examining cross sections of the leaf, it was confirmed that the NGTs of sunflower are indeed important for accumulation of the foliar applied Zn. Interestingly, Zn was found to accumulate rapidly in trichomes on both the adaxial and abaxial leaf surfaces for sunflower leaves, despite the Zn being only applied to the adaxial surface, with this translocation seemingly due to transport through bundle sheath extensions. For sunflower, not only was Zn found to accumulate rapidly in the NGTs, but using nanoscale secondary ion mass spectrometry (NanoSIMS), it was found that the cuticle also has a role in Zn foliar absorption, with Zn appearing to move across the cuticle before accumulating in the walls of the epidermal cells. In contrast, however, no marked accumulation of Zn was found within the stomatal cavity, indicating that movement of Zn through the stomata was not likely to be important. However, it is necessary to extend the findings of this study to other plant species and other forms of Zn fertilisers which have different chemical properties.

A fifth experiment using sunflower examined the influence of the plant Zn status on foliar Zn absorption. It was found that the absorption of Zn in Zn deficient sunflower was 50-66% lower than for Zn sufficient sunflower. However, in situ µ-XRF analyses showed that the pattern of Zn absorption was similar for both plants, indicating the mechanisms of Zn foliar absorption in Zn sufficient and deficient sunflower were likely the same. Rather, Zn deficiency decreased the leaf trichome density and altered the chemical composition of the cuticle and epidermal layer, with these presumably being the main causes of the reduced Zn absorption in Zn deficient sunflower. In addition, in situ synchrotron-based X-ray absorption

2 spectroscopy (XAS) analyses showed that the Zn in the leaves following foliar application was mostly present as Zn phytate (37-53%) in Zn sufficient leaves, but as Zn phosphate (about 55%) in Zn deficient leaves.

Across several of the experiments described above, the foliar absorption of ZnSO4 and nano-

ZnO were compared. It was found that the absorption of Zn when supplied as ZnSO4 was ca. 10-fold higher than when supplied as nano-ZnO in soybean, and ca. 100-fold higher in sunflower after 6 h. Using µ-XRF analysis, it was found that their pattern of absorption did not differ between the two types of fertilisers. Furthermore, using XAS analysis, it was found that the speciation of the Zn within the leaf tissue following absorption was similar. These results suggest that the nano-ZnO was absorbed as soluble Zn ions rather than as intact nanoparticles, and nano fertilisers could potentially be used as a slow release foliar fertiliser provided they can be retained on the leaf surface during plant growth.

Overall, the cuticle appears to be an important pathway for Zn foliar absorption, although depending upon the plant species and the properties of the trichomes, it is possible that trichomes are also important for foliar Zn absorption. Regardless of the plant species or the form of Zn supplied, the subsequent translocation of bulk Zn after its initial absorption was limited. The Zn status of sunflower plants was found to influence the extent of the absorption of foliar-applied Zn, with Zn deficiency leading to decreased absorption of foliar applied Zn.

Compared with ZnSO4, the absorption of nano-ZnO was markedly lower despite their distribution and speciation within the leaf tissues being similar, suggesting that nano-ZnO was absorbed as soluble Zn rather than as intact nanoparticles. However, if nano-ZnO can be retained on the leaf surface, it would potentially have great value as a slow release fertiliser to provide sustained release.

Declaration by author This thesis is composed of my original work, and contains no material previously published or written by another person except where due reference has been made in the text. I have clearly stated the contribution by others to jointly-authored works that I have included in my thesis.

I have clearly stated the contribution of others to my thesis as a whole, including statistical assistance, survey design, data analysis, significant technical procedures, professional editorial advice, financial support and any other original research work used or reported in my thesis. The content of my thesis is the result of work I have carried out since the commencement of my higher degree by research candidature and does not include a substantial part of work that has been submitted to qualify for the award of any other degree or diploma in any university or other tertiary institution. I have clearly stated which parts of my thesis, if any, have been submitted to qualify for another award.

I acknowledge that an electronic copy of my thesis must be lodged with the University Library and, subject to the policy and procedures of The University of Queensland, the thesis be made available for research and study in accordance with the Copyright Act 1968 unless a period of embargo has been approved by the Dean of the Graduate School.

I acknowledge that copyright of all material contained in my thesis resides with the copyright holder(s) of that material. Where appropriate I have obtained copyright permission from the copyright holder to reproduce material in this thesis and have sought permission from co- authors for any jointly authored works included in the thesis.

Publications during candidature 1. Li C, Wang P, Menzies NW, Lombi E, Kopittke PM (2018) Effects of methyl jasmonate on plant growth and leaf properties. Journal of Plant Nutrition and Soil Science 181: 409-418.

2. Li C, Wang P, Menzies NW, Lombi E, Kopittke PM (2017) Effects of changes in leaf properties mediated by methyl jasmonate (MeJA) on foliar absorption of Zn, Mn and Fe. Annals of Botany 120: 405-415.

3. Li C, Wang P, Lombi E, Cheng M, Tang C, Howard DL, Menzies NW, Kopittke PM (2018) Absorption of foliar applied Zn fertilizers by trichomes in soybean and tomato. Journal of Experimental Botany 69: 2717-2729.

4. Li C, Wang P, Van der Ent A, Cheng M, Jiang H, Read TL, Lombi E, Tang C, de Jonge M, Menzies NW, Kopittke PM (2018) Absorption of foliar-applied Zn in sunflower (Helianthus annuus): Importance of the cuticle, stomata, and trichomes. Annals of Botany. DOI: 10.1093/aob/mcy135.

5. Li C, Wang P, Lombi E, Wu J, Blamey FPC, Fernández V, Daryl HL, Menzies NW, Kopittke PM (2018) Absorption of foliar applied Zn reduced in Zn deficient sunflower (Helianthus annuus) due to the changing of leaf properties. Plant and Soil 433: 309-322.

6. Blamey FPC, McKenna BA, Li C, Cheng M, Tang C, Jiang H, Howard DL, Paterson DJ, Kappen P, Wang P, Menzies NW, Kopittke PM (2017) Manganese distribution and speciation help to explain the effects of silicate and phosphate on manganese toxicity in four crop species. New Phytologist 217: 1146-1160.

7. Cicchelli FD, Wehr JB, Dalzell SA, Li C, Menzies NW, Kopittke PM (2016) Overhead-irrigation with saline and alkaline water: Deleterious effects on foliage of Rhodes grass and leucaena. Agricultural Water Management 169: 173-182.

8. Doolette CL, Read TL, Li C, Scheckel KG, Donner E, Kopittke PM, Schoerring JK, Lombi E (2018) Foliar application of zinc sulfate and zinc EDTA to wheat

leaves: differences in mobility, distribution and speciation. Journal of Experimental Botany 69: 4469-4481.

9. Wang P, McKenna BA, Menzies N, Li C, Glover CJ, Zhao FJ, Kopittke PM (2018) Minimizing experimental artefacts in synchrotron-based X-ray analyses of Fe speciation in plant tissues. Journal of Synchrotron Radiation: Under Review.

Publications included in this thesis 1. Li C, Wang P, Menzies NW, Lombi E, Kopittke PM (2018) Effects of methyl jasmonate on plant growth and leaf properties. Journal of Plant Nutrition and Soil Science 181: 409-418. 2. Li C, Wang P, Menzies NW, Lombi E, Kopittke PM (2017) Effects of changes in leaf properties mediated by methyl jasmonate (MeJA) on foliar absorption of Zn, Mn and Fe. Annals of Botany 120: 405-415.

- incorporated as Chapter 3. Contributor Statement of contribution Author Li C (Candidate) Designed and performed experiments (60%) Analysis and interpretation (60%) Drafting and production (60%) Author Kopittke PM and Wang P Designed and performed experiments (40%) Analysis and interpretation (30%) Drafting and production (30%) Author Lombi E and Menzies NW Analysis and interpretation (10%) Drafting and production (10%)

- incorporated as Chapter 4. Contributor Statement of contribution Author Li C (Candidate) Designed and performed experiments (65%)

Analysis and interpretation (65%) Drafting and production (65%) Author Kopittke PM and Wang P Designed and performed experiments (25%) Analysis and interpretation (35%) Drafting and production (30%) Author Lombi E and Menzies NW Drafting and production (10%) Author Cheng M, Tang C, Lombi E and Performed experiments (10%) Howard DL

- incorporated as Chapter 5. Contributor Statement of contribution Author Li C (Candidate) Designed and performed experiments (55%) Analysis and interpretation (70%) Drafting and production (60%) Author Kopittke PM Designed and performed experiments (20%) Analysis and interpretation (20%) Drafting and production (25%) Author Wang P, Van der Ent A, Jiang H, Performed experiments (15%) and Lombi E, Analysis and interpretation (10%) Drafting and production (10%)

Author Cheng M, Jiang H, Tang C, de Performed experiments (10%) Jonge M, and Menzies NW Drafting and production (5%)

5. Li C, Wang P, Lombi E, Wu J, Blamey FPC, Fernández V, Daryl HL, Menzies NW, Kopittke PM (2018) Absorption of foliar applied Zn reduced in Zn deficient

sunflower (Helianthus annuus) due to the changing of leaf properties. Plant and Soil 433: 309-322.

- incorporated as Chapter 6. Contributor Statement of contribution Author Li C (Candidate) Designed and performed experiments (70%) Analysis and interpretation (70%) Drafting and production (65%) Author Kopittke PM Designed and performed experiments (20%) Analysis and interpretation (20%) Drafting and production (20%) Author Wang P, Lombi E, Fernández V, Drafting and production (15%) and Menzies NW Author Wu J and Daryl HL Performed experiments (10%)

Contributions by others to the thesis My supervisors Associate Professor Peter Kopittke, Professor Peng Wang, Professor Enzo Lombi, and Professor Neal Menzies have provided valuable advice and guidance on all work presented in this thesis, including designing the experiments, data interpreting, and editing my drafts. Associate Professor Pax Blamey has provided suggestions and assistance on my experiment work.

Professor Caixian Tang, and Dr Miamiao Cheng has provided assistance on part of experiment work in Chapter 4 and Chapter 5. Part of the experiment from Chapter 5 was collaborating with Dr Antony van der Ent and Dr Haibo Jiang. Dr Victoria Fernández gave valuable suggestions for Chapter 5 and helped editing Chapter 6.

Statement of parts of the thesis submitted to qualify for the award of another degree None Research Involving Human or Animal Subjects No animal or human subjects were involved in this research

Acknowledgements I would like to express my sincere gratitude to my supervisors Associate Professor Peter Kopittke, Professor Peng Wang, Professor Enzo Lombi, and Professor Neal Menzies for the valuable guidance, advice, inspiration, support, and patience that they give to me, especially to my professional, responsible, and supportive principal supervisor Peter who is always there to listening and feedback us in a timely manner. I have learned from him not only knowledge, but his way of thinking and work. It is indeed a privilege to having him as my PhD supervisor.

My sincere thanks also goes to Associate Professor Pax Blamey, he has generously shared me his experience and knowledge. I would also like to thank Dr Victoria Fernández who has kindly gave me suggestions and greatly helped me on Chapter 6. The assistance of Professor Caixian Tang, Dr. Miamiao Cheng, Dr. Haibo Jiang, Dr Brett Ferguson, Dr Antony van der Ent, and Jingtao Wu in my experimental work are also much appreciated. I also thank my milestone examiners Dr Ryo Fujinuma, Dr Bernhard Wehr, Dr Antony van der Ent and Associate Professor Longbin Huang for their suggestions and time. For the expert technique guidance and assistance I have received, I am highly appreciated the help from Dr Kathryn Green, Dr Kim Sewell, Mr Ron Rasch, Dr Erica Lovas, Mr Richard Webb, Ms Ying Yu, Dr Robert Sullivan, Dr Daryl Howard, and Dr Martin de Jonge.

Assistance provided to me by the staff of The University of Queensland are also thanked, especially to Dr Brigid McKenna, Ms Kerry Vinall, and Ms Katy Carroll for their technical services and the postgraduate administrator Ms Kaye Hunt. Thanks also to my student colleagues Lei Xiong, Christian Forster, Yunru Lai, Zhigen Li, Jingtao Wu, Robert Banks, Yaqi Zhang, Chelsea Stroppiana, Monia Anzooman, Sara Niaz, Alessandro Zucchelli, and Thea Lund Read for their companion, help and support.

Finally, to allow me have such a wonderful experience, I want to thank the financial support from Sonic Essentials and the Australian Research Council (ARC) for my PhD project as well as the China Scholarship Council and The University of Queensland for providing me the scholarships.

Financial support Financial support was provided by Sonic Essentials. This research was also supported by Australian Research Council (ARC) through the Linkage Projects funding scheme (LP130100741). Support was also provided to the Candidate through the China Scholarship Council (CSC) and The University of Queensland. A large portion of the experimental work was conducted at the XAS and XFM beamlines of the Australian Synchrotron, part of Australian Nuclear Science and Technology Organisation (ANSTO).

Keywords Zinc, foliar fertiliser, nano-fertiliser, foliar absorption, cuticle, trichomes, stomata, translocation

Australian and New Zealand Standard Research Classifications (ANZSRC) ANZSRC code: 070306, Crop and Pasture Nutrition, 50% ANZSRC code: 060705, Plant Physiology, 30% ANZSRC code: 050304, Soil Chemistry, 20%

Fields of Research (FoR) Classification FoR code: 0703, Crop and Pasture Production, 50% FoR code: 0607, Plant Biology, 30% FoR code: 0503, Soil Science, 20%

Table of Contents Abstract …………………………………………………………………………………….... 1 List of Abbreviations …………………………………………………………………..…...13

Chapter 1 Introduction ...... 14

Chapter 2 Review of literature ...... 15

2.1 Zinc status in soils and implications for plants and humans ...... 15

2.1.1 Zinc in soils and availability to plants ...... 15

2.1.2 Zinc in plants ...... 16

2.1.3 Zinc deficiency in crops and humans ...... 17

2.2 Overcoming Zn deficiency in crops ...... 19

2.2.1 Limitations of soil-application of Zn fertilisers ...... 20

2.2.2 Advantages of foliar-application of Zn fertilisers ...... 22

2.3 Absorption of the foliar applied nutrients ...... 22

2.3.1 Cuticle and foliar absorption ...... 23

2.3.2 Stomata and foliar absorption ...... 27

2.3.3 Trichomes and foliar absorption ...... 30

2.4 Translocation of foliar applied nutrients ...... 32

2.5 Effects of nutrient status on foliar nutrient absorption ...... 33

2.6 Use of nanomaterials as fertilisers ...... 34

2.7 Research questions ...... 35

Chapter 3 Effects of changes in leaf properties mediated by methyl jasmonate (MeJA) on foliar absorption of Zn, Mn, and Fe ...... 37

3.1 Introduction ...... 37

3.2 Paper 1 ...... 37

3.3 Paper 2 ...... 66

3.4 Conclusions ...... 101

Chapter 4 Absorption of foliar-applied Zn fertilizers by trichomes in soybean and tomato ...... 102

4.1 Introduction ...... 102

4.2 Paper 3 ...... 102

Chapter 5 Absorption of foliar-applied Zn in sunflower (Helianthus annuus): Importance of the cuticle, stomata, and trichomes ...... 143

5.1 Introduction ...... 143

5.2 Paper 4 ...... 143

Chapter 6 Absorption of foliar applied Zn is decreased in Zn deficient sunflower (Helianthus annuus) due to changes in leaf properties ...... 184

6.1 Introduction ...... 184

6.2 Paper 5 ...... 184

Chapter 7 General discussion and conclusions ...... 219

7.1 Absorption of foliar applied Zn ...... 219

7.1.1 The cuticle is an important pathway for foliar Zn absorption ...... 219

7.1.2 Some trichomes could be important for foliar Zn absorption ...... 220

7.1.3 Stomata appear to not be an important pathway for foliar Zn absorption ...... 222

7.1.4 Foliar Zn absorption is decreased in Zn deficient sunflower ...... 222

7.2 Comparison of the foliar absorption of ZnSO4 and nano-ZnO ...... 223

7.3 Translocation of the foliar absorbed Zn ...... 224

7.4 Conclusions ...... 225

References…………………………………………………………………………………..226

List of Abbreviations µ-XRF X-ray fluorescence microspectroscopy

BSEs Bundle sheath extensions

DW Dry weight

Fe Iron

FTIR Fourier transform infrared spectroscopy

ICP–MS Inductively coupled plasma mass spectrometry

ICP-OES Inductively coupled plasma optical emission spectroscopy

JA Jasmonic acid

LCF Linear combination fitting

MeJA Methyl jasmonate

Mn Manganese

NanoSIMS Nanoscale secondary ion mass spectrometry

NILs Near-isogenic lines

RH Relative humidity

SEM Scanning electron microscopy

XANES K-edge X-ray absorption near edge structure spectra

XFM X-ray fluorescence microscopy

YFELs Youngest fully-expanded leaves

Zn Zinc

Chapter 1 Introduction Zinc (Zn) is an essential nutrient for both plants and humans (Broadley et al., 2007, Tuerk et al., 2009), but many soils worldwide contain inadequate levels of this nutrient (Alloway, 2008, Cakmak, 2000). As a result, it is estimated that crops growing on 49% of the world’s important agricultural soils and one third of the world’s population suffer from Zn deficiency (Sillanpää, 1990, Cakmak et al., 2010). Foliar fertilisation has been proven as a biofortification strategy to overcome Zn deficiency, especially when plants are at the stage where Zn is in high demand or when the soil-application of Zn fertilisers is ineffective due to edaphic factors such as high pH, high carbonate contents and organic matter contents (Fernández et al., 2013a, Prasad et al., 2014). New functional fertilisers, for example, foliar fertilisers with sustained and controlled release, are needed. Nano-fertilisers are promising in this regard given that they can be easily functionalised and potentially release nutrients to crops in a controlled manner, including sustained rates and target-specific effects (Liu et al., 2015).

Currently, the processes regarding the foliar absorption of nutrients by plants and the translocation of the foliar absorbed nutrients in plants are not well understood. Particularly, the potential roles of the cuticle, trichomes, and stomata on the foliar absorption of Zn remain unclear (Fernández et al., 2013b). As a consequence, current foliar fertilisers are not designed on a physiological basis, and hence suffer from various limitations. For example, soluble Zn salts carry a high risk of scorching the leaves (Alloway, 2008) and chelated forms of Zn such as Zn-EDTA are associated with environmental concerns (Oviedo et al., 2003). The present study aims to investigate the process of the absorption of the foliar applied Zn, with the redistribution of Zn after absorption also considered. These factors will be investigated using traditional approaches as well as synchrotron-based X-ray fluorescence microscopy (µ-XRF), synchrotron-based X-ray absorption spectroscopy (XAS), and nanoscale secondary ion mass spectrometry (NanoSIMS) – these allowing for in situ analyses. The outcomes of this work will assist in improving the understanding of foliar Zn absorption and translocation, thereby promoting the design of physiologically-based Zn foliar fertilisers and simultaneously minimising the threat to food security and human health.

Chapter 2 Review of literature

2.1 Zinc status in soils and implications for plants and humans

2.1.1 Zinc in soils and availability to plants The average concentration of Zn in the Earth’s crust is 75 mg kg-1, with the typical concentration of total Zn in soils ranging 10 to 300 mg kg-1 (Swaine, 1956, Barak et al., 1993). The concentration of Zn in soil depends upon the parent material (for example, sandy soils tend to have low Zn concentrations while soils with higher clay contents tend to have higher Zn concentrations), inputs from atmospheric deposition, agricultural fertilisers (e.g. livestock manures, inorganic fertilisers, agrochemicals, and sewage sludge), and industrial waste products (Alloway, 2008). Zinc exists in soil in three primary forms: (1) soluble Zn, including free ions (Zn2+ and ZnOH+) and weakly complexed Zn; (2) exchangeable Zn adsorbed on surfaces of the colloidal fraction (associated with clay particles, organic matter, and Al and Fe hydroxides); and (3) secondary minerals and insoluble complexes (Alloway, 2008). It is reported that soils containing less than 0.5 mg Zn kg-1 as DTPA-extractable Zn are potentially Zn deficient and tend to respond well to Zn fertilisers (Lindsay et al., 1978).

Only free Zn2+ ions and labile Zn weakly complexed with inorganic or organic ligands are available to plant roots (Broadley et al., 2007, Alloway, 2008). Hence, Zn deficiency in plants is not only a result of the low total Zn content in soils, but also the low bioavailability of Zn for plants (Alloway, 2009). For example, a range of edaphic factors affect the availability of Zn, including soil pH, organic matter content, clay content, calcium carbonate content, redox condition, rhizospheric microbial activity, soil moisture, climate, and coexisting cations and 3- anions (e.g. PO4 ) (Alloway, 2008, Sadeghzadeh, 2013, Rehman et al., 2018). Indeed, plant Zn deficiency often occurs in calcareous soils, sandy soils, strongly weathered deep tropical soils, saline and sodic (salt affected) soils, and poorly drained/waterlogged soils. Among these soils, calcareous soils tend to result in the most severe Zn deficiency (Alloway, 2008).

The first identification of plant Zn deficiency in field conditions was reported in 1937 in the deciduous orchards of California, USA (Robson, 2012). Since then, considerable progress has been made in identifying the chemical forms and physiological functions of Zn in soils and plants. Low levels of plant-available Zn in soil cause plant Zn deficiency, and the resultant Zn deficiency in humans is now recognised as one of the most common micronutrient

15 deficiencies worldwide (Alloway, 2008, Du et al., 2015, Rehman et al., 2018). Indeed, it is estimated that 49% of the world’s important agricultural soils and nearly half of all cereals contain low levels of plant-available Zn (Sillanpää, 1990, Cakmak, 2008), mainly located in Afghanistan, Australia, Bangladesh, Brazil, China, India, Iran, Iraq, Pakistan, Philippines, Sudan, Syria, Turkey, many states in the USA, and parts of Africa and Europe (Alloway, 2008). Consequently, the low levels of plant-available Zn in soils are causing the world’s most common micronutrient deficiency in crops (ZNI, 2014, Rehman et al., 2018).

2.1.2 Zinc in plants 2.1.2.1 Functions of Zn in plants As an essential micronutrient for plants, Zn plays catalytic, structural, and regulatory roles in a wide range of biochemical pathways (Palmer et al., 2009, Lanquar et al., 2014, Caldelas et al., 2017), with Zn being the only element that is present in all six enzyme classes (oxidoreductases, transferases, hydrolases, lyases, isomerases, and ligases) in plants (Broadley et al., 2007, Alloway, 2008). Specifically, the primary functions of Zn in plants are: (1) carbohydrate metabolism, for example, Zn is important in photosynthesis and in the conversion of sugars to starch (Ohki, 1976, Samreen et al., 2017, Zhang et al., 2017); (2) protein metabolism, with Zn being essential in chromatin structure, DNA/RNA metabolism and gene expression (Sadeghzadeh, 2013); (3) auxin metabolism, with Zn associated with growth regulator (Begum et al., 2016); (4) pollen formation (Pandey et al., 2006); and (5) maintenance of the integrity of biological membranes (Alloway, 2008).

2.1.2.2 Forms and distribution of Zn in plants Within plants, Zn is not redox active and does not undergo valency changes, with Zn2+ ions having Lewis acid characteristics and having a flexible coordination sphere (Palmer et al., 2009, Laitaoja et al., 2013). Zinc is typically present within plants as soluble forms (including free Zn2+ ions and those associated with low molecular weight organic ligands), metalloproteins, or insoluble forms associated with the cell walls (Alloway, 2008). Depending on the plant species, the shoot Zn concentration can vary from 15-300 mg kg-1 (Caldelas et al., 2017), and the water-soluble form may account for 58-91% of the total Zn in the plant. This water-soluble fraction is regarded as a better indicator for plant Zn status than total Zn contents (Brown et al., 1993), and it is also considered to be the most physiologically active as the water-soluble fraction is moveable and may involve in homeostatic mechanisms by

16 which it may act as a buffer system to bind excess free Zn ions (Alloway, 2008). Within the leaves, it has been shown that in the hyperaccumulator plant Thlaspi caerulescens, Zn accumulates in the vacuoles of epidermal cells and cell walls of the inner leaf cells (Frey et al., 2000). However, due to detection limitations, it is not yet known what proportion of plant cytoplasmic Zn is present as free Zn2+ or as Zn bound to protein, amino acid, nucleotide or lipid ligands, or compartmentalised into organelles (Broadley et al., 2007).

2.1.3 Zinc deficiency in crops and humans 2.1.3.1 Consequences and symptoms of Zn deficiency in crops The low available Zn in soils worldwide causes Zn deficiency in crops, with this resulting not only in reduced crop yields but also in a reduced nutritional quality of the food (Sadeghzadeh, 2013, Cakmak et al., 2018). As described earlier, Zn is necessary for many physiological and biochemical processes, hence Zn deficiency also results in greatly reduced plant growth. For example, Cakmak et al. (1993) observed that Zn deficiency severely decreased plant growth and leaf concentrations of soluble protein and chlorophyll. With the exception of guaiacol peroxidase, the activities of all antioxidant enzymes were significantly decreased by Zn deficiency, in particular GSSG (oxidized glutathione) and SOD (superoxide dismutase), indicating that Zn deficiency also impairs the ability of the plant to enzymatically scavenge - O2 and H2O2.

Responses to Zn deficiency differ greatly among plant species. Jahiruddin (1986) reported that the normal concentrations of Zn in plants range from 25 to 150 mg Zn kg-1 (dry mass), with deficiencies occurring when leaf concentrations are < 20 mg Zn kg-1 and toxicity when they exceed 400 mg Zn kg-1. Robson (1993) suggested that critical deficiency levels in shoots are in the range of 10 - 15 mg Zn kg-1 dry weight for most graminaceous species, and 20 - 30 mg Zn kg-1 for most dicotyledonous species. Indeed, some plant species are particularly sensitive to Zn deficiency, including maize (Zea mays), sorghum (Sorghum bicolor) and grape (Vitis vinifera), and to a lesser extent for soybean and tomato, while wheat (Triticum) and alfalfa (Medicago sativa) are less sensitive (Moraghan, 1984, Alloway, 2008).

Zinc deficiency can be diagnosed through a series of symptoms in plants, including symptoms in both the older and younger leaves (Weir et al., 1993, Weir et al., 1995, Alloway, 2008). The symptoms generally include: (1) chlorosis (leaves yellowing or even whitening, and often

17 interveinal, due to the reduced amount, or absence of chlorophyll); (2) necrotic spots (occurring in areas of chlorosis due to the death of the leaf tissue in small concentrated areas); (3) bronzing of leaves, associated with chlorosis and the chlorotic areas which may turn bronze coloured; (4) rosetting of leaves (occurring in dicotyledonous crops when the internodes on the stems of dicotyledonous crops fail to elongate normally, clustering leaves on the stem); (5) stunting of plants (as a consequence of reduced dry matter production giving a smaller plant and/or reduced internode elongation of stems of developing crops); (6) dwarf leaves (also called ‘little leaf’); and (7) malformed leaves (Alloway, 2008). However, these symptoms differ substantially among plant species. For example, Cakmak et al. (1997), who observed the first visible symptom of Zn deficiency, reported a reduction in shoot elongation followed by the appearance of whitish-brown necrotic patches on the leaf blades. These symptoms occurred more rapidly and with greater severity in wheat varieties, particularly in durum wheats (Triticum durum), but were either absent or only slight in rye (Secale cereale) and triticale (× Triticosecale Wittmack). The same was observed for the decrease in shoot dry matter production and grain yield.

2.1.3.2 Consequences and status of Zn deficiency in humans Zinc plays an essential role in numerous biochemical pathways in human health, including physical growth and development, immune system, reproductive health, sensory functions, and neurobehavioral development (Hotz et al., 2004, Sharma et al., 2013). The functions of Zn can be broadly divided into three categories: catalytic, structural and regulatory activities (Tuerk et al., 2009). For example, Zn is ubiquitous in subcellular metabolism, and has a specific structural role in enzyme molecules as well as in many proteins and biomembranes (Hambidge, 2000), with 2800–3000 proteins containing a Zn prosthetic group (Tapiero et al., 2003). Apart from that, Zn is an integral component of Zn finger proteins that regulate DNA transcription (Levenson et al., 2011). Additionally, Zn is also essential for regulating intestinal absorption of Fe, with sufficient Zn and Fe being crucial for treating Fe deficient anaemia (Prasad et al., 2014). As discussed for plants, Zn is the only metal to be involved in all six classes of enzymes in the human body (Barak et al., 1993), and it is required for the activation of over 300 enzymes (Gibson, 2012). Finally, Zn is also a component of neurotransmitters, present in the cells of the salivary glands, prostrate, and immune system (Hershfinkel et al., 2007).

There are a large number of studies that have examined the many serious health problems caused by Zn deficiency. For example, it is known that Zn deficiency affects the functions of many organ systems, including the integumentary, gastrointestinal, central nervous system, immune, skeletal, and reproductive systems. Zinc deficiency is also associated with acute and chronic liver diseases and diarrheal disease (Hambidge, 2000, Tuerk et al., 2009). Furthermore, Zn deficiency is also responsible for increased risk of infections, DNA damage, and cancer development. Specifically, it is able to lead to stunting in children (Hotz et al., 2004, Gibson, 2006, Prasad, 2007). Conversely, there is a body of evidence to show Zn supplements can significantly prevent or treat diseases caused by Zn deficiency including immune dysfunction, acute lower respiratory infection, diarrhoea and liver diseases (Sandstead, 1994, Walker et al., 2004, Brooks et al., 2005).

According to a report by the World Health Organization (WHO 2002) on the risk factors responsible for development of illnesses and diseases, Zn deficiency ranks 11th among the 20 most serious factors in the world and 5th among the 10 most important factors in developing countries (Cakmak, 2008). In a similar manner, Hotz et al. (2004) reported that Zn deficiency affects, on average, one-third of the world’s human population, ranging from 4-73% of the population in various countries. The WHO also estimates that 800,000 people die annually due to Zn deficiency, with 450,000 of these being children under the age of five. Finally, studies by the World Bank and other organizations have estimated that micronutrient malnutrition could lead to a reduction in gross domestic product up to 5% globally (IZA, 2015).

2.2 Overcoming Zn deficiency in crops

Fortification of Zn in crops can significantly improve crop yield and nutrient quality as well as human health (Velu et al., 2014, Joy et al., 2015, Zhang et al., 2018). This can be achieved through both genetic practices (e.g. breeding crop cultivars with higher Zn concentration in grains) and agronomic practices (e.g. fertilising crops with Zn). Currently, genetic approaches remain a long term approach (Cakmak, 2008), with application of Zn fertilisers the main solution used at present. For example, the application of various Zn fertilizers at appropriate rates, either to the soil or to the foliage, have been shown to increase crop yield by up to 120% in wheat, 48% in rice (Oryza sativa), 50% in cotton (Gossypium), 18% in maize, 22% in potato (Solanum tuberosum), 30% in alfalfa, 50% in soybean, and 26% in citrus (Citrus

19 reticulatus) (Ahmad et al., 2012). Among different application methods, foliar application of Zn has proven to be an effective strategy to overcome Zn deficiency in crops (Ram et al., 2016), particularly for those soils which limit the uptake of Zn by the roots due to chemical constraints, for example, alkaline soils has a low Zn availability due to precipitation reactions (Cakmak, 2008, Cakmak et al., 2010). Indeed, Cakmak (2008) notes that foliar applications of Zn is the most efficient way to enhance Zn concentration in wheat grains, given that the foliar application is a direct delivery of the micronutrient to the plant, bypassing interactions with the soil phase, where adverse chemical conditions can diminish its bioavailability. The disadvantages and the advantages of soil-application and foliar-application of Zn fertilisers are discussed below.

2.2.1 Limitations of soil-application of Zn fertilisers Roots primarily take up Zn from the soil solution as Zn2+. Zinc deficiency in plants is caused either by inherently low Zn concentrations or low plant available Zn concentrations.

Specifically, a series of factors such as soil pH, redox conditions, calcite (CaCO3) and organic matter contents, concentrations of all ligands capable of forming chelators, clay content, temperature, and microoganisms have impact on soil Zn bioavailability and the mobility of Zn in soils (Alloway, 2008). Indeed, concentrations of Zn in the soil solutions are normally low (4270 µg L-1) compared with average total concentrations of around 64 mg Zn kg-1 (Alloway, 2009). Apart from an inherently low Zn concentration, the main factors limiting soil Zn availability to plants are illustrated below.

Among the soil chemical factors, soil pH has been identified as being a particularly critical parameter regulating micronutrient availability (Sims, 1986, Zeng et al., 2011), and it plays the most important role in regulating Zn solubility in soil solution. This is due to the greater adsorptive capacity of the soil solid surfaces resulting from increased pH-dependent negative charge, which results in formation of hydroxylated forms of Zn, chemisorption on calcite, and/or co-precipitation in Fe oxides (Marschner, 1993, Cakmak, 2008, Kabata-Pendias, 2010). For example, Sims (1986) reported that soil pH markedly altered the distribution of Mn and Zn, with soluble Zn concentration reaching as high as 7,137 µg L-1 (normally 4–270 µg L-1) in very acidic soils (Kabata-Pendias, 2010). Marschner (1993) found that at Zn concentration in soil solution decreased 30-45-fold for each unit as pH increase between 5.5 and 7.0. Therefore, Zn deficiency is more likely to occur at high soil pH values, such as calcareous soils high in CaCO3.

High levels of soil organic matter (e.g. peat or muck soils) are also associated with the development of Zn deficiency in crops. This is owning to that either these organic materials have an inherently low total Zn concentration or Zn combined with some organic materials and formed stable, solid-phase organic complexes (Alloway, 2008, Alloway, 2009). In a similar manner, Dabkowska-Naskret (2003) reported that a significant decrease in Zn was associated with soil organic matter, with the binding of Zn with organic matter being pH dependent. Similarly, Harfouche et al. (2009) found that most Zn was bound with organic matter and Zn complexation with mineral particles might intimately stacked on organic matter particles. However, when rapidly decomposable organic matter, for instance manure, is added to soils, Zn may become more available due to the formation of soluble organic Zn complexes, which are mobile and also probably capable of absorption into plant roots (Alloway, 2008, Zeng et al., 2011).

Soil moisture is another key factor influencing the root Zn uptake process. Transport of Zn to the root surface in soils occurs predominantly via diffusion (Wilkinson et al., 1968), and this process is highly sensitive to soil moisture. Soil moisture is a key physical factor providing a suitable medium for adequate Zn diffusion to plant roots. Low soil moisture can reduce the solubility and mobility of Zn. The plant available Zn in soils, therefore, is adversely affected under water stressed conditions (Cakmak, 2008). However, in prolonged waterlogged soils (e.g. paddy rice soils), the reducing conditions result in a rise in pH and high concentrations of bicarbonate ions, and the formation of insoluble Zn sulphide (ZnS) (Alloway, 2008). In periodically waterlogged soils, the reducing conditions also give rise to increased concentrations of divalent ferrous (Fe2+) and manganese (Mn2+) ions (from the dissolution of their hydrous oxides), which could compete with Zn ions for uptake into roots (Alloway, 2008).

Finally, Zn also interacts with other nutrients (Mousavi et al., 2012, Bouain et al., 2014, Khan et al., 2014, Briat et al., 2015). Specifically, it has been reported that large applications of P can causes Zn deficiency in plants (Robson et al., 1983, Soltangheisi et al., 2013). This could be due to (1) the formation of insoluble Zn3(PO4)2 in the soil which decreased the plant available Zn concentration (Soltangheisi et al., 2013); and (2) the uptake and transport of Zn and P in plants has a coordinating mechanism (Briat et al., 2015, Lei, 2018). Similarly, Zn

21 deficiency in crops also occurs after the application of a copper (Cu) fertiliser, due to the competition between Cu and Zn for the uptake sites (Mousavi et al., 2012).

2.2.2 Advantages of foliar-application of Zn fertilisers As described earlier, the bioavailability (and hence economic efficiency of soil-applied Zn fertiliser) can potentially be reduced due to interactions with the solid-phase. In contrast, foliar fertilisation is an efficient approach for crop Zn biofortification as it is more immediate and target-oriented than soil fertilisation, with nutrients directly delivered to plant tissues during critical stages of plant growth (Fernández et al., 2013b). Indeed, as Cakmak (2008) notes, foliar applications of Zn have proven to be the most efficient way to enhance Zn concentration in wheat grain. Cakmak et al. (2010) later found that when Zn is sprayed onto to wheat leaves at the late growth stage (e.g., milk and dough), Zn concentrations of whole grain and grain fractions are significantly increased. Also, Tian et al. (2015) reported that foliar application of Zn resulted in a greater improvement in Zn densities in rice and wheat grain when compared with soil applied Zn. Shivay et al. (2010a) and Shivay et al. (2010b) found that agronomic efficiency of Zn with foliar application was about four times of that for soil application and importantly rate of Zn application was much lower when applied on foliage compared to soil application.

Therefore, Zn foliar fertilisers are potentially useful, especially (1) when soil conditions limit availability of soil applied Zn; (2) in conditions when high loss rates of soil applied Zn may occur, or (3) at the stage of plant growth when the internal plant demands and the environmental conditions interact to limit delivery of Zn to critical plant organs (Fernández et al., 2013a). However, the process of foliar Zn absorption by plants and the subsequent translocation are far from comprehensively understood. As a consequence, current foliar fertilisers are not tailored on the basis of physiological processes and hence suffer various limitations.

2.3 Absorption of the foliar applied nutrients

The absorption of foliar applied nutrients is a complex process, which is affected by the characteristics of foliage (leaf shape, age and surface characters), physico-chemical properties of the fertiliser (concentration, molecular size, solubility, pH, suspending media), plant physiological status (growth stage, aggregate demand, and mobility of the nutrient within the

22 plant), and ambient environment conditions (temperature, humidity and light) (Fernández et al., 2009, Fernández et al., 2013b).

Unlike roots, the leaf surface is covered by a lipophilic cuticle which protects the plant from uncontrolled water loss, against attachment of pathogens, and particles (Schreiber et al., 2009). According to the structure of the leaf surface, it has been proposed that the absorption of nutrients across the plant surface may potentially occur via: (1) the cuticle, (2) cuticular cracks and imperfections, or (3) stomata, trichomes, lenticels (Fernández et al., 2013b). Lenticels are mainly present on stems, pedicels, old roots or fruit surfaces (Bezuidenhout et al., 2005). For most leaves, the cuticle, stomata (including the pore and guard cells) and trichomes (including trichome base and the protruding trichomes) could represent pathways by which foliar applied nutrients can penetrate into leaf tissues. To illustrate the potential contribution of these pathways in nutrient absorption, it is necessary to understand the structural and physico-chemical basis of the cuticular leaf surfaces. Properties of these organs and their potential roles in foliar absorption process are presented below.

2.3.1 Cuticle and foliar absorption 2.3.1.1 Leaf cuticle Plants have evolved a series of physical mechanisms to adapt to the environment. On its aerial surface, plants have developed a cuticle, which is essential in (1) controlling the passage of water vapour and gases; (2) restricting the loss of nutrients, metabolites and water from plants to the environment; (3) protecting the plants from insects and pathogens; (4) limiting the entrance of xenobiotics; and (5) providing a self-cleaning mechanism (the lotus effect) (Schreiber et al., 2009, Yeats et al., 2013a, Fernández et al., 2017). The plant cuticle is an extracellular polymer membrane, which covers all aerial primary organs such as stems, leaves, flowers and fruits. The hydrophobic cuticle often possesses modified epidermal cells such as trichomes or stomata, and is covered by waxes that may confer a hydrophobic character to the plant surface. The characteristics of the plant leaf cuticle have been reported to be related with the absorption of foliar applied nutrients (Fernández et al., 2013b, Fernández et al., 2017).

The plant cuticle is a polymer membrane which is heterogeneous in both chemical composition and structure (Schreiber et al., 2009). Generally, the cuticle consists of cutin (or in some species cutan), waxes, phenolics and polysaccharides (Domínguez et al., 2011). Cutin

23 is a polymer matrix formed by polyhydroxylated C16, C18 fatty acids and/or glycerol cross- linked with ester bonds. The waxes are composed of mostly mixtures of C20 – C40 n-alcohols, n-aldehydes, n-alkanes, and very long chain fatty acids. Phenolics are principally cinnamic acids and flavonoids, while polysaccharides in the cuticle are similar to those found in the plant cell wall, which mainly consist of cellulose, hemicellulose, or pectin (Caffall et al., 2009, Domínguez et al., 2011, Guzmán et al., 2014c). Based on their histochemical staining and presumed chemical composition, Yeats et al. (2013a) classified cuticles into two domains from the internal to the external surface : (1) cuticular layer: a cutin-rich domain embedded with polysaccharides, intracuticular waxes (waxes deposited within the cutin matrix), and phenolics; and (2) cuticle proper: an overlying layer that is less abundant in polysaccharides, but enriched in intracuticular waxes and epicuticular waxes (waxes accumulated on its surface as epicuticular wax crystals, or films), with phenolics also embedded in cutin. However, although the cuticle is usually considered separately from the underlying polysaccharide epidermic cell walls, the two structures are physically related and have some overlapping functions. Indeed, it has been reported that cellulose and pectin are present within the cuticle proper (Guzmán et al., 2014a). Moreover, Guzmán et al. (2014b) observed a cellulose network resembling the cell walls with different structural patterns in the regions ascribed to the cuticle proper and cuticular layer, after cutin depolymerisation of the isolated leaf cuticles. Furthermore, Domínguez et al. (2011) proposed that polysaccharides play a crucial role in the cuticle rheological behaviour. Therefore, it has been recently suggested that the current model of the cuticle needs to be revised, and it has been suggested that the cuticle may be a specialised lipidic modification of the epidermis cell walls, just as lignification is a common modification of plant secondary cell walls (Yeats et al., 2013a, Guzmán et al., 2014b).

Depending on the plant species and ontogeny, the cuticle varies considerably in architecture and composition, and also differ dramatically in thickness, from the nanometer to the micrometer scale (Jeffree, 2006). In addition, cuticles vary in response to environmental and physiological conditions during growth and development, although the chemical composition and structure of the cuticles of most plant species and organs remains unclear (Fernández et al., 2013a). Guzmán et al. (2014b) observed the cuticle thickness of ranged from 5.5-9.5 μm for adaxial leaves of Eucalyptus camaldulensis and 2.5-5.0 μm for E. globulus, with differences also observed in cutin components from the isolated cuticles. In a study in which Arabidopsis (Arabidopsis thaliana) was subjected to water stress, subirrigation of sodium chloride (NaCl), and exogenous sprayed abscisic acid (ABA) treatments, Kosma et al. (2009)

24 found that these treatments induced changes in the synthesis of waxes. Specifically, these stress treatments led to increases in cuticular wax per unit area by 32-80%, primarily due to increases in wax alkanes by 29-98%. In contrast, only water deficit increased the total cutin monomer amount (by 65%), whereas abscisic acid had little effect on cutin composition. Whitecross et al. (1972) and Baker (1974) showed that light reduction and shading during plant growth caused a decrease in the amount of wax per leaf area. In general, however, how cuticles respond to environmental and/or physiological conditions are still poorly understood.

2.3.1.2 Nutrient absorption through the cuticle The movement of nutrients into the plant through the cuticle has been studied for more than a century. The commercial significance of foliar sprays of plant herbicides, fertilisers, and plant growth regulators has resulted in a substantial interest in cuticular permeability (Fernández et al., 2013a). The penetration of nutrients through the leaf surface has been demonstrated in both living plants and isolated cuticle membranes (Koontz et al., 1957, Schlegel et al., 2001, Riederer et al., 2006, Fernández et al., 2009, Du et al., 2015). The cuticle is a lipid-rich layer with its outer compartment dominated by waxes (a mixture of aliphatic hydrocarbons and their derivatives) (Koch et al., 2008, Fernández et al., 2013b). These characteristics make it an effective barrier for the penetration of hydrophilic, polar compounds, although lipophilic and apolar compounds may cross the hydrophobic cuticle by diffusion. Therefore, two different pathways have been identified which are responsible for cuticular penetration: (1) the lipophilic pathway for non-ionic, apolar, and lipophilic compounds (e.g. pesticides and herbicides); and (2) the hydrophilic pathway for ionic, polar, hydrophilic substances (e.g. Ca, K and glyphosate salts) (Fernández et al., 2013b, Du, 2014).

2.3.1.2.1 Lipophilic pathway for non-ionic, apolar, and lipophilic compounds It is generally accepted that the lipophilic pathway is a physical diffusion process (Fernández et al., 2013a), which can be understood as an aqueous donor compartment (high concentration) across the cuticular membrane into an aqueous receiver compartment (low concentration). At molecular levels, the diffusion of a molecule in the cuticle can be seen as passing into and between voids in the polymer matrix arising by molecular motion (Elshatshat et al., 2007). Researchers have shown that the penetration of lipophilic substances was highly size selective with the size selectivity decreasing with increasing temperature (Fernández et al., 2013b, Buchholz et al., 1998). For example, a four-fold increase in molecular weight resulted in a decrease in mobility by a factor of >1000 (Buchholz et al., 1998, Schreiber,

2005). It was also found that extraction of cuticular waxes or an increase in temperature significantly increased the permeability to lipophilic molecules (Baur et al., 1997, Schönherr et al., 1989, Schreiber, 2002, Schreiber, 2005). This solution-diffusion model has been widely studied and is theoretically well-established. The model can predict and measure the permeability of a specifically given lipophilic compound through a particular cuticular membrane as per the equation below (Schönherr et al., 1989, Fernández et al., 2009, Fernández et al., 2013b, Riederer et al., 2006). 푃 = 퐷 ∗ 퐾 ∗ 푙−1 Where: P (m s-1) is the permeance; D (m2 s-1) is the diffusion coefficient in the cuticle (i.e. mobility of the compounds in the cuticle); K is the partition coefficient (solubility of the compounds in the cuticle), which is the ratio between the equilibrium molar concentrations in the cuticle and in the solution at the cuticle surface; and 푙 (m) refers to the path length of diffusion through the cuticle (Fernández et al., 2013b).

2.3.1.2.2 Hydrophilic pathway for ionic, polar, and hydrophilic substances In contrast, the penetration pathway of hydrophilic, polar solutes through the cuticle are not yet fully understood (Fernández et al., 2013a, Fernández et al., 2017). Hydrated ions and inorganic salts are insoluble in lipid phases such as cutin and cuticular waxes. Physically, the penetration pathway for hydrophilic substances is distinctive from the lipophilic pathway. The hypothesis of ‘polar pores’, which exist on the cuticle for the penetration of the hydrophilic substances was proposed by Schönherr (1976) who believed that most of the cuticle surface was covered by lipophilic domains, but polar domains were also present in the cuticle to a certain degree.

A series of experiments have demonstrated the existence of the putative ‘polar pores’. For example, it has indicated that an increase in temperature, and accelerators such as tributyl phosphate and diethyl sebacate can substantially increase the fluidity of amorphous waxes and cutin (Schönherr, 2000), but studies using isolated cuticular membranes showed that the penetration of inorganic ions and charged organic molecules was not affected by temperature or accelerators (Schönherr, 2000, Schönherr et al., 2001). Moreover, size selection for polar substances is substantially lower. For example, size selectivity for the penetration of Ca2+ across an isolated cuticle was markedly less pronounced than that for lipophilic molecules (Schönherr, 2000). The penetration of ions is also influenced by humidity (Schönherr, 2000, Schlegel et al., 2001) as the swelling of the aqueous pores is humidity dependent (Schönherr,

2000). It was also proposed from studies on isolated cuticles that the aqueous pores were preferentially located at the cuticle which covering stomata and trichomes (Schlegel et al., 2001, Schreiber, 2005, Schönherr, 2006, Schlegel et al., 2006). However, there is no experimental evidence for the presence of such pores in leaf cuticles (Fernández et al., 2009). Furthermore, Fernández et al. (2017) proposed a ‘dynamic aqueous continuum’ model which suggested that the cuticle has highly tortuous, relative humidity dependent, dynamic sorption and desorption of water connections associated with hydrophilic domains existing between the leaf surface and interior tissues. However, due to the complex chemical composition and structure of the cuticle, the mechanisms and importance of the cuticle in the foliar absorption of mineral nutrients remain unclear (Fernández et al., 2017).

It has been shown that adding a wetting agent or surfactant can significantly increase penetration of foliar applied nutrients. For example, Schönherr (2000) observed that adding a wetting agent Glucopon 215 SCUP resulted in a 10-fold increase in the rate constants of calcium chloride penetration of leaves, suggesting that this could be attributed to the improved contact between salt solutions and the cuticle.

2.3.1.2.3 An exception: water can use both pathways Interestingly, Schreiber (2005) suggested that as a small, polar but uncharged non-ionic molecule, water is an exception and it can use both pathways to cross cuticles. Firstly, it was shown that water permeability of cuticles is affected by accelerators (Riederer et al., 1990), wax extraction (Riederer et al., 2001) and increasing temperature (Schreiber, 2001). 3 Furthermore, a study on cuticular transpiration by tritiated water ( H2O) indicated that cuticular transpiration was strongly influenced by humidity (Schreiber, 2001). These studies demonstrate that water diffuses through the cuticle by the lipophilic pathway, which has low solubility but high mobility. On the other hand, water can also pass through the cuticle by the proposed aqueous polar pores (Schreiber, 2005).

2.3.2 Stomata and foliar absorption 2.3.2.1 Stomata Stomata are specialised epidermal structures that are responsible for gas exchange between the atmosphere and the plant body and are essential for plant survival and productivity. Stomata consist of two highly specialised guard cells around a small pore and are present on the leaves and stalks of nearly all land plants. Depending on the species and environmental

27 conditions, stomata range in size from about 10 to 80 μm in length and occur at densities between 5 and 1,000 mm-2 of the epidermis (Hetherington et al., 2003). For some species (e.g. citrus) stomata are only present on the abaxial leaf surface, whereas in other species (e.g. soybean, sunflower) they are present on both sides of the leaves. The stomatal development process has been well understood in the model plant Arabidopsis thaliana (Richardson et al., 2013, Chater et al., 2014), and the mechanisms that determine the opening and closure of stomata are well characterised (Daszkowska-Golec et al., 2013, Chater et al., 2014). Although the mechanisms that regulate the reaction of stomata to many exogenous and environmental cues remains unclear, it is now accepted that the mode of stomata action is subject to the integration of environmental and intracellular signals (Daszkowska-Golec et al., 2013).

Interestingly, Fernández et al. (2009) noted that cells of Arabidopsis are able to sense bacterial molecules, hence, provoking stomatal closure as a protecting mechanism, but there is no direct evidence of stomata reacting to the contact to solutions. Chater et al. (2014) stated that mature leaves can detect the environmental signals and relay messages to immature leaves to tell them how to adapt and grow, and they argued that the stomata on mature leaves were a key portal by which plants coordinate their carbon and water relations. In the past studies, the opening or closing of stomata was controlled by fusicoccin, ABA or light (Eichert et al., 1998, Eichert et al., 2008a). For example, to provoke stomata opening, leaves were illuminated or treated with fusicoccin. Stomatal closure was induced by shading or the application of ABA (Eichert et al., 2008a).

2.3.2.2 Penetration of nutrients through stomata Because of the special structure of stomata, their role, if any, in foliar absorption has been widely studied. Firstly, Schönherr et al. (1972) indicated that hydrophobicity of the leaf surface, surface tension of the liquid, and stomatal geometry control penetration of stomata by liquids, and suggested that spontaneous infiltration of stomata cannot occur unless pressure is applied or surfactants added to reduce surface tension. However, Eichert et al. (1998), using leaf segments and leaf epidermal strips of leek (Allium porrum L.), found that the penetration of uranine through stomata occurred under natural conditions, i.e., without surfactants or applied pressure. Using confocal laser scanning microscopy, Eichert et al. (2008b) observed visually that hydrophilic polystyrene particles with a diameter of 43 nm suspended in water can enter leaves through the stomata by diffusion along the walls of the pore, thereby proving

28 that penetration of the stomata is indeed possible. However, it was concluded that only a small portion of the stomata are actually penetrated, usually less than 10 %.

Previous studies on stomatal penetration can be divided into three categories according to their research approaches. Firstly, some studies have shown that stomata have a role in foliar absorption by comparing the absorption between opened and closed stomata (Sands et al., 1973, Eichert et al., 1998, Schlegel et al., 2002). Secondly, some studies have demonstrated that stomata play a role in foliar absorption by comparing the different absorption behaviours of adaxial/abaxial leaf sides given the different stomata density on the two sides (Knoche et al., 1992, Burkhardt et al., 2012, Will et al., 2012). For example, Burkhardt et al. (2012) observed greater uptake in the abaxial (stomatous) of apple leaves than the adaxial (astomatous) side, indicating that the differences are mainly caused by stomatal transport due to the structural difference of the two sides. Similar results were reported by Will et al. (2012) from a study using isotopically labelled B with lychee (Litchi chinensis Sonn.) and soybean (Glycine max cv. Merr.) plants. However, these results must be interpreted with care given the different leaf surface parameters (e.g. trichome density, cuticle composition and thickness) of the adaxial and abaxial leaf side. Thirdly, many studies have shown a positive correlation between uptake rates and stomata density which suggests that nutrient uptake through stomata is possible (Schönherr et al., 1978, Eichert et al., 2001).

The mechanism of stomatal uptake has not yet been characterised. It was suggested that the hydrophobicity and the geometry of the stomata limits the diffusion of solution through it (Burkhardt et al., 2012), but several possible pathways including depositing of hygroscopic particles and growing microbes in the pore could increase the wettability of the pores (Burkhardt et al., 2012, Eichert et al., 2012a) and potentially form a continuous liquid water film on the stomatal walls that enable diffusive solute transport (Fernández et al., 2017), with this process called “hydraulic activation of stomata” (Burkhardt, 2010, Burkhardt et al., 2009). The activation of this process is required separately for each stomata and it is potentially relevant to the composition and structure of the cuticle above the guard cells or the pore wall cells (Burkhardt et al., 2012). However, the chemical composition and structure of the pore walls and stomatal surfaces differ among species (Fernández et al., 2017). In addition, for the types of the fertilisers applied, hygroscopic leaf surface particles could facilitate the stomatal diffusion (Burkhardt et al., 2012). Therefore, at least for some species, there is now clear evidence that stomatal absorption of water and solutes is possible, but the

29 mechanism of the penetration and the overall contribution of stomata in foliar nutrient absorption are unknown.

2.3.3 Trichomes and foliar absorption 2.3.3.1 Leaf trichomes Trichomes are appendages that originate from epidermal cells and develop outwards on the surface of various plant organs (Werker, 2000). Different species have different types of trichomes on their aerial parts. Trichomes are classified as glandular or non-glandular, with the major distinction being that glandular trichomes have heads that contain or excrete various chemicals (Tian et al., 2012). Glandular trichomes are usually multicellular, consisting of differentiated basal, stalk and apical cells, and can be found on the surface of about 30 % of all vascular plants. In contrast, non-glandular trichomes are not only present on most angiosperms, but also on some gymnosperms and bryophytes (Glas et al., 2012). The morphology and chemical composition of trichomes vary with the species and the organs where they are located. They are also affected by environmental conditions. Moreover, the structure and outer surface of the trichomes are highly diverse; they may be one cell or multiple cells and may be smooth or exhibit micro-ornamentation (e.g. micropapillate, reticulate, striate) (Werker, 2000). There are numerous types of trichomes, which have not yet been described and defined.

The principal functions of trichomes include: (1) resistance against herbivories and pathogens by physically interfering with their movement, and by direct toxicity though chemicals (e.g. monoterpenes and sesquiterpenes) produced or released (Levin, 1973, Tian et al., 2012, Aschenbrenner et al., 2013), (2) protection against UV radiation, maintaining leaf surface temperature and preventing dehydration by increasing reflection, decreasing transpiration and reducing air movement on the leaf surface (Ghorashy et al., 1971, Nielsen et al., 1984, Taheri et al., 2014), (3) secreting secondary metabolites. For examples, glandular trichomes can exude various metabolites that not only repel insects but can also be used by pharmaceutical industry, in pesticides, and as food additives (Glas et al., 2012). Moreover, trichomes may be another pathway that plants communicate with other organisms and their environment, although little has been reported regarding this.

2.3.3.2 Leaf trichomes of tomato, sunflower and soybean Different types of trichomes have been identified on the tomato leaf surface. Glas et al. (2012) indicated that, typically, eight different types of trichomes occur in Solanaceae with four (i.e., Type I, IV, VI and VII) glandular trichomes and four (i.e., Type II, III, V and VIII) non-glandular, with all having distinctive morphology and chemical contents. Similarly, Tian et al. (2012) described seven different types of trichomes for tomato mutants and wild-types, with Types I, IV, VI and VII being glandular and Types II, III and V non-glandular. Kang et al. (2010) identified three types of trichomes on tomato (Solanum lycopersicum cv. Castlemart [LA2400]) and also suggested that trichome-based chemical defences play a major role in the resistance of cultivated tomatoes to opportunistic herbivores by influencing herbivore community composition under natural conditions. Schilmiller et al. (2012) mentioned that glandular secreting trichomes on the surface of tomato plants and many of its relatives in the Solanaceae family produce a mixture of O-acyl sugars that contribute to insect resistance.

Studies on sunflower leaf trichomes have consistently found three types of trichomes. For example, Spring et al. (1987) found three types of glandular and nonglandular trichomes on the leaf surface of a sunflower. Aschenbrenner et al. (2013) also reported that the sunflower leaf surface had three types of trichomes, namely non-glandular trichomes (NGT), linear glandular trichomes (LGT) and capitate glandular trichomes (CGT). Their study further indicated that, during leaf expansion, there were no substantial generation of new trichomes on the leaf surface, suggesting the trichome number per leaf was defined early and uninfluenced by external modification.

Fewer trichome types have been observed on soybean leaves. Franceschi et al. (1983) observed two types of trichomes (long nonglandular trichomes and short structure trichomes consisting of five cells in linear arrangement) on the very young expanding leaves of Glycine max cv. Wye. At least three types of trichomes were observed at the surfaces of the gynoecia of Glycine max cv. Clark (Healy et al., 2005); the linear glandular trichome found on gynoecia was similar to the glandular trichome on the leaf surface (Franceschi et al., 1983). Healy et al. (2009) also illustrated that this trichome usually consisted of five to seven linearly arranged cells, with the stalk cells with callose walls being highly vacuolate, and their cytoplasms have reduced numbers of Golgi bodies and endoplasmic reticulum. This study also indicated that during secretion, two to four distal cells develop dense cytoplasms

31 containing both prominent Golgi bodies with large vesicles and endoplasmic reticulum with enlarged lumens. Further study suggested there the trichomes were coved by cuticle, with the cuticle of the secretion region being thicker and more highly modified than a normal cuticle. Studies of the trichome densities of Glycine max cv. Clark and its isolines can be sourced from Bernard et al. (1969) and Gunasinghe et al. (1988).

2.3.3.3 Movement of nutrients through trichomes In contrast to the cuticle and stomata, the penetration property of trichomes has not been studied in much depth. It is known that glandular trichomes can secrete varied types of specialised metabolites (Schilmiller et al., 2008). Furthermore, some studies have found that some types of trichomes can accumulate excess heavy metals and are even able to excrete them out of the plant (Blamey et al., 1986, Sarret et al., 2006), suggesting that some channels exist on trichomes. In addition, absorption of water from dew (Pina et al., 2016) or fog (Ju et al., 2012) through trichomes has been reported in some species as a drought resistance mechanism, indicating that absorption of foliar-applied nutrients may potentially occur through trichomes in some plants. However, it remains unclear whether trichomes play a role in the absorption of foliar applied nutrients (Fernández et al., 2013a). Given the diverse varieties of trichomes with different structures and compositions, as stated above, it is possible that the penetration property of trichomes may vary among different types of trichomes.

2.4 Translocation of foliar applied nutrients

The foliar absorption of nutrients is a complex process. As Fernández et al. (2013b) notes, it mainly includes the following procedures: (1) foliar adsorption, (2) cuticular penetration, (3) diffusion in apoplastic and symplastic spaces, and (4) phloem loading into vascular veins and translocation out of the sprayed leaf tissues into other actively growing parts of the plants. After penetration through the cuticle, the nutrients need to cross the epidermal cells (or directly enter the mesophyll from the stomata pore) and then load into the vascular bundle and transfer to other part of the plants. A high translocation rate would not only protect the leaf from damage caused by the high concentration of the applied nutrients, but also influence the efficacy of the foliar applied nutrients (Fernández et al., 2013a).

Very few studies have focused on the mobility of foliar applied nutrients within the leaf tissues. For example, how the nutrients load to the phloem i.e., through the apoplastic and/or symplastic pathway is unclear. Given the role of apoplast in maintaining ion balance, element detoxification, transmission of signals leading to stomatal response, and regulation of pH fluctuations, Fernández et al. (2013a) suggested that apoplast might play a critical role in determining the fate of foliar applied nutrients. The apoplast constitutes all compartments beyond the plasmalemma – the interfibrillar and intermicellar space of the cell walls, and the xylem, including its gas- and water-filled intercellular space; the border of the apoplast is the outer surfaces of plants, i.e. the rhizoplane and the cuticle acher (Sattelmacher, 2001). The apoplast acts as a short-term reservoir for ions that regulates ion exchange and may accumulate cations and repel anions (Sattelmacher, 2001, Fernández et al., 2013a). For instance, the limited mobility of cations in plant leaves, such as Zn or Fe, might be due to their binding to the negative charges in the apoplastic space (Fernández et al., 2013a). However, the movement of the foliar applied nutrients in the leaf tissues is unclear.

It is already known that the mobility of nutrients in plants varies among nutrients. In terms of mobility in the phloem, essential nutrients have been classified as highly mobile (N, P, K, Mg, S, Cl, Ni), intermediate or conditionally mobile (Fe, Zn, Cu, B, Mo), and rather immobile (Ca, Mn) (Fernández et al., 2013a). For example, Marešová et al. (2012) found that < 1% foliar applied Zn is transported out of the applied leaf, whereas foliar applied P can substantially transfer to other parts of the plants (Koontz et al., 1957). The redistribution of nutrients is relevant to plant growth stages, for example, Pearson et al. (1994) found that Zn was remobilized from the flag leaf to grains of wheat especially during grain development stage. It is proposed that Zn could be transported through the phloem as Zn2+, Zn-nicotianamine (Nishiyama et al., 2012), Zn-malate, Zn-histidine complexes (Álvarez-Fernández et al., 2014, Gupta et al., 2016) or Zn-DMA (2’-Deoxymugineic acid) (Bashir et al., 2012). However, the mechanisms of the translocation of the foliar absorbed Zn within plants remain unclear.

2.5 Effects of nutrient status on foliar nutrient absorption

It was noted that generally foliar absorption was reduced in nutrient deficient plants. For example, Will et al. (2011) report that B foliar absorption in B deficient soybean was significantly lower than that in B sufficient plants, and the translocation of the B out of the applied leaf was also decreased. Similarly, Fernández et al. (2014a) found that absorption of

33 foliar-applied P was lower in P deficient wheat than P sufficient wheat. It was proposed that this was due to the changes in leaf properties, especially cuticle relevant properties and hence the permeability of the leaf surface has changed (Fernández et al., 2013b). For example, it was found the amount of wax per unit surface was decreased in Fe-deficient leaves of peach (Prunus persica) (Fernández et al., 2008) and P-deficient leaves of wheat (Fernández et al. 2014a). Eichert et al. (2010) found that xylem vessel size reduced for Fe deficient plants. Will et al. (2011) also found that the stomata of the B deficient leaves were shrunken and closed, and it was believed that this reduced foliar B absorption through the stomata pathway. Moreover, some find N deficiency induced an increase in epicuticular wax concentration (Prior et al., 1997, Bondada et al., 2006) but it was found N foliar absorption in N deficient plants was greater than that in N sufficient plants (Klein et al., 1984, Fernández-Escobar et al., 2011). In contrast, Erenoglu et al. (2002) reported that foliar absorption of Zn in wheat was not influenced by the Zn nutritional status, with Zn deficient leaves translocating more Zn out of the leaf than the Zn sufficient plants. How the Zn status of plant itself influence the absorption rate of the foliar applied Zn, and what has caused this effect are unknown.

2.6 Use of nanomaterials as fertilisers

Nanomaterials are engineered structures with at least one dimension of 1 to 100 nm. Nanomaterials lie in the transitional zone between individual atoms or molecules and bulk materials of the same chemical composition. They are increasingly being used for commercial purposes such as fillers, opacifiers, catalysts, semiconductors, cosmetics, microelectronics, and drug carriers (Ratner et al., 2003). The unusual physicochemical properties of engineered nanomaterials are attributable to their small size (surface area and size distribution), chemical composition (purity, crystallinity, electronic properties, etc.), surface structure (surface reactivity, surface groups, inorganic or organic coatings, etc.), solubility, shape, and aggregation (Nel et al., 2006). Currently, the application of nanotechnology in agriculture, e.g. nano-fertiliser has great potential (DeRosa et al., 2010, Wang et al., 2016).

As DeRosa et al. (2010) defined, a nanofertiliser refers to a product that delivers nutrients to crops in one of three ways: (1) nutrient is encapsulated inside nanomaterials such as nanotubes or nanoporous materials, (2) nutrient is coated with a thin protective polymer film, or (3) nutrient is delivered as particles or emulsions of nanoscale dimensions. Due to the large ratio of surface area to volume, and the advantage that nanofertiliser is easily surface

34 functionalised to meet multiple purpose e.g. Zn release rates, surface adhesion, and optimised Zn uptake, the effectiveness of nanofertilisers may surpass that of traditional foliar fertilisers (Naderi et al., 2013). Moreover, with the current emphasis to optimise the efficiency of fertilisers through the 4R Nutrient Stewardship principles, i.e. the use of the fertiliser from the right source, at the right rate and right time, with the right placement (Roberts, 2008), nanofertilisers may provide a strategy in this regard.

Indeed, some studies have demonstrated that nanoparticles can penetrate into plants. Lin et al. (2008) reported that ZnO nanoparticles were able to concentrate in the rhizosphere, enter the root cells. Du (2014) reported a short-term foliar Zn uptake from a newly synthesised Zn hydroxide nitrate [Zn5(OH)8(NO3)2·2H2O] nanocrystal suspension on tomato leaf surfaces. These studies imply that it is possible to develop nanofertilizer as a new nutrient delivery system for plants.

Generally, foliar fertilisers have lower environmental impact than most soil-applied fertilisers, primarily because of the lower rates applied, thereby minimising the leaching risk to soil or water (Fernández et al., 2013b). However, an uncertain risk that may be raised is associated with the use of nanofertilisers, given that it is a new artificial material and their interaction with natural systems remains unclear (Wang et al., 2013). For example, spray drift may occur and non-target organisms such as insects or soil microbes may be affected (Bindraban et al., 2014). Despite the advantages of nanofertilisers, however, their environmental impacts need further assessment.

2.7 Research questions

Zinc deficiency is a worldwide problem, which causes a decrease in crop yield, nutrient quality impairment, and subsequently serious human health problems. Due to the fact that the soil properties can potentially restrict uptake of Zn fertilisers when applied to the soil, foliar application of Zn has been demonstrated to often be the most efficient way to overcome Zn deficiency. In addition, nanofertilisers have been proposed as a promising strategy in biofortification of Zn. However, the mechanism of nutrient foliar absorption and the translocation of foliar absorbed nutrients remain unclear. The following knowledge gaps have been identified:

1) The role of cuticle, trichomes, and stomata in foliar Zn absorption is unknown, with the mechanisms responsible for cuticle hydrophilic penetration and stomatal uptake unclear;

2) It remains unclear whether ZnSO4 (traditional foliar fertiliser) and Zn foliar nanofertiliser have different absorption and translocation modes, as well as efficacy for crops. 3) The translocation of the foliar applied Zn in the plant is not clear, including the short term transfer in the applied leaf tissues and the long term transfer in the whole plant; 4) Whether the Zn status of the plant itself influences the absorption of foliar applied Zn and the underlying mechanisms are unknown.

This project was designed to test the following hypotheses: 1) Zinc foliar absorption through cuticle, stomata and trichome are all possible, but their

importance varies for different species and the applied forms of Zn (ZnSO4 and nano- ZnO);

2) The translocation of Zn foliar fertiliser differs for different applied Zn forms (ZnSO4

and nano-ZnO), with soluble forms of Zn (such as ZnSO4) being having higher translocation than forms of lower solubility (such as nano-ZnO); 3) Zinc status of the plant itself affects foliar Zn absorption and translocation, with the absorption and translocation of Zn being lower in Zn-deficient plants than in Zn- sufficient plants.

Testing these hypotheses will deliver new knowledge in the area of plant nutrition in relation to the absorption of nutrients through foliar fertilisation. This has great significance in improving understanding of absorption and translocation of Zn foliar fertilisers. Moreover, effects of the Zn status of plant itself on foliar absorption of Zn and the absorption of nano Zn foliar fertiliser were tested, hence contributing to understanding of the physiological foundations for applying foliar Zn fertilisers and development of foliar nanofertilisers.

Chapter 3 Effects of changes in leaf properties mediated by methyl jasmonate (MeJA) on foliar absorption of Zn, Mn, and Fe

3.1 Introduction

The pathways whereby foliar applied nutrients move across the leaf surface remain unclear. Based on the morphology of the leaf surface, it has been proposed that the cuticle, stomata, and trichomes are all possible pathways for the absorption of foliar applied nutrients (Fernández et al., 2013b). The first experiment therefore aimed to use an approach to modify the leaf properties in order to determine how these changes altered the foliar absorption of nutrients. In this regard, it is known that MeJA can cause an increase in leaf trichome density (Tian et al., 2012), thus in this experiment MeJA was used to induce changes in leaf properties such as trichome density. Using sunflower, tomato, and soybean, the leaf properties that might be related to foliar nutrient absorption including trichome density, stomatal density, leaf cuticle thickness, and cuticle composition were examined after treating plants with MeJA at five concentrations (0, 0.1, 0.5, 1.0, and 2.5 mM).

Based on the results of the first experiment, a MeJA concentration was selected (1 mM MeJA) which had been shown to alter leaf properties but not impact deleteriously on plant growth (plant mass and height). Using both traditional analytical approaches as well as synchrotron-based µ-XRF, these changes in leaf properties were then related to the foliar absorption of Zn, Mn, and Fe. The overall aim of these experiments was to investigate how leaf trichomes, stomata, and the cuticle influence foliar nutrient absorption.

3.2 Paper 1

Li C, Wang P, Menzies NW, Lombi E, Kopittke PM (2018) Effects of methyl jasmonate on plant growth and leaf properties. Journal of Plant Nutrition and Soil Science 181: 409-418. (https://doi.org/10.1002/jpln.201700373)

Effects of methyl jasmonate on plant growth and leaf properties Running title: methyl jasmonate and plant growth CUI LI1, PENG WANG2, 3*, NEAL W. MENZIES1, ENZO LOMBI4, PETER M. KOPITTKE1

1 The University of Queensland, School of Agriculture and Food Sciences, St Lucia, Queensland, 4072, Australia. 2 Nanjing Agricultural University, College of Resources and Environmental Sciences, Nanjing, 210095, China. 3 The University of Queensland, Centre for Soil and Environmental Research, School of Agriculture and Food Sciences, St Lucia, Queensland, 4072, Australia. 4 University of South Australia, Future Industries Institute, Mawson Lakes, South Australia, 5095, Australia

*To whom correspondence should be addressed. Phone: +86 25 84399055, Fax: 86 25 84399055, Email: [email protected]

Number of text pages: 15 Number of Figures: 9 (Figure 7 and Figure 9 in colour) Number of Tables: 0

Key Words —— Methyl jasmonate; Plant growth; Trichome density; Stomatal density; Cuticle thickness.

Abstract —— Exogenous application of methyl jasmonate (MeJA) can induce anatomical and chemical changes that are components of defence responses in plants. Particularly, MeJA is well-known to increase leaf trichome density to protect against insect herbivory, but surprisingly little is known about the effects of MeJA on other leaf properties and plant growth. Using sunflower (Helianthus annuus), tomato (Solanum lycopersicum), and soybean (Glycine max) treated with 0, 0.1, 0.5, 1, and 2.5 mM MeJA, we examined changes in leaf trichome density, stomatal density, cuticle thickness, cuticle composition, plant height, and biomass production. For all three plant species, MeJA (especially at the higher concentrations) caused significant decreases in plant height (up to 39 %) and biomass (up to 79 %). MeJA caused substantial increases in leaf trichome density (being 1.3- to 3.5-times higher) in all three species, with the magnitude of these effects increasing with MeJA concentration. However, we also observed that MeJA resulted in significant changes in cuticle composition and thickness, and stomatal density, although the magnitude of these changes was smaller relative to changes in trichome density. Specifically, high concentrations of MeJA increased the relative content of phenolic compounds and cutin in leaf cuticle while decreasing the relative content of polysaccharide. The changes in stomatal density varied with plant species and MeJA concentration. Also, MeJA increased cuticle thickness in tomato but decreased that in sunflower and soybean. Thus, studies investigating MeJA should also consider the importance of changes in other leaf properties and plant growth.

Introduction For agricultural crops, the ability to cope with biotic and abiotic stress is crucial for ensuring a healthy plant, high yield, and a high-quality product. Over the coming decades, agricultural production will be required to increase substantially whilst also increasing in efficiency – this being due to an increasing human population, climate change, and water and soil scarcity (Kazan 2015). Unlike animals, plants rely heavily on hormones to mediate responses to all kinds of stimuli, including jasmonates (JAs) (Dar et al., 2015). Jasmonates is a group of oxylipins derived from linolenic acid and it is universally exist in plant kingdom (Cheong and Choi 2003). Both the metabolism and mechanism of action of JAs have been widely studied (Creelman and Mullet 1997; Turner et al., 2002; Wasternack 2007), with JAs (as with other phytohormones) involved in multiple functions, such as seed germination (Yildiz et al., 2007), pollen germination (Yildiz and Yilmaz 2002), senescence (Kim et al., 2015), seedling growth (Kilic et al., 2008), fruit ripening (Mukkun and Singh 2009), and cellular and inter-plant communication (Cheong and Choi 2003). However, the most well-known function of JAs is in the regulation of defence-related genes (Howe 2010; Wasternack 2017), with JAs essential for triggering the biologically effective defence in plant–insect interaction (McConn et al., 1997; Zhang et al., 2017). For example, it was found that concentrations of JAs increased substantially in wounded leaves of Arabidopsis thaliana within 30 s (Glauser et al., 2009).

Methyl jasmonate (MeJA) is a naturally occurring fragrant volatile ester form of jasmonic acid, representing one of the best known members of JAs. It is known that MeJA plays an important role in defense mechanisms to protect against both biotic and abiotic stresses (Cheong and Choi 2003). Generally, JAs are used to regulate responses through: (i) alteration of plant growth and development through long-distance signalling (Huber et al., 2005; Motallebi et al., 2015), (ii) regulation of plant secondary metabolism (Meldau et al., 2012), for example, by inducing the production of toxic compounds to protect against herbivory (Moreira et al., 2012) or by increasing reactive oxygen species (ROS) activity and content to protect against abiotic stresses (Cao et al., 2009; Parra-Lobato et al., 2009; Anjum et al., 2011), and (iii) enhancing the rate of healing from damage (Peña-Cortés et al., 1995), for example, Balusamy et al., (2015) found that exogenous MeJA was able to promote the recovery of Panax ginseng leaves recovery from wounding.

Given that many of the functions of MeJA have been identified, there are increasing numbers of studies investigating the effects and potential uses of MeJA in agriculture (Reyes-Díaz et al., 2016). For example, it has been suggested that MeJA can be used to enhance grape (Vitis vinifera) and wine quality (Portu et al., 2015), improve the anthocyanin and flavonol content in blackberries (Rubus sp.) (Wang et al., 2008), raspberries (Rubus occidentalis L) (Wang and Zheng 2005), and apples (Malus pumila) (Shafiq et al., 2013). Given the important role of MeJA in the plant defence system, it has also been suggested that MeJA could be used as a chemical elicitor in young conifer (Pinophyta) trees to increase their self-resistance ability (i.e. the ability of the plants themselves to resist both biotic and abiotic stresses), thereby increasing the survival rate of seedlings (Moreira et al., 2012). Similarly, it has been reported that pre-treatment of plants with MeJA can alleviate the deleterious effects of salinity (Fedina and Tsonev 1997; Del Amor and Cuadra-Crespo 2011; Wargent et al., 2013). Interestingly, if MeJA can be used as a natural regulator in agriculture to reduce insect herbivory and disease incidence and to enhance the plant resistance to abiotic stress, it would benefit not only food production but also provide an environmentally friendly alternative to pesticides. However, it has also been reported that the exogenous application of MeJA can decrease photosynthesis (Attaran et al., 2014) and inhibit plant growth (Shyu and Brutnell 2015). Surprisingly little is known about the effects of exogenous MeJA on leaf properties. This lack of knowledge not only hinders research into the use of MeJA itself, but it is also important for understanding the effects of MeJA as related to a range of scientific fields. For instance, although MeJA is often used experimentally to investigate trichomes and herbivory (Tian et al., 2012), consideration is not normally given to the potential effects of MeJA on other leaf properties, such as stomatal density and cuticle thickness. Indeed, we are not aware of any studies that have provided detailed information regarding the effects of MeJA on a range of leaf properties, such as leaf cuticle composition, cuticle thickness, and trichome and stomatal densities – these being the focus of the present study.

The aim of the present study was to investigate the effect of MeJA on plant growth, giving particular consideration to changes in biomass and leaf properties such as stomatal density, trichome density, cuticle composition, and cuticle thickness. We investigated, three plant species, being sunflower (Helianthus annuus), tomato (Solanum lycopersicum), and soybean (Glycine max) and used five MeJA concentrations (0, 0.1, 0.5, 1, and 2.5 mM). To investigate leaf properties, we utilized scanning electron microscopy (SEM), light microscopy, and

Fourier transform infrared spectroscopy (FTIR). The results of this experiment will assist in understanding the effects of MeJA on plants.

Materials and methods Experimental design and plant growth The plant-growth experiment was conducted at The University of Queensland (St Lucia, Australia), in a growth room at 25 °C with high pressure sodium lamps providing light (photon flux density of 500 μmol m–2 s–1) for 14 h/d. Seeds of tomato (cv. Red Luck), soybean (cv. Bunya) and sunflower (cv. Hyoleic 41) were germinated in rolled paper towel moistened with tap water for either 3 or 4 d before being transferred to 11 L buckets filled with a basal nutrient solution (below). Each container had four holes with a total of either eight (for tomato and soybean) or four (for sunflower) plants – each container forming one experimental unit. The nutrient solution was similar to that used by Blamey et al., (2015), but - + the composition was modified to contain (µM): 910 N (94 % NO3 and 6 % NH4 ), 475 K, 20 P, 1126 Ca, 227 Mg, 1251 Cl, 556 S, 25 Fe(III)EDTA, 3 B, 0.5 Mn, 0.5 Zn, 0.2 Cu, and 0.01 Mo. Nutrient solutions were changed after one week, and then every 4 d thereafter. To progressively replace P which had been taken up by the plants, once the plants had been growing in the basal solution for 10 d, 5 mL of 44 mM KH2PO4 was added every second day thereafter.

The experiment consisted of three plant species and five concentrations of MeJA 0 (control), 0.1, 0.5, 1, and 2.5 mM, with these being in the general range used in previous studies [for example, ranging between 0.001 and 15 mM (Reyes-Díaz et al., 2016; Tian et al., 2012)]. We conducted a preliminary experiment (0, 0.1, 1, 2.5, 5, 7.5, 10, and 15 mM MeJA) which identified that concentrations > 2.5 mM were potentially lethal, and hence these concentrations were not included in the final experiment. Each of the 15 treatments was replicated four times (i.e. there were four containers for each treatment, with each container having four or eight plants), yielding a total of 60 experimental units. Application of the MeJA occurred after the plants had been growing in basal solution for 7 (sunflower), 8 (soybean) or 10 d (tomato). The MeJA (in 0.8 % ethanol) was sprayed onto the foliage until the leaves were fully saturated (ca. 300 mL per container) (Boughton et al., 2005; Tian et al., 2012). After application, the plants were allowed to grow for 4 h without lighting. Thereafter, plants were grown for a further 10 d under the same condition as described previously before

42 measurement of root length, plant height, and biomass. For the assessment of biomass, each plant was separated into roots, stems, and leaves before oven-dried at 65 °C for 2 d. The youngest fully-expanded leaves (YFELs) were used for examining leaf morphology (leaf area, stomatal density, and trichome density), cuticle thickness, and cuticle composition as described below.

Examination of leaf characteristics Using the YFELs, leaf morphology was examined using SEM, with four replicates of each treatment. Firstly, photos of the YFELs were taken using a digital camera (Canon EOS 5D Mark III) and the area of the YFELs measured by ImageJ (available free-of-charge at https://imagej.net). For leaf trichome and stomatal densities, fresh material (ca. 3 mm × 3 mm) was collected from the base, middle, and tip of leaves from each treatment and immediately fixed in 3 % glutaraldehyde in 0.1 M sodium cacodylate. Microwave processing was performed using a Pelco BioWave (Ted Pella Inc., California, USA) with a ColdSpot water recirculating device. Following fixation with glutaraldehyde, samples were post-fixed with 1 % osmium tetroxide, subjected to a dehydration series using ethanol (30, 50, 70, 90, 100, and 100 %) before being processed using the BioWave. The tissues were then subjected to critical point dryer (Autosamdri-815 CPD, USA) before being coated with Au. Finally, samples were examined using a SEM (JEOL NEOSCOPE, Japan) at 10 kV accelerating voltage to determine the density of trichomes and stomata on the adaxial leaf surface. For measuring trichome density, images were obtained using SEM at a magnification of 100×, with the number of trichomes manually counted across a total area of ca. 5 mm2, with this repeated for each of the four replicates. Similarly, for stomatal density, images were obtained using SEM at a magnification of 500×, with the number of stomata manually counted across a total area of ca. 0.2 mm2.

Cuticle composition was examined in a manner similar to that described by Chatterjee et al., (2012). Briefly, using the middle part of each YFEL, the abaxial side was scraped using a scalpel, with the abaxial cuticles and leaf tissues scraped as clean as possible, and the adaxial cuticles placed into an enzyme solution containing 2 % pectinase and 0.1 % cellulase in 50 mM sodium acetate buffer. These tissues were digested overnight at room temperature before being washed twice using deionised water. Samples were examined using light microscopy to ensure that the underlying tissues were completely removed before being air dried and then

43 the four replicates of each treatment were mixed evenly and analysed using FTIR (Guzmán et al., 2014).

The combined thickness of the cuticle and epidermal cell wall was measured using sections cut from resin-embedded samples examined with a light microscope. Briefly, the middle portion of the fresh YFELs was cut into pieces ca. 1 × 4 mm. The segments were fixed in 3 % glutaraldehyde in 0.1 M sodium cacodylate at 4 °C overnight, rinsed twice with 0.1 M sodium cacodylate before being post-fixed with 1 % OsO4 in 0.1 M sodium cacodylate for 4 h, and dehydrated in a graded ethanol series of 20, 30, 40, 50 (10 mins each) 70, 90, 100 (30 mins each), and 100 % (overnight). Epon resin was used to infiltrate the tissues with a graded resin: 10, 20, 30, 40 (90 mins), 50 (overnight), 65, 80 (4 h), and 100 % (overnight) mixtures of Epon in ethanol. The tissues were then embedded in 100 % Epon blocks and polymerized at 60 °C overnight. Sections (1 µm) were cut using an ultramicrotome (LEICA EM UC6), stained with Toluidine blue (0.5 % in 1 % borax) and observed under the light microscope (O'brien 1995), with four replicates examined.

Statistical analyses Data were analyzed using a two-way analysis of variance (GenStat 18th Edition). Comparisons between means were made using Fisher’s protected least significant difference (LSD) test. Where a significant interaction was observed between factors, the LSD value for that interaction is presented, and means can be compared between both factors. Where no significant interaction was found, LSD values are presented for each individual factor as appropriate.

Results Effects of methyl jasmonate on plant growth Except for root length of sunflower, application of MeJA reduced plant growth, measured as plant height, root length, and biomass for all three plant species, with the application of elevated concentrations of MeJA resulting in marked decreases in these parameters (Figure 1a-d). For example, for soybean, 1 mM MeJA decreased root dry mass by 15 %, shoot dry mass by 22 %, root length by 26 %, and plant height by 17 % (Figure 1a-d). However, the magnitude of the decrease in plant height, root length, and biomass depended upon the concentration of MeJA. For example, in soybean and tomato, 0.1 mM MeJA did not cause a

44 significant decrease in these parameters. In sunflower, the same concentration resulted in significant decreases in plant height and biomass (Figure 1a-d), indicating that the plants differed in the nature of their response to MeJA. We also assessed leaf area for the YFELs, with the application of MeJA having no effect on leaf area for any of the three plant species, regardless of the concentration of the MeJA (Figure 1e).

Effects of methyl jasmonate on leaf trichome and stomatal density For the control treatments, the average trichome density was found to be 1,300 cm-2 for soybean, 1,600 cm-2 for sunflower, and 1,800 cm-2 for tomato. For all the three species, trichome density was found to vary with the portions of the leaves, with the tip having the highest density (Figure 2-4). Trichomes are identified as either glandular or non-glandular [see Franceschi and Giaquinta (1983), Glas et al., (2012), Tian et al., (2012), and Aschenbrenner et al., (2013) for details of the glandular and non-glandular trichomes in soybean, tomato, and sunflower], with the major distinction being that glandular trichomes can contain or excrete various chemicals (Tian et al., 2012). As expected, application of MeJA was found to result in significant increases in the density of trichomes, with the average trichome density of the MeJA treatments being 2.3-times higher than the control for soybean, 1.6-times higher for sunflower and 1.6-times higher for tomato (Figure 2-5). Generally, the magnitude of the increase in trichome density increased with MeJA concentration. For example, for soybean, the average trichome density at 0.1 mM was 1.5-times higher than the control whilst it was 3.5-times higher at 2.5 mM. Whilst the densities of both glandular and non-glandular trichomes tended to increase for both soybean and sunflower, in tomato, only the density of glandular trichomes tended to increase with increasing MeJA, with densities of non-glandular trichomes increasing significantly only at 2.5 mM MeJA (Figure 5). For the glandular trichomes on the tip of the soybean leaf (Figure 5b), it was also noted that the density increased significantly at 0.5 mM MeJA, before decreasing again at 1 and 2.5 mM MeJA. The cause of this increase observed at 0.5 mM MeJA is not known. Regardless, the leaf area of the YFELs was not affected by MeJA (Figure 1e), and hence the changes in trichome density were not due to concomitant changes in leaf area.

The average stomatal density in the control treatments was 20,100 cm-2 for soybean, 22,900 cm-2 for sunflower, and 27,400 cm-2 for tomato (Figure 6). However, these stomatal densities depended upon the plant species, leaf position, and MeJA concentration. Overall, MeJA

45 generally increased stomatal density for soybean (average increase of 26 %) and sunflower (average increase of 16 %), while decreased stomatal density for tomato (average decrease of 4.5 %), although it was noted that the relative magnitude of these changes in stomatal density (Figure 6) was substantially less than observed for the trichomes (Figure 5).

Effects of methyl jasmonate on cuticle thickness Although the cuticle is usually considered to be separate from the underlying polysaccharide epidermal cell walls, recent studies have suggested that the cuticle is simply an extension of the cell wall region that contains additional lipids (Guzmán et al., 2014; Yeats and Rose 2013). Therefore, in the present study, we referred to cuticle thickness as the combined thickness of the epidermal cell wall and cuticle (Figure 7). In untreated plants, the cuticle was thicker in sunflower (3.0 µm) than in soybean (2.1 µm) and tomato (1.9 µm) (Figure 8a). A significant interaction was found between MeJA and the plant species, indicating that although the MeJA concentration influenced cuticle thickness, the pattern of this change depended upon the plant species. For example, in tomato it was found that cuticle thickness tended to increase significantly with increasing MeJA concentration (e.g. from 1.9 μm at 0 mM MeJA to 2.7 μm at 2.5 mM MeJA), whilst in sunflower it was observed that cuticle thickness tended to decrease significantly with increasing MeJA concentration (e.g. from 3.0 μm at 0 mM MeJA to 1.7 μm at 2.5 mM MeJA). For soybean, only 2.5 mM MeJA significantly reduced leaf cuticle thickness, being from 2.07 μm at 0 mM MeJA to 1.53 μm at 2.5 mM MeJA (Figure 8a).

Not only did cuticle thickness vary among species, but the thickness of the overall leaf also varied (Figure 8b). Sunflower was found to have the thickest leaf, although thickness increased only slightly with increasing MeJA (from 250 μm at 0 mM MeJA to 310 μm at 2.5 mM MeJA, Figure 8b). In contrast, the thickness of tomato leaves increased markedly (from 170 μm at 0 mM MeJA to 290 μm at 2.5 mM MeJA) whilst the thickness of soybean leaves tended to decrease slightly (from 160 μm at 0 mM MeJA to 110 μm at 2.5 mM MeJA, Figure 8b).

Effects of methyl jasmonate on cuticle composition Analysis of isolated cuticles using FTIR showed that the three plant species had the same function groups (Figure 9). The broad band at 3400 cm-1 is assigned to the stretching vibration

46 of hydroxyl groups that interact by H bonding, ν(O-H…O), with cutin and polysaccharides being the components. The two bands at around 2900 cm-1 are assigned to the asymmetrical and symmetrical stretching vibrations of CH2 groups, νa (CH2) and νs(CH2), with the major contribution being from cutin and waxes. A weak band at approximately 1730 cm-1 is associated with cutin, whilst the band at 1600 cm-1 is related to aromatic and C=C functional groups from phenolic compounds or cutan. The highest band at 1020 cm-1 is associated with polysaccharides. For all the plant species, application of MeJA had no marked effects on cuticle compositions. The only exception was at the highest MeJA concentration (2.5 mM), with the relative content of phenolic compounds, cutin or cutan (corresponding to the bond at 1600 cm-1) increasing while the relative content of polysaccharides decreased (corresponding to the bond at approximately 1020 cm-1) (Guzmán et al., 2014; Heredia-Guerrero et al., 2014) (Figure 9).

Discussion Jasmonates are an important group of phytohormones, despite their importance and their widespread use of JAs in scientific studies, the effects of MeJA application on a range of leaf properties, such as cuticle thickness and composition, stomatal density remain unknown. In the present study, we found that increasing concentrations of MeJA reduced plant height and plant biomass (Figure 1), while increased leaf trichome densities (Figure 2-5). Depending upon plant species and MeJA concentration, it was also found that MeJA induced changes in cuticle thickness (Figure 8a), leaf thickness (Figure 8b), cuticle composition (Figure 9), and leaf stomatal density (Figure 6). This data is important not only for understanding the effects of MeJA per se, but also provides important information for studies utilizing MeJA.

Effects of methyl jasmonate on plant growth Methyl jasmonate is an important phytohormone involved in diverse developmental process (Parra-Lobato et al., 2009). Of particular interest, MeJA plays an important role in resistance to a range of biotic and abiotic stresses (Cheong and Choi 2003). For example, it is known that exogenous application of MeJA can induce chemical and anatomical changes leading to a reduction in insect herbivory in plants (Parra-Lobato et al., 2009; Anjum et al., 2011; Moreira et al., 2012). In the present study, we observed that the application of MeJA reduced plant height and biomass in soybean, sunflower and tomato, with the rate of decrease increasing with MeJA concentration (Figure 1). Even the lowest concentration used, 0.1 mM,

47 resulted in significant decreases in shoot mass and root mass for sunflower, although significant differences were not observed for soybean and tomato at this concentration (Figure 1). The differences observed among the three plant species are likely due to differences in the sensitivity of each species to MeJA. The observation that elevated levels of MeJA reduce plant growth is in accordance with previous studies (Heijari et al., 2005; Gould et al., 2009; Sampedro et al., 2011; Moreira et al., 2012).

Nevertheless, the mechanisms by which MeJA inhibits plant growth are not well understood. It has been suggested that jasmonate can inhibit the cell division cycle (Świa̧ tek et al., 2002; Świątek et al., 2004). In addition, Noir et al., (2013) found that MeJA decreased both cell number and size in Arabidopsis through the jasmonate receptor COI1. More recently, Kim et al., (2015) proposed that since MeJA was also important in the control of plant senescence, the exogenous application of MeJA could reduce growth by triggering the plant senescence system. However, given that MeJA generally causes a decrease in plant biomass, using of MeJA to improve plant defence ability should also consider this adverse effect, and should be carefully assessed (for example, MeJA concentrations and plant species should be considered).

Effects of methyl jasmonate on leaf trichome and stomatal density Trichomes are appendages that originate from epidermal cells and develop outwards on the surface of plant organs. The primary functions of trichomes include: (i) resistance against herbivories and pathogens by physically interfering with their movement, or by direct toxicity though the production or release of various chemicals (e.g. monoterpenes and sesquiterpenes) (Levin 1973; Tian et al., 2012; Aschenbrenner et al., 2013), (ii) protection against UV radiation, maintaining leaf surface temperature and preventing dehydration by increasing reflection, decreasing transpiration and reducing air movement on the leaf surface (Ghorashy et al., 1971; Nielsen et al., 1984; Taheri et al., 2015), and (iii) secretion of secondary metabolites, such as glandular trichomes which can exude various metabolites that not only repel insects but can also be used by pharmaceutical industry, in pesticides, and as food additives (Glas et al., 2012).

In the present study, the exogenous application of 0.1 to 2.5 mM MeJA to plants of soybean, tomato and sunflower significantly increased trichome densities in YFELs (Figure 5), with the

48 magnitude of the difference increasing with MeJA concentration. For soybean and sunflower, it was observed that both glandular and non-glandular trichome densities increased (Figure 5a-d). For tomato, the density of the glandular trichomes increased as the MeJA concentration increased from 0 to 2.5 mM, while the density of the non-glandular trichomes only increased at a MeJA concentration of 2.5 mM (Figure 5e-f). These findings are generally in agreement with previous studies. For example, Boughton et al., (2005) observed that application of 7.5 mM MeJA to tomato (c.v. Trust) increased density of glandular trichomes on YFELs. Similarly, Tian et al., (2012) reported that the application of 3.3 mM MeJA to tomato mutants increased the trichome density of YFELs by 37.8 to 85.8 %. These increases in trichome density, observed both in the present and previous studies, are due to the role of jasmonate signalling in the plant defence system. These systemic defence mechanisms can be triggered in response to herbivores or other damage, with higher densities of trichomes often improving defence against insects (Traw and Bergelson 2003; Tian et al., 2012).

In contrast to trichomes, MeJA had only a comparatively small (and variable) effect on stomatal density for the three plant species (Figure 6). For example, in sunflower, stomatal density increased only an average of 17.3 % at 1 mM MeJA, whilst in tomato, stomatal density actually decreased an average of 16.9 % at 1 mM MeJA (Figure 6). It has reported that MeJA can induce stomatal closure (Hossain et al., 2011), but we are unaware of any previous studies that have examined the effects of MeJA on stomatal density. Thus, the effect of MeJA on stomatal density was comparatively small, and depended upon the plant species and the applied MeJA concentration.

Effects of methyl jasmonate on leaf cuticle The cuticle is an extracellular polymer membrane, which covers aerial organs such as stems, leaves, flowers and fruits, serving as an important protective barrier. For example, it is the first line of defence against pests and pathogens (Yeats and Rose 2013). Interestingly, we found that the application of MeJA decreased the thickness of the epidermal cell wall and cuticle in soybean and sunflower but increased for tomato under the application of MeJA (Figure 8a). Again, although significant changes were observed, it was noted that the magnitude of these changes in cuticle thickness was less than observed for trichome density. Indeed, cuticle thickness changed an average of 13 % in soybean, 35 % in sunflower, and 37 % in tomato (Figure 8a). Leaf thickness is an important character in plants particularly

49 when assessing plants’ productivity or ecological performance (Vile et al., 2005). For example, leaf thickness is an important component of specific leaf area (SLA), leaf dry matter content, and leaf physical strength (Pérez-Harguindeguy et al., 2013). In the present study, we found that increasing concentration of MeJA increased the leaf thickness of tomato and sunflower while decreased the leaf thickness of soybean, with the amplitude of variation similar to the changes in cuticle thickness (Figure 8b). Further studies need to be conducted regarding this aspect to understand the mechanisms of the effects of MeJA on cuticle and leaf thickness.

Cuticle composition was analyzed using FTIR, with all three plant species having functional groups (Figure 9). (c.f. the diversity found among some species, see Heredia-Guerrero et al., (2014) for example). Furthermore, the application of up to 1 mM MeJA was found to have little influence on cuticle composition, with the spectra for all concentrations being similar to the control (Figure 9). However, for all three plant species, the application of 2.5 mM MeJA increased the relative content of phenolic compounds (cutan or cutin) while decreasing the relative content of polysaccharides. Given that cutin is important in plant-pathogen interaction (Yeats and Rose 2013), toxic compounds such as terpenoids or phenolics can be induced by wounding and herbivory (Kessler and Baldwin 2002). Therefore, the increased cutin and phenolics content reflect that cuticle is involved in plant defence system through changing its composition other than thickness. In addition, tomato showed a comparatively larger increase in phenolics and cutin. Given that we have found that tomato had four types of glandular trichomes (more than soybean and sunflower ) and its glandular trichomes density increased significantly compared to non-glandular trichomes (Figure 2-4), this observation might potentially be because some of the glandular trichomes contained or excreted phenolics.

Conclusion In conclusion, application of MeJA to the above-ground tissues of soybean, sunflower and tomato had a substantial influence on plant growth and leaf properties. It was found that MeJA inhibited plant growth, evident through decreases in plant height and biomass. However, it was also observed that MeJA increased leaf trichome density and altered cuticle composition – the magnitude of these changes in plant growth and leaf properties increased with increasing concentrations of MeJA, and the sensitivity of plant to MeJA was different among the three species. In addition, depending upon the concentration of MeJA and the plant

50 species, it was also observed that MeJA altered stomata density, epidermal cell wall and cuticle thickness, and leaf thickness. Thus, in addition to the commonly-studied effects of changes in trichome density, studies utilizing MeJA should also consider the importance of changes in other leaf properties and plant growth.

Acknowledgments This research was supported by Sonic Essentials (Melbourne, Australia) and through Australian Research Council’s Linkage Projects funding scheme (project number LP130100741). Support was also provided to Peter Kopittke as a recipient of an ARC Future Fellowship (FT120100277) and to Cui Li who received support from the China Scholarship Council and The University of Queensland (UQ International Scholarship).

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Figure 1. Effects of methyl jasmonate (MeJA) on plant height (a), root length (b), shoot biomass (c), root biomass (d), and leaf area of the youngest fully-expanded leaves (YFELs) (e) of soybean, tomato, and sunflower. In (a-d), the two-way ANOVA indicated a significant interaction; in (e), significant differences were only found among the three species, with LSD (5 %) values shown accordingly.

Figure 2. Scanning electron micrographs showing the effects of methyl jasmonate on trichome density of soybean leaves, examined either at the base, middle, or tip. The scale bar applies to all images. GT refers to glandular trichomes and NGT refers to non-glandular trichomes.

Figure 3. Scanning electron micrographs showing the effects of methyl jasmonate on trichome density of sunflower leaves, examined either at the base, middle, or tip. The scale bar applies to all images. GT refers to glandular trichomes and NGT refers to non-glandular trichomes.

Figure 4. Scanning electron micrographs showing the effects of methyl jasmonate on trichome density of tomato leaves, examined either at the base, middle, or tip. The scale bar applies to all images. GT refers to glandular trichomes and NGT refers to non-glandular trichomes.

Figure 5. Effects of methyl jasmonate (MeJA) on leaf non-glandular and glandular trichome density of soybean (a, b), tomato (c, d) and sunflower (e, f). For glandular trichomes of sunflower (d), the two-way ANOVA indicated a significant interaction among leaf position (base, middle and tip) and MeJA concentration (0, 0.1, 0.5, 1 and 2.5 mM). Elsewhere, the interaction was not significant, with LSD values presented for each factor individually where significant differences were found.

Figure 6. Effects of methyl jasmonate (MeJA) on stomatal density of soybean (a), tomato (b) and sunflower (c). For soybean, the two-way ANOVA indicated a significant interaction among leaf position (base, middle and tip) and MeJA concentration (0, 0.1, 0.5, 1 and 2.5 mM). For sunflower and tomato, the interaction was not significant, with LSD values presented for each factor individually.

Figure 7. Light micrographs showing cross sections of leaves of untreated soybean, sunflower and tomato, stained with toluidine blue. The scale bar applies to all images.

Figure 8. Effects of methyl jasmonate (MeJA) on the thickness of the adaxial epidermal cell wall and cuticle (a), and overall leaf thickness (b). In both instances, the two-way ANOVA indicated a significant interaction between the two factors.

Figure 9. Fourier transform infrared spectroscopy (FTIR) spectra of isolated cuticles from the adaxial leaf surface of tomato, soybean and sunflower.

3.3 Paper 2

Li C, Wang P, Menzies NW, Lombi E, Kopittke PM (2017) Effects of changes in leaf properties mediated by methyl jasmonate (MeJA) on foliar absorption of Zn, Mn and Fe. Annals of Botany 120: 405-415. (https://doi.org/10.1093/aob/mcx063)

ORIGINAL ARTICLE Effects of changes in leaf properties mediated by methyl jasmonate (MeJA) on foliar absorption of Zn, Mn, and Fe

Running Title: Leaf properties and foliar nutrients absorption

Cui Li1, Peng Wang2, 1,*, Neal W. Menzies1, Enzo Lombi3, Peter M. Kopittke1

1 The University of Queensland, School of Agriculture and Food Sciences, St Lucia, Queensland, 4072, Australia; 2 Nanjing Agricultural University, College of Resources and Environmental Sciences, Nanjing, 210095, China; 3 University of South Australia, Future Industries Institute, Mawson Lakes, South Australia, 5095, Australia

* For correspondence. Email: [email protected]

Background and Aims Foliar fertilization to overcome nutritional deficiencies is becoming increasingly widespread. However, the processes of foliar nutrient absorption and translocation are poorly understood. The present study aimed to investigate how cuticular leaf properties affect the absorption of foliar-applied nutrients in leaf tissues. Methods Given that methyl jasmonate (MeJA) can cause alterations in leaf properties, we applied 1 mM MeJA to sunflower (Helianthus annuus), tomato (Solanum lycopersicum), and soybean (Glycine max) to assess changes in leaf properties. Using traditionally analytical approaches and synchrotron-based X-ray fluorescence microscopy, the effects of these changes on the absorption and translocation of foliar-applied Zn, Mn, and Fe were examined. Key Results The changes in leaf properties caused by the application of MeJA increased foliar absorption of Zn, Mn, and Fe up to 3- to 5-fold in sunflower but decreased it by 0.5- to 0.9-fold in tomato, with no effect in soybean. These changes in the foliar absorption of nutrients could be not explained by changes in overall trichome density, which increased in both sunflower (86%) and tomato (76%) (with no change in soybean). Similarly, the changes could be not attributed to changes in stomatal density or cuticle composition, given these properties remained constant. Rather, the changes in the foliar absorption of Zn, Mn, and Fe was related to the thickness of the cuticle and epidermal cell wall. Finally, the subsequent translocation of the absorbed nutrients within the leaf tissues was limited (< 1.3 mm) irrespective of treatment. Conclusions The present study highlights the potential importance of the combined thickness of cuticle and epidermal cell wall in the absorption of foliar-applied nutrients. This information will assist in increasing the efficacy of the foliar fertilization.

Key words: Biofortification; cuticle; foliar absorption; methyl jasmonate; nutrients; trichomes; sunflower; soybean; tomato

INTRODUCTION Foliar fertilization is becoming increasingly widely used, with this approach potentially being more environmentally-friendly and more efficient compared to soil-based applications of fertilizers (Fernández and Eichert, 2009). For example, Ram et al. (2016) found that foliar application of Zn significantly increased both grain Zn concentrations and grain yield for wheat (Triticum aestivum L.) and rice (Oryza sativa L.) when examined at 31 sites across seven countries. However, the processes of foliar absorption and the subsequent translocation of absorbed nutrients are not well understood. As a consequence, foliar fertilizers are generally not designed upon a physiological basis, and hence potentially suffer from various limitations. For instance, the use of soluble Zn salts for foliar fertilization can result in leaf scorching (Drissi et al., 2015), while chelated forms of Zn, such as Zn-EDTA, are under increased scrutiny due to associated environmental concerns (Oviedo and Rodríguez, 2003).

The penetration of nutrients through the leaf surface has been demonstrated in both living plants and isolated cuticle membranes (Koontz and Biddulph, 1957; Schlegel and Schönherr, 2001; Riederer and Friedmann, 2006; Fernández and Eichert, 2009; Du et al., 2015). The process by which nutrients are absorbed through aerial tissues differs from uptake through roots, with the cell walls of leaves covered by a cuticle. Given that the cuticle is hydrophobic, it has been proposed that the cuticle has two separate pathways responsible for the transport of lipophilic and hydrophilic substances (Buchholz, 2006; Schönherr, 2006). The process by which lipid-insoluble (hydrophilic) ions penetrate through the cuticle has been studied extensively (Koontz and Biddulph, 1957; Schönherr, 1976; Buchholz et al., 1998; Schreiber, 2001; Schlegel et al., 2005, 2006; Schreiber, 2005; Fernández et al., 2014). It is generally assumed that the cuticle has small aqueous pores that can allow hydrated ions to pass. Although this hypothesis has been proven theoretically (Schönherr, 2006), the chemical composition, structure, and existence of such pores have never been directly confirmed. Through the use of fluorescent dyes and silver nitrate to localize the foliar-absorption pathway, it has also been proposed that the aqueous pores preferentially occur in guard cells, trichome bases (especially glandular trichomes), and cuticular anticlinal walls (Schlegel et al., 2005; Schönherr, 2006). Therefore, it is possible that the absorption of nutrients across leaf surfaces may occur via (i) the cuticle, (ii) cuticular cracks and imperfections, or (iii) stomata, trichomes, and lenticels (Fernández et al., 2013).

It is known methyl jasmonate (MeJA), as a phytohormone, is involved in plant defence systems and can cause alterations in leaf properties, such as trichome density (Tian et al., 2012). In the present study, using sunflower (Helianthus annuus), tomato (Solanum lycopersicum), and soybean (Glycine max), we utilized 1 mM MeJA to alter leaf properties and examined whether changes in leaf properties influence foliar absorption and the subsequent translocation of Zn, Fe, and Mn. Firstly, the effects of 1 mM MeJA on leaf properties were assessed by examining changes in thickness of the cuticle (measured as the combined thickness of the cuticle and epidermal cell wall, see later), cuticle composition, trichome density, and stomatal density. Next, the absorption of foliar-applied nutrients was assessed by examining changes in bulk nutrient concentrations of the leaf tissues. Finally, synchrotron-based X-ray florescence microscopy (μ-XRF) was used to provide in situ analyses of the distribution of nutrients within hydrated and fresh leaf tissues. The results of the present study will assist in understanding the process by which nutrients move across the leaf surface, and thereby provide underlying information required for improving the efficacy of foliar fertilizers.

MATERIALS AND METHODS Experimental design and plant growth The plant-growth study was conducted at The University of Queensland (St Lucia, Australia), in a growth room at 25 °C with high-pressure sodium lamps providing light (photon flux density of 500 μmol m–2 s–1) for 14 h d-1. Seeds of soybean (cv. Bunya), sunflower (cv. Hyoleic 41), and tomato (cv. Red Luck) were germinated in rolled paper towel moistened with tap water for either 3 (soybean and sunflower) or 4 d (tomato) before being transferred to 11 L black buckets. Each bucket had four holes in the lid with a total of either eight (tomato and soybean) or four (sunflower) plants, with each bucket forming one experimental unit. The - buckets were filled with a basal nutrient solution which contained (µM): 910 N (94 % NO3 + and 6 % NH4 ), 475 K, 20 P, 1126 Ca, 227 Mg, 1251 Cl, 556 S, 25 Fe(III)EDTA, 3 B, 0.5 Mn, 0.5 Zn, 0.2 Cu, and 0.01 Mo (Blamey et al., 2015). Nutrient solutions were changed after the first week, and then after every 4 d. After the plants had been growing for 10 d, 5 mL of

44 mM KH2PO4 was added to each 11 L bucket every second day thereafter to progressively replace P which had been taken up by the plants.

The experiment consisted of three plant species and two concentrations of MeJA [0 (control) and 1 mM] with four replicates, yielding a total of 24 experimental units. Application of the

MeJA occurred after the plants had been growing for 7 (sunflower), 8 (soybean) or 10 d (tomato). The MeJA (0 or 1 mM in 0.8 % ethanol) was sprayed onto the foliage until the leaves were fully saturated (ca. 300 mL per container) (Boughton et al., 2005; Tian et al., 2012), with the growth lamps switched off for 4 h following the application of the MeJA. Thereafter, plants were grown until the youngest leaves were fully-expanded (ca. 10 d after application of MeJA). The youngest fully-expanded leaves (YFELs) were then used for examining leaf morphology (stomatal and trichome density), the thickness of the cuticle and epidermal cell wall, and cuticle composition.

Assessment of leaf characteristics Using the YFELs, leaf (adaxial) stomatal density and trichome density were examined using scanning electron microscopy (SEM). Fresh materials (ca. 3 mm × 3 mm) were collected from the YFELs of each treatment and immediately fixed in 3 % glutaraldehyde in 0.1 M sodium cacodylate, followed by a microwave processing which was performed using a Pelco BioWave (Ted Pella Inc., California, USA) with a ColdSpot water recirculating device. Then the samples were post-fixed with 1 % osmium tetroxide, subjected to a dehydration series using ethanol (30, 50, 70, 90, 100, and 100 %) and processed using the BioWave. The tissues were then processed using a critical point dryer (Autosamdri-815 CPD, USA) before being coated with Au. Finally, samples were examined using a SEM (JEOL NEOSCOPE, Japan) at 10 kV accelerating voltage, with the density of trichomes and stomata determined.

Cuticle composition was examined using Fourier transform infrared spectroscopy (FTIR). Briefly, the abaxial side of each YFEL was scraped as clean as possible using a scalpel, and the remaining adaxial cuticles were placed into an enzyme solution containing 2 % pectinase and 0.1 % cellulase in 50 mM sodium acetate buffer to remove the residual cell debris from the cuticle. This enzymatic digestion was performed overnight at room temperature (25 °C) before the cuticles were washed twice using deionized water. The samples were examined using light microscopy to ensure that the underlying tissues were completely removed before being air-dried and analysed by FTIR (the obtained spectrum were then normalized) (Guzmán et al., 2014).

Although the cuticle has traditionally been thought to be independent of the underlying epidermal cell wall, it has recently been proposed that the cuticle is an extension of the epidermal cell wall (Guzmán et al., 2014; Fernández et al., 2016). Therefore, in the present

71 study, we measured the combined thickness of the cuticle and epidermal cell wall using light microscopy by examining sections cut from resin-embedded leaf samples. Briefly, the fresh YFELs of each treatment were cut into pieces ca. 1 mm × 4 mm. The segments were fixed in 3 % glutaraldehyde in 0.1 M sodium cacodylate at 4 °C overnight, rinsed twice with 0.1 M sodium cacodylate, post-fixed with 1 % OsO4 in 0.1 M sodium cacodylate for 4 h, and dehydrated in a graded ethanol series of 20, 30, 40, 50 (10 mins each), 70, 90, 100 (30 mins each), and 100 % (overnight). A series of graded Epon mixtures (Epon in ethanol) were used to infiltrate the tissues: 10, 20, 30, 40 (90 mins), 50 (overnight), 65, 80 (4 h), and 100 % (overnight). The tissues were then embedded in 100 % Epon blocks and polymerized at 60 °C overnight. Sections (1 µm thick) were cut using an ultramicrotome (LEICA EM UC6), stained by Toluidine blue (0.5 % in 1 % borax) and observed by light microscopy (Obrien, 1995).

Foliar-application of Zn, Mn, and Fe Bulk leaves (six replicates) were analysed by inductively coupled plasma mass spectrometry (ICP-MS) in order to obtain total elemental concentrations. A half-leaf loading method was used (Vu et al., 2013), with half of each leaf receiving nutrients and the other half receiving deionized water. To increase contact with the leaf surface, 0.05 % Tween 20 was used as a mild non-ionic surfactant (Reuveni et al., 1994). Specifically, to one half of each leaf, 20 -1 droplets (5 μL per droplet) of 1000 mg Zn L (15.4 mM, pH 5.2, supplied as ZnSO4.7H2O),

Mn (18.2 mM, pH 4.1, supplied as MnSO4.4H2O), and Fe (17.9 mM, pH 3.8, supplied as

FeSO4.7H2O) were applied with 0.05 % Tween 20, whilst the other half of each leaf received 60 droplets (5 μL per droplet) of deionized water with 0.05 % Tween 20 (pH 5.3). The leaves were incubated in Petri dishes for 6 h before been cut from the plants and then blotted dry with filter paper. Thereafter, the leaves were cut in half along the mid-vein, rinsed separately using 2 % HNO3, followed by 3 % ethanol, and deionized water (Du et al., 2015). The samples were oven dried, digested using a 5:1 mixture of nitric acid and perchloric acid, and analysed using ICP-MS. Blanks and reference materials were included to ensure accuracy.

Finally, to examine the in situ distribution of foliar absorbed Zn, Mn, and Fe in leaf tissues, the same three plant species were grown at La Trobe University (Bundoora, Australia) and exposed to either 0 or 1 mM MeJA as described earlier. Two droplets (5 μL each) of each nutrient (1000 L-1 of Zn, Mn or Fe with 0.05 % Tween 20) were applied to the adaxial YFELs of each of the three plant species. For sunflower and soybean, six droplets were applied near the tip of the leaf with the three nutrients applied on one side of the midrib and three replicate

72 droplets on the other side of the midrib. For tomato, the six droplets were applied on the base, middle or tip of a leaflet of a compound leaf with three nutrients on each side (see later). Following application of the droplets, the leaves were sealed inside in Petri dishes containing moist filter paper for 6 h, with lights on. For this entire incubation period, the leaves remained attached to the plants, with petiole passing through a small hole in the side of the Petri dish [Supplementary Information Fig. S1]. After incubation for 6 h, leaves were cut and rinsed thoroughly (as described earlier) before analysis using µ-XRF at the XFM beamline at the Australian Synchrotron (Melbourne, Australia).

Details of the XFM beamline as used to analyse plant tissues have been provided previously (Kopittke et al., 2011; Paterson et al., 2011). Briefly, X-rays were selected by a Si (111) monochromator and focused (ca. 2 µm × 2 µm) on the specimen by a pair of Kirkpatrick-Baez mirrors. The X-ray fluorescence emitted by the specimen was collected using the 384-element Maia detector system in a backscatter position at an excitation energy of 12,900 eV. Washed leaves were carefully blotted dry and mounted between two pieces of 4-µm thick Ultralene film, forming a tight seal around the leaf to limit dehydration. There was generally < 5 min between excision of the leaf and commencement of the µ-XRF analysis. For each sample, two scans were performed. The first scan (‘survey scan’) was comparatively rapid and aimed to examine the entire sample and identify the portion of the leaf surface to which the Zn, Mn, and Fe had been applied. The second scan (‘detailed scan’) was performed on a smaller area of the tissues surrounding the portion of the leaf to which the nutrients had been applied. This detailed scan was conducted with a smaller step size to increase spatial resolution (see details below).

Elemental mapping of each specimen involved continuous ‘on the fly’ analysis in the horizontal direction and discrete steps in the vertical direction. For the survey scan, the step size (i.e. pixel size) was 200 µm with a horizontal stage velocity of 18 mm s-1 (resulting in a pixel transit time of 11 ms). For the detailed scan, the step size was 20 µm with a horizontal stage velocity of 3 mm s-1 (resulting in a pixel transit time of 6.7 ms). The full X-ray fluorescence spectra were analysed using the CSIRO Dynamic Analysis method in GeoPIXE (http://www.nmp.csiro.au/dynamic.html) which enables quantitative, true-elemental images (Ryan, 2000; Ryan and Jamieson, 1993). All images were corrected for variations in leaf thickness through the use of Compton scattering as an internal standard.

Statistical analyses Data were analysed using IBM SPSS Statistics version 24 (IBM Corporation, New York). Comparisons between means were made using the T-test (95 %).

RESULTS Effects of methyl jasmonate on leaf properties Application of 1 mM MeJA resulted in changes in leaf properties, with these changes varying among the three plant species. Firstly, in regard to trichome density, it was observed that 1 mM MeJA significantly increased non-glandular trichome density in sunflower by 56 %, glandular trichome density in tomato by 134 %, but had no significant effect on trichome density in soybean (Fig. 1). Secondly, in contrast to trichome density, 1 mM MeJA had no significant effect on stomatal density in any of the three plant species (Fig. 2). Next, consideration was given to the thickness of the cuticle and epidermal cell wall. For sunflower, application of 1 mM MeJA significantly decreased thickness by 35 %, but in direct contrast, the thickness of the cuticle and epidermal cell walls in tomato actually increased by 30 %, whilst no significant differences were observed for soybean (Fig. 3). Finally, FTIR was used to examine changes in cuticle composition, but no marked differences were observed following the application of 1 mM MeJA for any of the three plant species (Fig. 4).

Comparison of foliar absorption of Zn, Mn, and Fe Upon detailed quantitative examination of bulk tissue concentrations using ICP-MS (Table 1), it was found that foliar absorption occurred in all treatments, although differences were observed depending upon both the plant species and the MeJA treatment (control or 1 mM MeJA). Specifically, for soybean, the application of 1 mM MeJA did not have a significant influence on foliar absorption for any of the three nutrients (Table 1). In contrast, for sunflower, treatment with 1 mM MeJA resulted in marked increases (ca. 3- to 5- fold) compared to the controls (Table 1), whilst for tomato, treatment with 1 mM MeJA actually decreased absorption of the three nutrients by 50 to 90 % (Table 1). Across the control treatments of the three plant species, soybean (< 1 μg leaf-1) had the lowest levels of absorption for all the three nutrients, followed by sunflower (ca. 4 – 6 μg leaf-1), and tomato (ca. 5 - 12 μg leaf-1) (Table 1).

Using μ-XRF analysis, it was found that foliar-application of Zn, Mn, and Fe resulted in substantial increases in their concentrations in the underlying leaf tissues, indicating that after 6 h, the three nutrients had crossed the leaf cuticle and penetrated into leaf tissues of soybean (Fig. 5A), sunflower (Fig. 5B), and tomato (Fig. 5C). In these tri-colour images (Fig. 5), red (Zn), green (Mn), and yellow (Fe) indicate the concentrations of these nutrients. It was noted that, below the site of application, the concentration of the nutrients in the veins were higher than the surrounding interveinal tissues. Indeed, for all three nutrients in both sunflower and tomato, projected concentrations in the veins were 2- to 4-fold higher than the corresponding interveinal tissues [Supplementary Information Fig. S2 and Fig. S3].

Effects of MeJA on subsequent movement of nutrients following their absorption Using µ-XRF, it was possible to examine the movement of Zn, Fe, and Mn by comparing their concentrations at increasing distance from the surface-applied droplets (Fig. 6). The distance that nutrients moved in the interveinal tissues and veins was calculated based upon measurements of droplet size from light microscopy and from maps of elemental distribution obtained from µ-XRF analyses (Fig. 6). As shown in Table 2, except for a significant reduction in the movement of Fe in the veins of sunflower, MeJA had no significant effect on the movement of Zn, Fe, or Mn in either the interveinal tissues or the veins from the site of application. In all instances, the extent to which Zn, Fe, and Mn moved from the site of their application was small, ranging from 0.22 to 1.32 mm regardless of the plant species or the nutrient, with the movement distance in veins being slightly greater than in the interveinal tissues. For both sunflower and tomato, it was noted that Mn had a similar movement distance within the vein and the interveinal tissues, whereas Zn had a greater movement distance within the vein (ca. 2-fold longer) than the interveinal tissues (Table 2 and Fig. 7). However, of the three nutrients, Mn had a slightly greater movement in interveinal tissues (mean 0.76 mm) than Zn (mean 0.45 mm) and Fe (mean 0.32 mm) (Table 2).

DISCUSSION Given that the leaf surface is covered with a cuticle, the underlying process whereby foliar- applied nutrients are absorbed through the leaf surface has been debated for many years. In the present study, we have utilized MeJA to alter leaf properties in order to examine the effects of leaf properties on foliar nutrient absorption. We found that foliar nutrient absorption was not related to overall trichome density – the application of 1 mM MeJA increased trichome density in both sunflower (86 %) and tomato (76 %) (Fig. 1), but the foliar

75 absorption of Zn, Mn, and Fe increased for sunflower but actually decreased in tomato (Table 1). Similarly, changes in foliar absorption could not be attributed to changes in either stomatal density or cuticle composition (Figs. 2 and 4), given that these leaf properties remained constant. Rather, we suggest that the foliar absorption was related to the combined thickness of the cuticle and epidermal cell wall. Indeed, in sunflower, foliar absorption of the three nutrients increased by 300 to 530 %, corresponding to a decrease in the cuticle and epidermal thickness from 3.0 to 1.9 μm (Table 1 and Fig. 3). Similarly, an increase in the thickness of the cuticle and epidermal cell wall from 1.9 to 2.5 μm was accompanied by a decrease (50 to 90 %) in the absorption of the three nutrients in tomato (Table 1 and Fig. 3). Finally, in soybean, we observed no changes in either nutrient absorption or in the thickness of the cuticle and epidermal cell wall. We also found that following their movement across the leaf surface, the absorbed Zn, Fe, and Mn moved only a very limited distance (0.22 to 1.32 mm) from the site of application before concentrations decreased to background levels, with MeJA (and the concomitant changes in leaf properties) having no effect on this translocation regardless of the plant species (Table 2).

The role of the cuticle in the foliar absorption of nutrients Given that the cuticle is generally defined as a lipid-rich layer with its outer compartment dominated by waxes (Koch and Ensikat, 2008; Fernández et al., 2013), it is generally assumed that the cuticle is a layer distinct from the underlying epidermal cell wall. As a result, almost all studies examining the foliar absorption of nutrients have utilized isolated cuticles, with the underlying epidermal cell wall not taken into account (Baur et al., 1997; Riederer and Schreiber, 2001; Schreiber, 2005; Buchholz, 2006; Riederer and Friedmann, 2006; Schönherr, 2006; Jetter and Riederer, 2016). However, given that it has recently been reported that the cuticle and epidermal cell wall have similar chemical constitution and function, it has been proposed that the cuticle was actually an extension of the epidermal cell wall region (Guzmán et al., 2014; Fernández et al., 2016). To the knowledge of the authors, the present study is the first to relate changes in foliar absorption of nutrients (Zn, Mn, and Fe) to the combined thickness of the epidermal cell wall and thickness of cuticle.

We found that the absorption of these three nutrients, in all the three plant species, was related to changes in cuticle and epidermal cell wall thickness (Fig. 3 and Table 1). Nevertheless, in contrast to these findings, some previous studies have suggested that cuticle thickness is not related to the cuticular permeability, although it must be noted that these previous studies

76 have utilized isolated cuticles without also considering the underlying epidermal cell wall (Riederer and Schreiber, 2001; Jetter and Riederer, 2016).

In this present study, we also found differences in foliar nutrients absorption among the three plant species. Overall, soybean (ca. 0.1 - 0.7 μg per leaf) had a markedly lower absorption of the three nutrients than sunflower or tomato (ca. 4 - 12 μg per leaf) (Table 1). We suggest that this is potentially because the soybean cuticle has a higher wax content than sunflower and tomato, as evidenced by the FTIR analyses [Supplementary Information Fig. S4], with a relatively high content of waxes (two bands at c. 2900 cm-1) and a low content of polysaccharides (the band at 3400 cm-1) which have been reported previously in soybean compared with sunflower and tomato (Guzmán et al., 2014; Heredia-Guerrero et al., 2014). Also, we noted that it was substantially more difficult to apply Supplementary Information the droplets to the leaves of soybean than it was to either sunflower or tomato because of higher surface tension (Fig. 5A), consistent with the analyses using SEM revealing a waxy layer covering leaves of soybean [Fig. S5]. This is in agreement with the previous studies that have reported the role of cuticular waxes in determining the permeability of plant cuticles (Riederer and Schreiber, 1995; Zhang et al., 2005; Bondada et al., 2006). In addition, by using µ-XRF, we found that the nutrient concentrations in the tissues underlying the droplets were higher in veins compared with the interveinal tissues (Figs. 6 and 7). Given to the low mobility of nutrients in the leaf tissues, this could be due to nutrient absorption being higher in veins than the interveinal leaf tissues.

Leaf cuticle composition is expected to be important for the foliar absorption of nutrients. Indeed, it has been reported that MeJA increased polysaccharide content in the callus of Rhodiola sachalinensis (Li et al., 2014) and Dendrobium officinale (Yuan et al., 2016) through upregulating the metabolism enzymes and biosynthetic genes of sucrose. Similarly, MeJA can influence the lipid metabolism, especially for fatty acyls metabolism (Cao et al., 2016). In the present study, however, FTIR analysis did not identify any marked changes in cuticle composition among the three plant species upon treatment with MeJA (Fig. 4). The effects of MeJA on leaf properties varied among different species. For example, MeJA increased the trichome density of sunflower and tomato, but did not change the leaf trichome for soybean (Fig. 1). Therefore, the effects of MeJA on leaf cuticle composition are likely species-dependent.

The role of trichomes in the foliar absorption of nutrients Trichomes are appendages that originate from epidermal cells and develop outwards on the surface of various plant organs, classified either as non-glandular or glandular trichomes, with one of the well-accepted functions of being protecting the plant from herbivories (Werker, 2000). In the present study, the foliar absorption was not related to the overall trichome density – although application of 1 mM MeJA increased the overall trichome density in both sunflower (86 %) and tomato (76 %) (Fig. 1), foliar absorption of Zn, Mn, and Fe increased for sunflower but actually decreased in tomato (Table 1). However, the changes in nutrient absorption were consistent with changes in the densities of non-glandular trichomes. Specifically, 1 mM MeJA increased non-glandular trichome density of sunflower by 56 % while decreased non-glandular trichome density of tomato by 62 %, and no changes occurred in non-glandular trichome density of soybean under the application of MeJA. We contend, however, that it is unlikely that these non-glandular trichomes play an important role in the foliar absorption of nutrients based on the fact that non-glandular trichomes are just appendages at leaf surface which have no connections to the inside leaf tissues. However, it has been suggested that trichomes can change the surface roughness and in turn influence leaf wettability (Bickford, 2016). This could potentially impact upon the foliar absorption of nutrients, with leaf wettability decreasing with increasing trichome density (Brewer et al., 1991). In the present study, the non-glandular trichomes of both sunflower and tomato were larger than the glandular trichomes [Supplementary Information Fig. S6] and therefore non- glandular trichomes would be expected to have a larger impact on leaf wettability than glandular trichomes. Regardless, the change in leaf wettability induced by trichome density could not explain differences in foliar absorption of nutrients in the present study, due to the fact that although leaf wettability decreased in sunflower (and increased in tomato), the foliar absorption of Zn, Mn, and Fe increased for sunflower (and decreased for tomato).

As trichomes vary markedly in structure, morphology, and function, it is difficult to generalize regarding the functions of all trichomes. Interestingly, several studies have examined foliar absorption by trichomes in epiphytic Bromeliaceae (Benzing et al., 1976; Benzing et al., 1985; Winkler and Zotz, 2010). These studies have, for example, reported that foliar absorption of water, Ca, and Zn was associated with leaf trichome densities of Bromeliaceae (Benzing and Burt, 1970; Ohrui et al., 2007; Winkler and Zotz, 2010). However, given that these epiphytic Bromeliaceae are generally highly drought-resistant and do not have absorbent roots, the leaf trichomes of Bromeliaceae could potentially be highly

78 specialized, with the peltate trichomes different to trichomes of other species (Ohrui et al., 2007).

Translocation of Zn, Fe, and Mn away from site of application The efficacy of foliar nutrient application not only depends upon the rate at which the nutrients move through the leaf surface, but also the mobility of nutrients from the application site of the treated leaves to other parts of the plant. The mobility of these nutrients depends upon three factors: (i) the ability of the nutrients to enter the phloem; (ii) the ability of the nutrients to move within the phloem; and (iii) the ability of the nutrients to move out of the phloem into the sink tissues (Fernández et al., 2013). In the present study, we examined the distribution of Zn, Fe, and Mn 6 h after their application. It was observed that these three nutrients moved only a limited distance from their site of application, regardless of the plant species. This result is in agreement with previous studies which have reported that Zn, Mn, or Fe have only a low mobility in plants (Eddings and Brown, 1967; Kannan and Charnel, 1986; Ferrandon and Chamel, 1988; Zhang and Brown, 1999; Marešová et al., 2012; Du et al., 2015). The reason for the low mobility of these nutrients remains unclear, although Fernández and Brown (2013) suggested that the low mobility of Zn may be due to the binding of Zn to the negative charge of the apoplastic space. Further studies are required to examine the location and movement of foliar-applied nutrients at a cellular and sub-cellular level.

In conclusion, using three plant species (soybean, sunflower, and tomato), we utilized 1 mM MeJA to induce changes in leaf properties in order to examine the effects of these changes on absorption of foliar-applied nutrients. For all three plant species, we found that the foliar absorption of Zn, Mn, and Fe was related to the thickness of the cuticle and epidermal cell wall. In contrast, foliar absorption of the nutrients was not related to changes in the total trichome density. In addition, using µ-XRF to provide in situ data from hydrated and fresh leaves, we found that the translocation of Zn, Mn, and Fe away from the sites of application was limited (0.22 to 1.32 mm) either in the vein or in the interveinal tissues, although the concentration of three nutrients within the underlying leaf tissues were higher in the veins than in the surrounding interveinal tissues.

ACKNOWLEDGEMENTS

This research was supported under Australian Research Council’s Linkage Projects funding scheme (LP130100741). Support was also provided to Peter Kopittke as a recipient of an ARC Future Fellowship (FT120100277) and to Cui Li who received support through the China Scholarship Council and through The University of Queensland (UQ International Scholarship). We thank Kathryn Green (Centre for Microscopy and Microanalysis, The University of Queensland) for assistance with the operation of the SEM and the ultramicrotome. Parts of this research were undertaken on the XFM beamline at the Australian Synchrotron (AS152/XFM/9487), Victoria, Australia, for which we thank the assistance of Daryl Howard.

Supplementary Information Supplementary Information Fig. S1. Foliar application of Zn, Mn, and Fe droplets on a sunflower leaf for X-ray fluorescence microscopy (μ-XRF) analysis. Supplementary Information Fig. S2. Compton-corrected μ-XRF images showing the distribution of Mn, Zn, and Fe in a control leaf of tomato after 6 h of foliar application. Supplementary Information Fig. S3. Compton-corrected μ-XRF images showing the distribution of Mn, Zn, and Fe in a control leaf of sunflower after 6 h of foliar application. Supplementary Information Fig. S4. Fourier transform infrared spectroscopy (FTIR) spectra obtained from isolated cuticles from the adaxial leaf surfaces of tomato, soybean, and sunflower (all controls). Supplementary Information Fig. S5. Scanning electron micrographs showing the surfaces of the leaves of soybean (A-B), tomato (C), and sunflower (D). Supplementary Information Fig. S6. Scanning electron micrographs showing non-glandular (NGT) and glandular trichomes (GT) on leaf surface of tomato (A) and sunflower (B).

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Kopittke PM, Menzies NW, de Jonge MD, et al. 2011. In situ distribution and speciation of toxic copper, nickel, and zinc in hydrated roots of cowpea. Plant Physiology, 156: 663-673. Li Y, Lian M, Shao C, Jin C, Piao X. 2014. Effect of methyl jasmonate on salidroside and polysaccharide accumulation in Rhodiola sachalinensis callus. Zhongguo Zhong yao za zhi= Zhongguo zhongyao zazhi= China journal of Chinese materia medica, 39: 4252-4257. Marešová J, Remenárová L, Horník M, Pipíška M, Augustín J, Lesný J. 2012. Foliar uptake of zinc by vascular plants: radiometric study. Journal of Radioanalytical and Nuclear Chemistry, 292: 1329-1337. Obrien S. 1995. Extrafloral nectaries in Chamelaucium uncinatum: a first record in the Myrtaceae. Australian Journal of Botany, 43: 407-413. Ohrui T, Nobira H, Sakata Y, et al. 2007. Foliar trichome-and aquaporin-aided water uptake in a drought-resistant epiphyte Tillandsia ionantha Planchon. Planta, 227: 47- 56. Oviedo C, Rodríguez J. 2003. EDTA: the chelating agent under environmental scrutiny. Quimica Nova, 26: 901-905. Paterson D, De Jonge M, Howard D, et al. 2011. The X-ray Fluorescence Microscopy Beamline at the Australian Synchrotron. the 10th international conference on X-ray microscopy: AIP Publishing. Ram H, Rashid A, Zhang W, et al. 2016. Biofortification of wheat, rice and common bean by applying foliar zinc fertilizer along with pesticides in seven countries. Plant and Soil, 403: 389-401. Reuveni R, Agapov V, Reuveni M, Raviv M. 1994. Effects of foliar sprays of phosphates on powdery mildew (Sphaerotheca pannosa) of roses. Journal of Phytopathology, 142: 331-337. Riederer M, Friedmann A. 2006. Transport of lipophilic non-electrolytes across the cuticle. Annual Plant Reviews, 23: 250-279. Riederer M, Schreiber L. 1995. Waxes: the transport barriers of plant cuticles. The Oily Press: Dundee, Scotland. Riederer M, Schreiber L. 2001. Protecting against water loss: analysis of the barrier properties of plant cuticles. Journal of Experimental Botany, 52: 2023-2032. Ryan C. 2000. Quantitative trace element imaging using PIXE and the nuclear microprobe. International Journal of Imaging Systems and Technology, 11: 219-230.

Ryan C, Jamieson D. 1993. Dynamic analysis: on-line quantitative PIXE microanalysis and its use in overlap-resolved elemental mapping. Nuclear Instruments and Methods in Physics Research Section B: Beam Interactions with Materials and Atoms, 77: 203- 214. Schlegel T, Schönherr J. 2001. Selective permeability of cuticles over stomata and trichomes to calcium chloride. International Symposium on Foliar Nutrition of Perennial Fruit Plants, 594: 91-96. Schlegel TK, Schönherr J, Schreiber L. 2005. Size selectivity of aqueous pores in stomatous cuticles of Vicia faba leaves. Planta, 221: 648-655. Schlegel TK, Schönherr J, Schreiber L. 2006. Rates of foliar penetration of chelated Fe (III): role of light, stomata, species, and leaf age. Journal of Agricultural and Food Chemistry, 54: 6809-6813. Schönherr J. 1976. Water permeability of isolated cuticular membranes: the effect of pH and cations on diffusion, hydrodynamic permeability and size of polar pores in the cutin matrix. Planta, 128: 113-126. Schönherr J. 2006. Characterization of aqueous pores in plant cuticles and permeation of ionic solutes. Journal of Experimental Botany, 57: 2471-2491. Schreiber L. 2001. Effect of temperature on cuticular transpiration of isolated cuticular membranes and leaf discs. Journal of Experimental Botany, 52: 1893-1900. Schreiber L. 2005. Polar paths of diffusion across plant cuticles: new evidence for an old hypothesis. Annals of Botany, 95: 1069-1073. Tian D, Tooker J, Peiffer M, Chung SH, Felton GW. 2012. Role of trichomes in defense against herbivores: comparison of herbivore response to woolly and hairless trichome mutants in tomato (Solanum lycopersicum). Planta, 236: 1053-1066. Vu DT, Huang L, V Nguyen A, et al. 2wo013. Quantitative methods for estimating foliar uptake of zinc from suspension-based Zn chemicals. Journal of Plant Nutrition and Soil Science, 176: 764-775. Werker E. 2000. Trichome diversity and development. Advances in Botanical Research, 31: 1-35. Winkler U, Zotz G. 2010. ‘And then there were three’: highly efficient uptake of potassium by foliar trichomes of epiphytic bromeliads. Annals of Botany, 106: 421-427. Yuan Z, Zhang J, Liu T. 2016. Enhancement of polysaccharides accumulation in Dendrobium officinale by exogenously applied methyl jasmonate. Biologia Plantarum: 1-7.

Zhang JY, Broeckling CD, Blancaflor EB, Sledge MK, Sumner LW, Wang ZY. 2005. Overexpression of WXP1, a putative Medicago truncatula AP2 domain-containing transcription factor gene, increases cuticular wax accumulation and enhances drought tolerance in transgenic alfalfa (Medicago sativa). The Plant Journal, 42: 689-707. Zhang Q, Brown PH. 1999. Distribution and transport of foliar applied zinc in pistachio. Journal of the American Society for Horticultural Science, 124: 433-436.

Fig 1. Effects of methyl jasmonate (MeJA) on leaf trichome densities. (A) – (C) Density of non-glandular and glandular trichomes following the application of 1 mM MeJA in soybean (A), sunflower (B), and tomato (C). Values are mean with SD (n=4). Within each plant species, P < 0.05 between 0 and 1 mM MeJA is indicated with an asterisk (*). (D) Scanning electron micrographs showing leaf trichome densities of 0 and 1 mM MeJA of soybean, sunflower, and tomato. The scale bar applies to all images.

Fig 2. Comparison of leaf stomatal density of 0 and 1 mM methyl jasmonate (MeJA) treated leaves of soybean, sunflower, and tomato.

Fig 3. Effects of methyl jasmonate (MeJA) on the thickness of cuticle and epidermal cell wall. (A) A light micrograph showing cross section of soybean leaf, with the red rectangle showing the area in (B); (B) Light micrographs showing the cuticle and epidermal cell wall thickness (the blue layer above the epidermal cells) of 0 and 1 mM MeJA treated leaves of soybean, sunflower, and tomato (the scale bar applies to all images); (C) Comparison of leaf cuticle and epidermal cell wall thickness of 0 and 1 mM MeJA treated leaves of soybean, sunflower, and tomato. Values are mean with SD (n=4). Within each plant species, P < 0.05 between 0 and 1 mM MeJA are indicated with an asterisk (*).

Fig 4. Fourier transform infrared spectroscopy (FTIR) spectra obtained from isolated cuticles from the adaxial leaf surfaces of tomato, soybean, and sunflower, either from 0 or 1 mM methyl jasmonate (MeJA) treated leaves.

Fig 5. Zn, Mn, and Fe distribution (after 6 h of foliar application) in control leaf of soybean (A) and tomato (C), and 1 mM methyl jasmonate (MeJA) treated leaf of sunflower (B). In each image, the upper images are light microscopy before µ-XRF analysis, with the orange rectangle indicating the area examined by µ-XRF. The images below are tricolor µ-XRF map of Zn (red), Fe (green), and Mn (blue) distribution.

Fig 6. Comparison of Mn movement through the vein and interveinal tissues of a tomato control leaf after 6 h of foliar application. (A) Light micrograph showing the Mn-containing droplet. (B) Same image as (A), showing the corresponding analysed area using µ-XRF in (C). White circles indicate the edge of the Mn droplet, while dashed lines represent a position of interveinal tissue, and solid lines indicate the position of a vein. The distance of ab = ac, namely, the white solid (vein) and dash (interveinal) line have same length in the droplet. (C) Compton-corrected Mn concentration of the area of (A) and (B). Brighter colour correspond to higher Mn concentrations. The projected Mn concentration underlying the solid (vein) and dash (interveinal) line were shown in (D). (D) Compton-corrected Mn concentration in vein and interveinal tissue of tomato control leaf with the shaded part (shaded distance = ab = ac) representing the section under the Mn droplet; According to the Mn concentration of background level, d1 is the Mn movement distance in the interveinal tissues, while d2 is Mn movement distance in the vein.

Fig 7. Comparison of nutrient diffusion in vein and interveinal tissues of sunflower control (A-C) and tomato control (D-E) leaves after 6 h of foliar application. The shaded area represents the portion of the leaf under the droplet. Soybean control and Fe for tomato control are not shown due to the comparatively low absorption or no detectable veins under the droplets.

Table 1. Comparison of foliar absorption of Mn, Zn, and Fe in soybean, tomato, and sunflower after 6 h of foliar application. Plants were treated with either 0 mM (control) or 1 mM methyl jasmonate (MeJA). Elemental absorption (μg leaf-1) = (elemental concentration in the treated half – elemental concentration in the control half) × dry weight of the treated half of the leaf. Nutrient absorption (μg leaf-1) Treatment Mn Zn Fe Soybean - MeJA 0.11 ± 0.05 A 0.11 ± 0.07 A 0.69 ± 0.27 A + MeJA 0.18 ± 0.06 A 0.26 ± 0.15 A 0.82 ± 0.03 A Sunflower - MeJA 6.45 ± 0.40 A 4.15 ± 0.35 A 5.77 ± 1.16 A + MeJA 25.6 ± 3.66 B 18.4 ± 3.22 B 36.5 ± 6.85 B Tomato - MeJA 7.44 ± 1.41 A 5.32 ± 0.93 A 11.5 ± 4.52 A + MeJA 3.69 ± 0.66 B 0.88 ± 0.12 B 1.47 ± 0.63 B Values are mean ± SD (n = 6). Where P < 0.05 between control and 1 mM MeJA within each species is indicated with different capital letters.

Table 2. Movement distance of Zn, Fe, and Mn in leaves of sunflower, tomato, and soybean after 6 h of foliar application

Movement distance (mm) Treatments Mn Zn Fe interveinal vein interveinal vein interveinal vein Sunflower - MeJA 0.72 ± 0.25 A 0.95 ± 0.19 A 0.53 ± 0.03 A 0.96 ± 0.26 A 0.41 ± 0.08 A 1.31 ± 0.19 A + MeJA 0.62 ± 0.02 A 0.79 ± 0.10 A 0.40 ± 0.22 A 0.83 ± 0.03 A 0.22 ± 0.02 A 0.27 ± 0.05 B Tomato - MeJA 1.32 ± 0.11 A 1.27 ± 0.14 A 0.43 ± 0.05 A 0.88 ± 0.13 A NA NA + MeJA 0.76 ± 0.23 A 1.01 ± 0.13 A 0.57 ± 0.10 A 0.95 ± 0.13 A NA NA Soybean - MeJA 0.44 ± 0.08 A NA 0.30 ± 0.04 A NA NA NA + MeJA 0.69 ± 0.12 A NA 0.46 ± 0.06 A NA NA NA Values are mean ± SD (n = 4). P < 0.05 occurred between control and 1mM MeJA for each species is indicated with different capital letters. NA data not available due to the comparatively low concentrations under the droplet.

SUPPLEMENTARY INFORMATION Effects of changes in leaf properties mediated by methyl jasmonate (MeJA) on foliar absorption of Zn, Mn, and Fe

Cui Li1, Peng Wang1, 2,*, Neal W. Menzies1, Enzo Lombi3, Peter M. Kopittke1

1 The University of Queensland, School of Agriculture and Food Sciences, St Lucia, Queensland, 4072, Australia; 2 Nanjing Agricultural University, College of Resources and Environmental Sciences, Nanjing, 210095, China; 3 University of South Australia, Future Industries Institute, Mawson Lakes, South Australia, 5095, Australia

* For correspondence. Email [email protected]

Fig. S1 Foliar application of Zn, Mn, and Fe droplets on a sunflower leaf for X-ray florescence microscopy (μ-XRF) analysis.

Fig. S2 Compton-corrected μ-XRF images showing the distribution of Mn, Zn, and Fe in a control leaf of tomato. For each nutrient, the relative concentration along a transect (indicated by the white, dashed line) is also presented, demonstrating the higher concentrations in the veins relative to the surrounding tissues.

Fig. S3 Compton-corrected μ-XRF images showing the distribution of Mn, Zn, and Fe in a control leaf of sunflower. For each nutrient, the relative concentration along a transect (indicated by the white, dashed line) is also presented, demonstrating the higher concentrations in the veins relative to the surrounding tissues.

Fig. S4 Fourier transform infrared spectroscopy (FTIR) spectra obtained from isolated cuticles from the adaxial leaf surfaces of tomato, soybean, and sunflower (all controls).

Fig. S5 Scanning electron micrographs showing the surfaces of the leaves of soybean (A-B), tomato (C), and sunflower (D).

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3.4 Conclusions

From the first experiment, it was found that MeJA increased the leaf trichome density in all three plant species (increasing density by 1.3-3.5 times), with the magnitude of the increase being related to the concentration of MeJA. However, it was also found that MeJA caused changes in leaf stomatal density, thickness of the epidermal cell wall and cuticle, and cuticle composition. The magnitude of these changes varied among plant species, but these changes were generally smaller than the changes in trichome density. In addition, it was found that plant height and biomass decreased significantly, particularly at the higher MeJA concentrations. Importantly, it was found that 1 mM MeJA significantly increased trichome density in sunflower and tomato but did not alter trichome density in soybean. It was also found that 1 mM MeJA had no impact on stomatal density or cuticle composition in any of the three plant species, but it significantly decreased the thickness of the cuticle and the epidermal cell wall in sunflower but increased it in tomato and had no impact in soybean.

Based upon these results, the second study examined how these changes in leaf properties altered the foliar absorption of Zn, Mn, and Fe, comparing in 0 and 1 mM MeJA in soybean, sunflower, and tomato. The foliar absorption of the three nutrients increased 3- to 5-fold in sunflower, but decreased by 0.5- to 0.9-fold in tomato, with no change in soybean. Comparing these changes in foliar nutrient absorption with the changes in leaf properties, it was found that the thickness of the cuticle and the epidermal cell wall were related to foliar nutrient absorption. These results suggest that the cuticle is important for foliar nutrient absorption. In addition, using of synchrotron-based µ-XRF, it was found that the subsequent translocation of the absorbed nutrients within the leaf tissues was limited (<1.3 mm away from the site of application) irrespective of treatment, with the movement distance in veins being slightly greater than in the interveinal tissues.

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Chapter 4 Absorption of foliar-applied Zn fertilizers by trichomes in soybean and tomato

4.1 Introduction

The previous Chapter highlighted the importance of the cuticle for the foliar absorption of Zn. However, whether trichomes also play a role in foliar Zn absorption remains unclear, with this being the focus of the present study. Using soybean and tomato, the leaf trichomes

were characterised using SEM. Then, using nano-ZnO and ZnSO4, the in situ distribution of foliar applied Zn in soybean and tomato was examined using µ-XRF across a time series to examine whether Zn accumulated within the trichomes. In addition, foliar Zn absorption was also compared using ICP–MS in four soybean near-isogenic lines (NILs) that varied 10-fold in trichome density to test the role of trichomes in foliar Zn absorption.

4.2 Paper 3

Li C, Wang P, Lombi E, Cheng M, Tang C, Howard DL, Menzies NW, Kopittke PM (2018) Absorption of foliar applied Zn fertilizers by trichomes in soybean and tomato. Journal of Experimental Botany 69: 2717-2729. (https://doi.org/10.1093/jxb/ery085)

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Role of leaf trichomes in absorption of Zn from three types of foliar- applied fertilizers in soybean and tomato

Cui Li1, Peng Wang1, 2, *, Enzo Lombi3, Miaomiao Cheng4, Caixian Tang4, Daryl L. Howard5, Neal W. Menzies1, Peter M. Kopittke1

1 The University of Queensland, School of Agriculture and Food Sciences, St Lucia, Queensland, 4072, Australia 2 Nanjing Agricultural University, College of Resources and Environmental Sciences, Nanjing, 210095, China 3 University of South Australia, Future Industries Institute, Mawson Lakes, South Australia, 5095, Australia 4 La Trobe University, Centre for AgriBioscience, Bundoora, Victoria, 3086, Australia 5 ANSTO, Australian Synchrotron, Clayton, Victoria, 3168, Australia

Email addresses for authors: Cui Li: [email protected] Peng Wang*: [email protected], Tel: +86 25 84399055, Fax: 86 25 84399055 Enzo Lombi: [email protected] Miaomiao Cheng: [email protected] Caixian Tang: [email protected] Daryl L. Howard: [email protected] Neal W. Menzies: [email protected] Peter M. Kopittke: [email protected]

Date of Submission: 13 November, 2017

Total word count (from Introduction to the end of acknowledgements): 6406 No. of Tables 0 No. of Figures 8 (Figures 1-5 in color) No of Supplementary Files 7 Figures (Figures 2-3, 5-7 in color)

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Role of leaf trichomes in absorption of Zn from three types of foliar- applied fertilizers in soybean and tomato Running title: Role of trichomes in foliar absorption of Zn

Highlight Leaf trichomes are not part of the primary pathway whereby foliar-applied Zn fertilizer moves across the leaf surface in soybean and tomato.

Abstract The present study investigated the role of trichomes in absorption of foliar-applied Zn fertilizers in soybean and tomato. Using synchrotron-based X-ray fluorescence microscopy for in situ analyses of hydrated leaves, we found that upon the foliar-application of ZnSO4, Zn accumulated within 15 min in some non-glandular trichomes of soybean, but not tomato. However, analyses of cross sections of soybean leaves did not show any marked accumulation of Zn in tissues surrounding trichomes. Furthermore, when near-isogenic lines of soybean differing 10-fold in trichome density were used to compare Zn absorption, it was found that foliar Zn absorption was not related to trichome density. Therefore, it is suggested that trichomes are not part of the primary pathway whereby foliar-applied Zn moves across the leaf surface in soybean and tomato. However, this does not preclude trichomes being important in other plant species, as trichomes are known to be highly diverse. We also compared the absorption of Zn when supplied as ZnSO4, with nano- and bulk-ZnO, finding that the absorption of Zn from ZnSO4 was ca. 10-fold higher than from nano- and bulk-ZnO, suggesting that Zn was mainly absorbed as soluble Zn. This study improves our understanding of the absorption of foliar-applied nutrients. Key words: Foliar absorption, foliar fertilizer, trichomes, Zn, soybean, tomato, nanoparticles

Abbreviations: Near-isogenic lines NILs Inductively coupled plasma mass spectrometry ICP-MS Synchrotron-based X-ray fluorescence microscopy μ-XRF Youngest fully-expanded leaves YFELs Relative humidity RH

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Introduction Zinc (Zn) is an essential micronutrient for the nutrition of both plants and humans. However, it has been estimated that 49 % of the world’s important agricultural soils have inadequate levels of Zn and that one-third of the world’s human population suffers from Zn deficiency (Sillanpää, 1990; Cakmak et al., 2010). Foliar fertilization is potentially an efficient approach for the correction of Zn deficiency in crops (Zhang et al., 2017; Jaksomsak et al., 2018). The use of foliar fertilizers is particularly useful for those soils which limit the uptake of Zn by the roots due to chemical constraints, such as alkaline soils in which Zn availability is low due to precipitation reactions (Cakmak, 2008; Cakmak et al., 2010). However, much remains unknown regarding the underlying processes whereby foliar-applied nutrients move across the leaf surface into the underlying plant tissue (Fernández et al., 2017; Li et al., 2017). This poor understanding of the physiological processes involved in the absorption of foliar-applied nutrients hinders efforts to increase the effectiveness of foliar fertilizers (Fernández et al., 2013). Of particular interest, it is unclear how nutrients move across the hydrophobic cuticle covering the leaf surfaces (Riederer and Schreiber, 2001; Fernández and Eichert, 2009; Fernández et al., 2017). For example, the importance of the stomata, cuticle, and trichomes in foliar nutrient absorption remains unclear (Schönherr, 2006; Fernández et al., 2013).

In the present study, we focus on the role of trichomes for the foliar-absorption of Zn. Trichomes are generally found on both the abaxial and adaxial surfaces of leaves, and can be found at high densities on the leaves of many plant species. For example, glandular trichomes have been reported on leaves of tomato (Solanum lycopersicum) at a density of 11,100 cm-2 (Maluf et al., 2007). Trichomes are appendages that originate from epidermal cells and develop outwards on the surface of various plant organs (Werker, 2000). Depending on the plant species, the growth environment, and the organs where they are located, trichomes vary considerably in structure, morphology, size, and function (Werker, 2000; Ohrui et al., 2007). Trichomes can be broadly classified as being either glandular or non-glandular, with the major distinction being that glandular trichomes can contain or excrete various chemical compounds (Tian et al., 2012). The most well known function of trichomes is for the protection of plants from herbivores, both through a physical interference and through the excretion of toxic or repellent compounds (Levin, 1973; Tian et al., 2012; Aschenbrenner et al., 2013; Stratmann and Bequette, 2016;). Although it has been reported for several plant species that trichome bases are cutinized (Fahn, 1986; Mahlberg and Kim, 1991; Ascensão et

105 al., 1999; Huang et al., 2008; Fernández et al., 2011; Wu et al., 201; Guo et al., 2013; Fernández et al., 2014), it has been reported that trichomes can absorb water in some plant species, for example, trichomes of cactus (Opuntia microdasys) were involved in its fog collection system (Ju et al., 2012) and trichomes of Phlomis fruticosa can absorb water under drought condition (Grammatikopoulos and Manetas, 1994). In contrast, Burrows et al. (2013) reported that the stellate trichomes of Solanum elaeagnifolium were water repellant and were likely to be barriers for absorption of foliar-applied chemicals. In addition, it was found that depending upon their density, structure and chemical composition, trichomes could influence leaf surface wettability and droplet retention ability. Specifically, higher trichome densities generally lead to lower wettability but higher droplet retention (Brewer et al., 1991; Grammatikopoulos and Manetas, 1994; Fernández et al., 2014;), and it was found that some trichomes have a higher water sorption capacity than the cuticle (Fernández et al., 2011). However, much uncertainty remains regarding the role (if any) of trichomes in the absorption of foliar-applied nutrients.

Given that trichomes are the outermost aerial structure of leaves, it is important to elucidate their role in the absorption of foliar-applied nutrients – this being the aim of the present study. Using soybean (Glycine max) and tomato, we first examined the morphology of trichomes on the leaf surface using scanning electron microscopy (SEM) and light microscopy. Next, we used synchrotron-based X-ray fluorescence microscopy (µ-XRF) for the in situ examination of fresh (hydrated) leaves to obtain laterally-resolved elemental maps showing the distribution of foliar-applied Zn within leaf tissues, with the spatial Zn distribution in soybean leaf also examined. Finally, for soybean, we used four near-isogenic lines (NILs) that varied 10-fold in trichome density to compare foliar Zn absorption by examining bulk leaf Zn concentrations. For this study, we also compared three types of Zn -1 fertilizers, being ZnSO4, nano-ZnO, and bulk-ZnO (all applied at 1000 mg Zn L ). This is important given that there is increasing interest in the potential use of nanoparticles as a foliar fertilizer (De Rosa et al., 2010; Nair et al., 2010; Khot et al., 2012; Wang et al., 2016). The results of the present study provide important information regarding the movement of nutrients across the leaf surface, with the aim to assist in improving the efficiency of Zn foliar fertilizers.

Materials and Methods

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Plant growth and assessment of leaf trichomes Plants were grown at The University of Queensland (St Lucia, Australia), at 25 °C, with artificial light provided by high pressure sodium lamps (photon flux density of 1500 μmol m– 2 s–1) for 12 h d-1. Seeds of soybean (cv. Bunya) and tomato (cv. Red Luck) were placed and rolled up in paper towel which suspended vertically in tap water (to keep moisture) for either 3 d (soybean) or 4 d (tomato). Seedlings were transferred to 11 L black buckets, with the lids having four holes in which the seedlings were suspended using shade cloth. Each bucket had eight plants (two plants per hole). The buckets were filled with a nutrient solution which - + contained (µM): 910 N (94 % NO3 and 6 % NH4 ), 475 K, 20 P, 1126 Ca, 227 Mg, 1251 Cl, 556 S, 25 Fe(III)EDTA, 3 B, 0.5 Mn, 0.5 Zn, 0.2 Cu, and 0.01 Mo (Blamey et al., 2015). Nutrient solutions were changed after one week, and then every 4 d. In addition, to replace P being taken up from solution, 5 mL of 44 mM KH2PO4 was added every second day once the plants had been growing in the nutrient solution for 10 d.

The youngest fully-expanded leaves (YFELs) were used to examine leaf trichome morphology after 14 (soybean) and 18 d (tomato) as outlined by Li et al. (2017). Briefly, the fresh YFELs were cut into small segments (2 × 4 mm), and then fixed in 3 % glutaraldehyde in 0.1 M sodium cacodylate with a microwave processing, followed by 1 % OsO4 in 0.1 M sodium cacodylate. Next, samples were dehydrated using an ethanol gradient with a Pelco BioWave (Ted Pella Inc., California, USA), dried using a critical point dryer (Autosamdri- 815 CPD, USA), and coated with Au. The trichomes were analyzed using an SEM (JEOL NEOSCOPE, Japan) at 10 kV accelerating voltage.

To examine trichome structure, cross sections of the leaves were cut and analyzed using light microscopy as described by Li et al. (2017). Briefly, the YFELs were cut in 1 mm × 4 mm segments and fixed overnight in 3 % glutaraldehyde with 0.1 M sodium cacodylate at 4 °C. Using 0.1 M sodium cacodylate, the samples were rinsed before being post-fixed in 1 %

OsO4 in 0.1 M sodium cacodylate for 4 h, and dehydrated in a graded ethanol series. The samples were infiltrated with a series of graded Epon mixtures (Epon in ethanol), and embedded in 100 % Epon blocks and polymerized at 60 °C overnight. An ultramicrotome (LEICA EM UC6) was used to cut the sections (1 µm) before being stained using toluidine blue (0.5 % in 1 % borax) (O'Brien, 1995).

Examining the distribution of Zn using µ-XRF 107

Soybean and tomato were grown at La Trobe University (Bundoora, Australia) to allow for their subsequent transport to the Australian Synchrotron (Clayton, Australia). Plants were grown as described earlier. After growth in the nutrient solution for 14 (soybean) and 18 d -1 (tomato), two droplets (5 µL each) of 1000 mg Zn L solution (supplied as ZnSO4.7H2O, 15.4 mM, pH 5.2) were applied to the adaxial surface of the YFELs. The droplets also contained 0.05 % Tween 20 to decrease surface tension (Reuveni et al., 1994). The leaves were sealed in Petri dishes which had a hole to allow for the petiole, with the leaves still attached to the plants. The Petri dishes also contained moist filter paper, with the droplets left on the leaves for 0.25, 0.5, 1, 3, or 6 h. During this time, the droplets did not dry out but remained as a liquid, with relative humidity (RH) inside the sealed Petri dish increasing rapidly to 98 % (temperature of 30 °C). Afterwards, the droplets were blotted dry with filter paper, and the leaves were excised and rinsed sequentially using 2 % HNO3, 3 % ethanol, and deionized water (Du et al., 2015).

The thoroughly-rinsed leaves were blotted dry and analysed using µ-XRF at the XFM beamline at the Australian Synchrotron (Clayton, Australia). The leaves were mounted tightly between two pieces of 4 µm thick Ultralene film which formed a seal around the leaf to limit dehydration. The time between excision of the leaf and commencement of the µ-XRF analysis was < 5 min. Basic information of the XFM beamline is given by Kopittke et al. (2011) and Paterson et al. (2011). Briefly, X-rays are selected by a Si (111) monochromator and focused (ca. 2 µm × 2 µm) on the specimen by a pair of Kirkpatrick-Baez mirrors. A 384-element Maia detector system in a backscatter position (180°) at an excitation energy of 12,900 eV was used to collect the X-ray fluorescence emitted by the specimen. For each specimen, a ‘survey scan’ and a ‘detail scan’ were performed. The survey scan was a comparatively quick scan which was used to allow for the selection of an area of interest; the step size (i.e. virtual pixel size) was 30 µm with a horizontal stage velocity of 4 mm s-1, resulting in a pixel transit time of 7.5 ms. For the subsequent detailed scan, the step size was 1 µm with a horizontal stage velocity of 1 mm s-1, resulting in a pixel transit time of 1 ms. Although it varied, the survey scans were often ca. 10 × 6 mm in size (resulting in a scan duration of ca. 10 min) and the detailed scans were often ca. 4 × 5 mm in size (resulting in a scan duration of ca. 75 min). The full X-ray fluorescence spectra were analyzed using the CSIRO Dynamic Analysis method in GeoPIXE (http://www.nmp.csiro.au/dynamic.html) (Ryan, 2000; Ryan and Jamieson, 1993).

108

For soybean, cross sections of the leaf tissues were also prepared for examining the distribution of Zn. The samples were prepared at The University of Queensland, with soybean plants grown for 14 d as described earlier. Then, a 40 µL droplet of 1000 mg Zn L-1

(supplied as ZnSO4.7H2O with 0.05 % Tween 20) was applied on the adaxial leaf surface before being incubated in Petri dishes for 0 (control), 0.25, 1, and 6 h. We applied a larger volume in this experiment (40 µL instead of 5 µL) in order to ensure that the resulting cross section that was cut was from an area entirely under the droplet. Thereafter, the area below where the Zn had been applied (or the corresponding position on control leaf) was excised after being rinsed, with these leaf segments embedded in 4 % agar solution at ca. 30 °C. After the agar solidified, the block was attached to the sample holder of a vibratome (Leica VT 1000s) using super glue, with 150 µm thick cross sections obtained without use of a buffering solution. Finally, the sections were sealed in the Ultralene film with XRF Sample cups (SC- 8047, Premier Lab Supply), small holes were made in the film, and the samples were freeze dried.

At the XFM beamline, the sections were carefully removed and placed between two sheets of Ultralene film (4 µm) and stretched over a Perspex frame which was magnetically mounted on the motion stage. A Vortex detector system in a 90° geometry at an excitation energy of 15,800 eV was used to collect the X-ray fluorescence emitted by the specimen. Again, a ‘survey scan’ was used for each sample to first identify the area of interest before obtaining a higher resolution image (‘detail scan’). For the ‘survey scan’, the step size was 50 µm with a horizontal stage velocity of 1 mm s-1, resulting in a pixel transit time of 50 ms. For the subsequent ‘detailed scan’, the step size was 2 µm with a horizontal stage velocity of 0.6 mm s-1, resulting in a pixel transit time of 3.33 ms. The survey scans often took ca. 3 min, and the detailed scans often took ca. 30 min.

Zinc absorption using soybean mutants Four NILs of soybean (cv. Clark) were obtained from the USDA Soybean Germplasm Collection, Department of Crop Sciences, USDA/ARS University of Illinois, Urbana IL, USA. The NILs we obtained were wild-type Clark (L58-231) (CS), sparse trichome (L63- 2999) (Ps), dense trichome (L62-1686) (Pd1), and glabrous (L62-1385) (P1). The plant growth experiment was conducted at The University of Queensland (St Lucia, Australia) as described earlier except that plants were grown in 5 L buckets with six plants in each bucket. Each of these buckets formed a single experimental unit, with six replicates per genotype. 109

For the current experiment we also compared three forms of Zn, with Zn supplied as

ZnSO4.7H2O, nano-ZnO (ca. 30 nm diameter), and its homologous bulk-ZnO (ca. 200-400 nm diameter) (Supplementary Fig. S1 and Table S1). The two ZnO forms (nano and bulk) were analysed using dynamic light scattering (DLS) using a zeta potential / particle sizer NICOMP 380 ZLS at a concentration of 333 mg Zn L-1. For this plant-growth experiment, all -1 three forms were used at a total Zn concentration of 1000 mg Zn L . Although ZnSO4.7H2O is highly soluble, the nano-ZnO and bulk-ZnO are only sparingly solubility, and hence although the total Zn concentration of the suspensions of these two compounds was 1000 mg Zn L-1, the concentration of soluble Zn would be lower than this value. Therefore, we measured the concentration of soluble Zn in suspensions of 1000 mg Zn L-1 nano- and bulk- ZnO. To do this, suspensions were mixed for 15 min in an ultrasonic bath before being filtered using centrifugal filter units (3 kDa, Millipore, Sigma). The Zn concentrations were measured in the filtered solutions using inductively coupled plasma mass spectrometry (ICP- MS). For both the nano- and bulk-ZnO suspensions, the Zn concentration was 3 mg Zn L-1 (46 µM). Therefore, to assist in comparing the 1000 mg Zn L-1 as nano- and bulk-ZnO -1 suspensions with 1000 mg Zn L as ZnSO4.7H2O solution, we also included a treatment with -1 3 mg Zn L as ZnSO4.7H2O. All solutions contained 0.05 % Tween 20. In addition, we examined two application times, being 1 and 6 h. Thus, the experiment consisted of a total of 32 treatments, with four NILs of soybean differing in trichome density, two application times -1 -1 (1 and 6 h), and four Zn treatments: 1000 mg Zn L as ZnSO4.7H2O, 3 mg Zn L as -1 -1 ZnSO4.7H2O (pH 5.2), 1000 mg Zn L as nano-ZnO (pH 7.1), and 1000 mg Zn L as bulk- ZnO (pH 7.0). Each treatment had six replicates, yielding a total of 192 experimental units.

After growth for two weeks, 30 droplets (5 μL each) of the appropriate Zn fertilizer was applied to half of the adaxial surface of the YFELs, with 30 droplets (5 μL each) of deionised water (with 0.05 % Tween 20) applied to the other half of the leaf (Vu et al., 2013). Following application of the droplets, the leaves were sealed inside Petri dishes with moistened filter paper, with the lights remaining switched on during the incubation period. As noted previously, RH inside the sealed Petri dishes increased rapidly to 98 %, and the droplets remained as a liquid (i.e. did not dry out) during the experimental period. After the appropriate length of time (1 or 6 h), the leaves were cut in half along the mid-vein, and rinsed separately using 2 % HNO3, 3 % ethanol, and deionized water. The leaf tissues were oven dried (temperature 65 °C), digested using a 5:1 mixture of nitric acid and perchloric 110 acid, and analysed using ICP-MS. Blanks and reference materials were included to ensure accuracy. The data in the present study were calculated as Zn absorption (μg half leaf-1) = (elemental concentration in the treated half – elemental concentration in the control half) × dry weight of the treated half of the leaf.

To allow for analyses of the trichome and stomatal density, each of the four NILs was grown for two weeks before analysis using the SEM as described earlier, with six replicates examined. Finally, as it has been proposed that trichomes may alter leaf surface wettability (Fernández et al., 2014), we measured the contact angle of 5 µL droplets for 1000 mg Zn L-1 -1 -1 as ZnSO4.7H2O, 1000 mg Zn L as nano-ZnO, and 1000 mg Zn L as bulk-ZnO (all with 0.05 % Tween 20) for the four NILs using the drop shape method (Woodward, 1999).

Statistical analyses Data were analyzed using IBM SPSS version 24 and GenStat version 18. Comparisons between means were made using the one-way analysis of variance least significant difference (LSD) at 5 %.

Results Types and morphology of trichomes on leaves of soybean and tomato For soybean, two types of trichomes were found (Fig. 1); a non-glandular trichome (Type V) and a glandular trichome (Type IX) (Franceschi and Giaquinta, 1983). The Type IX trichomes were ca. 0.04 mm in length and consisted of ca. four stalk cells with one small cell on the tip (Fig. 1A, C, and F). In contrast, the Type V trichomes were hollow, consisting of a multicellular base and one long stalk cell up to 1 mm in length (Fig. 1A,B,D,E). The overall trichome density of the soybean adaxial leaf surface was 1300 cm-2, with 65 % being non- glandular trichomes (Type V) (Fig. 1).

Tomato leaves had a total of seven types of trichomes, with four types of glandular trichomes (Types I, II, VI, and VII) and three types of non-glandular trichomes (Types III, IV, and V) (Fig. 2). The Type I trichomes were the smallest of the seven, consisting of a short stalk cell with a capitate round cell on the top (Fig. 2C). The Type II and VI trichomes were similar in length, with both having a unicellular base connected to a stalk cell, but the Type VI trichomes had a four-cell head (Fig. 2C,D,E). The Type III and IV trichomes were both non-

111 glandular and were similar in shape, with the Type III trichome having two stalk cells while the Type IV trichomes had only one long stalk cell (Fig. 2B,C,F,H). Type V trichomes were the largest trichome observed on the tomato leaf, consisting of a multi-cellular base connected with a long stalk (up to > 2 mm) (Fig. 2A,G). The Type VII trichomes were similar to Types II, but the Type VII trichomes had a small round cell on the trichome tip (Fig. 2D). Overall, the tomato adaxial leaf trichome density was 1,800 cm-2 , with about 48 % of the trichomes being either Type II or VI trichomes, followed by ca. 24 % of Type III and IV, then Types VI (ca. 13 %), I (ca. 9 %), V (ca. 5%), and < 1 % VII (Fig. 2).

Laterally-resolved elemental maps showing the movement of Zn into the leaf tissues Firstly, the distribution of Zn (and other elements) was analysed in situ within fresh (hydrated) leaf tissues of soybean and tomato using synchrotron-based µ-XRF following -1 exposure to 1000 mg Zn L of ZnSO4 for 15 min to 6 h. Initially, a rapid survey scan was used to allow for the selection of an area of interest, with these survey scans (Fig. 3 and Supplementary Fig. S2) showing that within only 15 min the Zn had already moved across the leaf surface and was present in the leaf tissues underlying the Zn droplets for both plant species. The concentration of Zn within the leaf tissues gradually increased as the length of exposure increased from 15 min to 6 h. It was also observed that elevated levels of Zn were found almost exclusively in the tissues underlying the droplets. In other words, the movement of Zn from the site of the application was limited.

Next, detailed scans were used to more closely examine Zn distribution in the tissues directly underlying the Zn droplets. Soybean and tomato had a similar Zn distribution pattern (Fig. 4), with Zn first observed to accumulate (i.e. within 15 min to 1 h) in the veins (Fig. 4A-C and F- H). In addition, for soybean, some Zn was also observed to accumulate in some of the basal cells of the non-glandular trichomes (Type V) (Fig. 4A-C). However, not all Type V trichomes underneath the droplet accumulated Zn. Assuming a trichome density of 1300 cm-2 (with 65% being Type V) (Fig. 1), it would be expected that there would be ca. seven Type V trichomes within the area shown in Fig. 4A-E, but it was only two Type V (ca. 29 %) trichomes could be identified based upon the accumulation of Zn (Fig. 4A-D and Supplementary Fig. S3). For both plant species after exposure for 3 h, Zn had accumulated in almost all of the underlying veins as well as in the interveinal tissues (albeit at a lower concentration) (Fig. 4D,I). After 6 h, the extent of accumulation increased further, and within

112 the interveinal tissues, high concentrations of Zn were found to accumulate in the apoplast (Fig. 4E,J).

Finally, to further examine the role of the trichomes in foliar Zn absorption in soybean, we also used µ-XRF to examine cross sections of leaves cut from tissues underneath where droplets of Zn had been applied (in these instances, the entire area of the cross sections had been obtained from underneath the Zn droplets). For these cross sections, concentrations of Zn increased after only 15 min and continued increasing up to 6 h (Fig. 5). Interestingly, although Zn was found to accumulate in some of the non-glandular trichome bases in soybean especially before 3 h of foliar application (Fig. 4), analysis of the cross sections by µ-XRF showed that the Zn concentration was not higher in the cells surrounding the base of the trichomes (Type V) (Fig. 5). This is likely because we could only observe a small number of trichomes on these cross sections, while only a limited number of trichomes (ca. 29 % Type V of the total Type V) observed to have elevated levels of Zn (Fig. 4A-D and Supplementary Fig. S3). In addition, Zn was detected in the inner tissues of the leaves (presumably the mesophyll cells), especially after 6 h application, implying that Zn had been transported into the leaf tissues after penetrated through the cuticle and epidermis (Fig. 5B). Given that the cross sections were 150 µm thick (ca. eight cellular layers deep) (Fig. 1E,F), it was not possible to identify determine subcellular distribution from the cross sections, and the apparent apoplastic accumulation observed from intact leaves (Fig. 4E) could not be confirmed from cross sections (Fig. 5).

Foliar absorption in soybean NILs as related to trichome density Given that Zn was observed to accumulate rapidly in some trichomes of soybean (Fig. 4), we used four soybean NILs (CS refers wild-type Clark L58-231, Ps refers sparse trichome L63- 2999, Pd1 refers dense trichome L62-1686, and P1 refers glabrous L62-1385) that varied 10- fold in trichome density to examine the potential importance of trichomes in the absorption of foliar-applied Zn. Given that stomata are also potentially important for the absorption of foliar-applied nutrients (Fernández et al., 2013), we determined the densities of both the trichomes and stomata for leaves of the four soybean NILs. As expected, trichome density differed markedly among the four NILs, with Pd1 having a trichome density (ca. 1100 cm -2) significantly higher than the other three NILs. Although Ps had the lowest trichome density (ca. 100 cm -2), it was not significantly different to either P1 (ca. 220 cm -2) or CS (ca. 190 cm -2) (Fig. 6A,C). The trichomes on leaves of Pd1, CS, and Ps were similar to those described 113 earlier in cv. Bunya (Fig. 1), with all three NILs having ca. 95 % Type V trichomes and ca. 5 % Type IX trichomes. In contrast, P1 had shorter Type V trichomes compared to Pd1, CS, and Ps, with 35 % of the trichomes for P1 being Type IX and the remainder being Type V (Fig. 6). The stomatal density was ca. 8,000-10,000 cm-2, with no significant differences among the four NILs (Fig. 6B).

Next, using ZnSO4, we compared differences in the absorption of foliar-applied Zn among the four soybean NILs. A half leaf loading method was used, with Zn applied to half the leaf and absorption calculated as the difference between the two halves of each leaf. After -1 exposure for 1 h to 1000 mg Zn L of ZnSO4, the quantity of Zn absorbed ranged from 0.7 to 2.5 μg half leaf-1, with no significant differences found among the four NILs (Fig. 7). After -1 exposure to ZnSO4 for 6 h, the amount of Zn absorbed increased to 25 - 45 μg half leaf . Although significant differences were found among the four NILs, these differences were not related to trichome density. Indeed, comparing Pd1, CS, and Ps (all of which had the same types of trichomes), it was found that although Ps had the lowest trichome density (110 cm-2) it had the highest Zn absorption (45 μg half leaf-1) (Figs. 6 and 8). In contrast, Pd1 (which had the highest trichome density among the three NILs, 1100 cm-2) had the second highest Zn absorption (35 μg half leaf-1) (Figs. 6 and 8).

Absorption of Zn using a nano-fertilizer Given the increasing interest in the use of nanoparticles as a Zn fertilizer, we also compared -1 three types of Zn fertilizers, being 1000 mg Zn L as ZnSO4, nano-ZnO (ca. 30 nm diameter), and bulk-ZnO (ca. 200-400 nm diameter) (Supplementary Fig. S1). To facilitate -1 comparison, the ZnSO4 was applied at two concentrations, being 1000 mg Zn L and 3 mg Zn L-1. This latter concentration of 3 mg Zn L-1 is equal to the concentration of soluble Zn measured in suspensions of both 1000 mg Zn L-1 of nano- and bulk-ZnO. After 6 h, leaves -1 exposed to 1000 mg Zn L of ZnSO4 had the highest rates of Zn absorption (25 – 44 μg half leaf-1), with absorption for leaves exposed to 1000 mg Zn L-1 nano- or bulk-ZnO being ca. 10-fold lower (1 – 7 μg half leaf-1). Interestingly, although the concentration of soluble Zn -1 was the same in the nano- and bulk-ZnO treatments as it was in the 3 mg Zn L ZnSO4 -1 treatment, the rate of absorption for leaves exposed to 3 mg Zn L ZnSO4 was up to ca. 10- fold lower (0.5 – 0.7 μg half leaf-1) than for leaves exposed to ZnO (Fig. 8). Across the different forms of Zn fertilizer, significant differences were found among the four NILs, but again, these differences were not related to trichome density (Figs. 6 and 8). 114

Discussion Foliar fertilization is an agronomic strategy of increasing interest, particularly for micronutrients such as Zn where low concentrations within edible plant tissues also impact adversely upon human nutrition (Fernández et al., 2013). However, despite their importance, the processes whereby foliar-applied nutrients move across the leaf surface remain unclear (Fernández et al., 2017; Li et al., 2017). Of particular interest, in the present study we investigated the role of trichomes in foliar absorption of Zn in soybean and tomato. Two types of trichomes were found on the leaf surface of soybean and seven types for tomato. Using synchrotron-based μ-XRF for in situ analyses of hydrated leaves of soybean and tomato after foliar Zn (ZnSO4) application, it was found for soybean that ca. 29 % of Type V trichomes accumulated Zn at their base, but in tomato, no trichomes were observed to accumulate Zn. Next, given that Zn was observed to accumulate in some trichomes on leaves of soybean, we examined this plant species in more detail using NILs differing up to 10-fold in trichome density as well as by using in situ analyses of Zn distribution in leaf cross sections with μ-XRF. However, using these approaches, we found that trichomes were not a primary pathway for the foliar absorption of Zn in soybean either.

Role of trichomes in absorption of foliar-applied Zn First, we examined the distribution of Zn in leaves in order to determine if Zn accumulates in trichomes upon its foliar application. Using μ-XRF for the in situ analysis of Zn distribution within hydrated leaves we found that, for both soybean and tomato, Zn moved rapidly across the leaf surface. In fact, Zn was observed in the mesophyll within only 15 min from application (Fig. 3 and Supplementary Fig. S2). It was noted that Zn accumulated first within the veins, with Zn concentrations subsequently increasing in the interveinal tissues, especially in the apoplast of interveinal tissues (Fig. 4). Of particular interest, it was also noted that Zn accumulated at the base of ca. 29 % non-glandular trichomes in soybean especially before 3 h (Fig. 4 and Supplementary Fig. S3). This accumulation in trichome bases possibly occurred because it has been reported that the base of non-glandular trichomes may be less waxy and have a higher wettability compared to the cuticle (Brewer et al., 1991; Grammatikopoulos and Manetas, 1994; Fernández et al., 2014). Our finding is similar to the results of Schlegel and Schönherr (2002) who used Ag to visualise foliar uptake through the precipitation of AgCl in broad bean (Vicia faba L.) and maize (Zea mays L.). However, analysis of the cross

115 sections by µ-XRF showed that the Zn concentration was not higher in the cells surrounding the base of the trichomes (Fig. 5). Interestingly, we did not observe the strong binding of Zn to the epidermal cells (i.e. Zn was moved into the underlying mesophyll cells) (Fig. 5) which has been proposed as a reason to account for the limited translocation of some foliar absorbed nutrients, including Ca, Mn, Fe, and Zn (Zhang and Brown, 1999; Du et al., 2015; Li et al., 2017). Instead, we found that the foliar absorbed Zn had moved further into the leaf tissues (into the mesophyll cells), even after only 15 min (Fig. 5).

Next, to further quantify the overall importance of trichomes in foliar Zn absorption, we compared the absorption of foliar-applied Zn in four soybean NILs differing up to 10-fold in -1 their trichome density (Fig. 6). When Zn was supplied as a 1000 mg Zn L ZnSO4, it was found that after 1 h of exposure, although Zn tissue concentrations had increased relative to the control, there were no significant differences among the four soybean NILs despite the differing trichome densities (Fig. 7). Furthermore, after exposure for 6 h, although significant differences were found among the NILs, these differences were not related to leaf trichome densities (Figs. 6 and 7). Thus, although Zn was found to accumulate at the base of some trichomes upon the foliar-application of Zn (Fig. 4), trichomes were not the primary organ for the movement of Zn across the leaf surface (Figs. 6 and 7).

Our observation that, in soybean and tomato, trichomes do not appear to be the primary pathway for the movement of Zn across the leaf surface (Figs. 7 and 8) is consistent with Burrows et al. (2013) who demonstrated that trichomes on the leaves of silverleaf nightshade (Solanum elaeagnifolium; Solanaceae) did not facilitate the uptake of a fluorescent tracer. However, in contrast to these observations, it is known that peltate trichomes in the highly drought-resistant Bromeliaceae are associated with foliar absorption of water, Ca, and Zn (Ohrui et al., 2007; Winkler and Zotz, 2010). Similarly, trichomes in drought-tolerant cactus are involved in water collection from fog (Ju et al., 2012), and trichomes in drought-stressed Croton can facilitate water penetration into leaves (Vitarelli et al., 2016). On the other hand, it has been reported that trichomes of temperate woodland ferns under of high air humidity may have a role in foliar water uptake (Schwerbrock and Leuschner, 2017). Therefore, it seems that the role of trichomes differs widely, depending upon the plant species and the climatic and ecological conditions (Werker, 2000). Indeed, although it was been reported that the peltate trichomes of some Bromeliaceae species under drought conditions can absorb water and nutrients which is helpful to overcome the drought conditions (Ohrui et al., 2007; 116

Winkler and Zotz, 2010), while the stellate or peltate trichomes of some terrestrial Bromeliaceae species are water repellant and are hydrophobic which could potentially obstruct pathogens and aid in self-cleaning (Pierce et al., 2001). However, much remains unknown regarding the physico-chemical and ecological significance of trichomes (Bickford, 2016). Thus, although we have found that trichomes are not important for foliar absorption of Zn in soybean and tomato, it is possible that trichomes could play a role in foliar absorption in other plant species. In addition, temperature, humidity and the point of deliquescence of the nutrient compound should also be considered when assessing foliar absorption (Kerstiens, 2006; Fernández et al., 2017).

Importance of other properties for absorption of foliar-applied Zn Given that Zn absorption was not related to trichome density in the four soybean NILs, we also examined stomatal density. However, stomatal density did not differ among the four NILs (Fig. 6) and hence changes in foliar absorption of Zn (Fig. 8) were not due to changes in stomatal density. Next, we considered whether the differences in absorption of Zn among the four NILs could be due to differences in contact between the leaves and the droplets, with trichomes known to alter leaf surface wettability (Brewer et al., 1991; Fernández et al., 2014). However, we found that contact angles did not differ among the four NILs (Supplementary Figs. S4 and S5). This is consistent with Brewer et al. (1991) who found that leaf water repellence is related to trichome density, particularly when the leaf trichome density is > 25 mm-2 (in constrast, the trichome density of the NILs in the present study was 1 – 11 mm-2). Thus, differences in Zn absorption in the soybean NILs could not be attributed to differences in wetting angle in the present study. Rather, it is possible that the differences in foliar Zn absorption observed among the NILs could be due to different cuticle properties, for example, although this was not examined in the present study. Also it must be noted that the RH during the foliar application period of the present study was very high (98 %). This perhaps increased the cuticular permeation in the present study, as it has previously been suggested that the cuticular permeation increases with RH, particularly when RH > 90 % (Schonherr, 2001; Fernández and Eichert, 2009; Fernández et al., 2017).

Pattern of accumulation of Zn in leaf tissues It was observed that foliar-applied Zn accumulated more rapidly in the veins than in the interveinal tissues for both plant species (Fig. 4). This comparatively rapid accumulation in the vein is consistent with previous studies, including Du et al. (2015) who reported that 117 foliar-applied Zn accumulated to the highest concentrations in veins of tomato and citrus (Citrus reticulatus). Similarly, Li et al. (2017) found the concentrations of foliar absorbed Fe, Mn, and Zn in the veins were 2- to 4-fold higher than in the corresponding interveinal tissues in both sunflower (Helianthus annuus) and tomato. However, the reason for this rapid accumulation in the veins remains unclear. Given that leaf wettability is an important plant- related factor influencing the absorption of foliar-applied nutrients (Fernández and Brown, 2013; Fernández et al., 2013), we measured the contact angle of a Zn droplet overlaying either a vein or the interveinal tissues, it was found that the contact angle of a droplet above the vein was 2-fold smaller than for the interveinal tissues (Supplementary Fig. S6). Therefore, we suggest that the initial rapid accumulation of Zn in the vein (Fig. 4) potentially occurs due to the higher wettability of the vein compared to the interveinal tissues, thereby resulting in better adhesion and contact.

Effect of the form of Zn on foliar absorption It has been proposed that nano-based foliar fertilizers may be more effective than traditional foliar fertilizers. For example, it is possible to alter the surface functionalisation of nano- based fertilizer products, such as for preparing fertilizers with sustained release rates (De Rosa et al., 2010). It has previously been shown that nano-fertilizers can be absorbed by both the leaves and roots (Slomberg and Schoenfisch, 2012; Hussain et al., 2013), but it is unclear as to whether the nanoparticles themselves are taken up, or whether it is their dissolution products (Wang et al., 2016; Wang et al., 2017). In the present study, we examined nano- ZnO (ca. 30 nm diameter) and homologous bulk-ZnO (ca. 200-400 nm diameter)

(Supplementary Fig. S1). For an additional control, we also examined ZnSO4 at a Zn concentration of 3 mg Zn L-1, with this corresponding to the same soluble Zn concentration measured for the nano- and bulk-ZnO suspensions. We exposed the leaves of the four soybean NILs to the Zn-containing compounds for 6 h, and found that absorption upon -1 exposure to 1000 mg Zn L ZnSO4 was on average ca. 10-fold higher than for nano- and bulk-ZnO suspensions. Moreover, there were no marked differences in foliar Zn absorption between the nano- and bulk-ZnO particles which differed 10-fold in their particle size. The markedly lower absorption upon exposure to the nano- and bulk-ZnO suspensions compared -1 to 1000 mg Zn L ZnSO4, together with the similar absorption for nano- and bulk-ZnO, suggests that the movement of Zn across the leaf surface mainly occurs as soluble Zn, rather than as solid ZnO. This is perhaps surprising, given that the nano-ZnO had an average particle diameter of 30 nm, which is smaller than typical stomatal openings (ca. 2 × 9 µm) 118

(Supplementary Fig. S7). It appears that ZnO was not greatly absorbed through the stomata, although the role of stomata in foliar absorption remains unclear (Fernández and Brown, 2013). Regardless, it is possible that the slower absorption of Zn from the nano- and bulk- ZnO suspensions is actually advantageous, not only in avoiding the ‘burning’ that can occur with soluble forms (Drissi et al., 2015), but also in providing a prolonged source of Zn (i.e. a slow-release fertilizer) if these ZnO particles can be retained on the leaf surface. This should form the focus of future research efforts in this regard.

Interestingly, the extent of absorption upon exposure to 1000 mg Zn L-1 nano- and bulk-ZnO -1 suspensions was on average ca. 6-fold higher than when exposed to 3 mg Zn L ZnSO4. This difference occurred despite the concentration of soluble Zn (3 mg Zn L-1) being the same across the three treatments. Given that absorption upon exposure to nano- and bulk-ZnO -1 suspensions was higher than for 3 mg Zn L ZnSO4, it is possible that the foliar-absorption of soluble Zn from the ZnO suspensions on the leaf surface drives the continual dissolution of -1 the ZnO, and increased Zn absorption compared to the 3 mg Zn L ZnSO4 treatment.

Finally, the point of deliquescence of different compounds can influence their rate of foliar absorption (Fernández and Eichert, 2009; Fernández et al., 2017). However, this is particularly important in longer term experiments, such as those where droplets of the foliar- applied compound have had sufficient time to potentially dry out. In the present study, however, droplets were applied for ≤ 6 h and remained as liquids during this period. Given that the absorption of Zn differed by 10-fold among the three Zn compounds, this cannot be attributed to differences in the point of deliquescence. Rather, this was due to differences in the soluble Zn concentration.

Conclusions Despite the importance of foliar fertilization for improving crop nutrition, the mechanisms by which foliar-applied nutrients move across the leaf surface remain unclear, including the potential role of trichomes. Using synchrotron-based μ-XRF and NILs of soybean differing up to 10-fold in trichome density, we found that trichomes are not part of the primary pathway by which foliar-applied Zn moves across the leaf surface in soybean and tomato. However, it is important to extend this work to other plant species which are known to differ in their leaf properties and trichome types. In addition, we also compared soluble Zn as

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ZnSO4 with sparingly soluble Zn as nano- and bulk- ZnO, finding that Zn was mainly absorbed as soluble Zn rather than as ZnO (whether supplied in nano or bulk forms), with Zn absorption upon exposure to ZnSO4 ca. 10-fold higher than for either nano- or bulk-ZnO after 6 h of application. However, the continued absorption of soluble Zn from suspensions of nano- and bulk-ZnO appears to result in their gradual dissolution, suggesting that they may potentially be useful as a slow release fertilizer provided that they can be retained on the leaf surface, with further studies required in this regard.

Acknowledgements The authors thank Professor Randall Nelson (University of Illinois, USA) for providing the seeds of the four soybean NILs, and Kerry Vinall and Brett Ferguson (The University of Queensland) for assistance with growing the plants. The authors also thank facilities, and the scientific and technical assistance, of both the Queensland Brain Institute and Australian Microscopy & Microanalysis Research Facility at the Centre for Microscopy and Microanalysis, The University of Queensland. Parts of the research were undertaken on the XFM beamline at the Australian Synchrotron, part of Australian Nuclear Science and Technology Organisation (ANSTO). This work was supported by Sonic Essentials, as well as by the Australian Research Council (ARC) through the Linkage Projects funding scheme (LP130100741). Support was also provided to Peter Kopittke by the ARC Future Fellowship funding scheme (FT120100277) and to Cui Li through the China Scholarship Council and The University of Queensland International Scholarship award.

Supplementary Data Fig. S1 Scanning electron micrographs showing particles of the nano- and bulk-ZnO. Fig. S2 Tomato µ-XRF survey scans. Fig. S3 Tri-color image (detailed scan; red is Zn, K is green, and Ca is blue) of a soybean leaf scanned from underneath where a ZnSO4 droplet had been applied for 1 h. Fig. S4 Comparison of the contact angles of the three Zn fertilizers with the leaves of the four soybean near-isogenic lines. -1 Fig. S5 Light micrographs showing contact angles of 1000 mg Zn L ZnSO4.7H2O (5 µL droplets) with the four soybean near-isogenic lines.

Fig. S6 Comparison of contact of foliar applied ZnSO4.7H2O droplets with the vein and the interveinal tissue of a soybean leaf.

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Fig. S7 Scanning electron micrograph showing the size of the opening stomata of a soybean leaf. Table S1 Results from dynamic light scattering of the nano- and bulk-ZnO compounds.

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Fig. 1 Trichomes on the soybean leaf surface, with ‘V’ and ‘IX’ indicating two types of trichomes (Type V and Type IX). Leaves were analysed using both light microscopy (B, E, and F) and scanning electron microscopy (A, C, and D). (A-D) Trichomes on the adaxial leaf surface, with the Type V trichome in (D) broken and showing a cross section. (E-F) Leaf cross sections stained with toluidine blue.

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Fig. 2 Trichomes on the tomato leaf surface, with ‘I’ – ‘VII’ indicating seven types of trichomes. Leaves were analysed using both light microscopy (E-H) and scanning electron microscopy (A-D). (A-D) Trichomes on the adaxial leaf surface. (E-H) Leaf cross sections stained with toluidine blue.

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Fig. 3 (A-E) Light micrographs showing the droplets (ZnSO4) on the soybean leaf before being removed and rinsed. (F-J) Results from the µ-XRF survey scans showing the distribution of Zn. The images in (F-J) have the same color scale, with brighter colors corresponding to higher Zn concentrations (mg/kg). Droplets were left on the leaf surface for 15 min, 30 min, 1 h, 3 h, or 6 h. The scale bar in (A) applies to (A-E), and the scale bar in (F) applies to (F-J).

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Fig. 4 Detailed µ-XRF scans showing the distribution of Zn in hydrated (fresh) leaves of soybean (A-E) and tomato leaves (F-J). All of the image areas were obtained from the area underlying a Zn droplet. The Zn droplets were applied to the leaf surface for 15 min, 30 min, 1 h, 3 h, or 6 h, all the images have the same color scale, with brighter colors corresponding to higher Zn concentrations (mg/kg). The scale bar in (A) applies to (A-E), the scale bar in (F) applies to (F-J).

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Fig. 5 (A) A light micrograph of soybean leaf cross section which had Zn applied for 1 h before being scanned by µ-XRF; (B) µ-XRF scans showing the distribution of Zn in soybean leaves (150 µm leaf cross sections). The cross sections were obtained from tissues underlying Zn droplets after 0 min (control), 15 min, 1 h, and 6 h of Zn application. In each cross section, the left side is the adaxial side where the Zn had been applied. Brighter colors corresponded to higher Zn concentrations. ‘V’ indicates the Type V trichome on soybean leaf surface. Note that the color is not comparable among the four samples, and Zn color scales have been provided for each cross section.

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Fig. 6 Densities of trichomes (A) and stomata (B) for leaves of four soybean near-isogenic lines (NILs): P1 (L62-1385), Ps (L63-2999), Pd1 (L62-1686), and CS (L58-231). In (A), ‘IX’ and ‘V’ refer to two types of trichomes. In A and B, values are mean ± standard deviation (n = 6). Significant differences (P < 0.05) are indicated by different lowercase letters. (C) Scanning electron micrographs showing the trichomes in the four NILs. The scale bar for P1 applies to all four images in (C).

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Fig. 7 Comparison of the foliar absorption of Zn after exposure for 1 h to 1000 mg Zn L-1

ZnSO4. Four near isogenic lines (NILs) of soybean were compared that differed in trichome density, being P1 (L62-1385), Ps (L63-2999), Pd1 (L62-1686), and CS (L58-231). Values are mean ± standard deviation (n = 6). Different lowercase letters show significant (P < 0.05) differences among the four NILs. Foliar absorption of 1000 mg Zn L-1 nano- and bulk- ZnO, -1 and 3 mg Zn L ZnSO4 were not presented due to the absorption was too low that cannot detected by ICP-MS.

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Fig. 8 Comparison of the foliar absorption of Zn after exposure for 6 h to 1000 mg Zn L-1 -1 -1 -1 ZnSO4 (A), 1000 mg Zn L nano-ZnO (C), 1000 mg Zn L bulk-ZnO (D), and 3 mg Zn L

ZnSO4 (B). Four near isogenic lines (NILs) of soybean were compared that differed in trichome density, being P1 (L62-1385), Ps (L63-2999), Pd1 (L62-1686), and CS (L58-231). Values are mean ± SD (n = 6). Different lowercase letters show significant (P < 0.05) differences among the four NILs.

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SUPPLEMENTARY DATA Role of leaf trichomes in absorption of Zn from three types of foliar- applied fertilizers in soybean and tomato

Cui Li1, Peng Wang1, 2, *, Enzo Lombi3, Miaomiao Cheng4, Caixian Tang4, Daryl L. Howard5, Neal W. Menzies1, Peter M. Kopittke1

Author for correspondence: Peng Wang Tel: +86 25 84399055, Fax: 86 25 84399055 Email: [email protected]

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Fig. S1 Scanning electron micrographs showing particles of the nano-ZnO (A) and the bulk- ZnO (B).

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Fig. S2 (A-E) Light micrographs showing the Zn droplets (ZnSO4) on tomato leaves before being removed and rinsed. (F-J) Results from the µ-XRF survey scans showing the distribution of Zn. The images in (F-J) have the same color scale, with brighter colors corresponding to higher Zn concentrations (mg/kg). Droplets were left on the leaf surface for 15 min, 30 min, 1 h, 3 h, or 6 h. The scale bar in (A) applies to (A-E), and the scale bar in (F) applies to (F-J).

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Fig. S3 Tri-color image (detailed scan; red is Zn, K is green, and Ca is blue) of a soybean leaf scanned from underneath where a ZnSO4 droplet had been applied for 1 h. Many Type V trichomes can be seen (ca. 200-400 µm long), with some containing elevated concentrations of Zn.

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-1 -1 Fig. S4 Contact angles for 5 µL droplets of 1000 mg Zn L ZnSO4 (a), 1000 mg Zn L nano- ZnO (b), and 1000 mg Zn L-1 bulk-ZnO (c) droplets (all solutions with 0.05 % Tween 20). The droplets were applied to leaves of four near-isogenic lines (NILs) of soybean. Measurments were taken 0, 15, 30, and 60 min after applying the droplets (n = 6). In all instances, for the two-way analysis of variance there was no significant interaction (NIL and time) or any significant differences among NILs. Therefore, the only LSD values (5 %) shown are for time, with contact angles generally decreasing over time.

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Fig. S5 Light micrographs from which the contact angles were calculated for four near isogenic lines (NILs) of soybean. Droplets (5 µL) were applied to interveinal leaf tissues -1 using 1000 mg Zn L ZnSO4, with images captured after 0, 15, 30, and 60 min. All droplets -1 contained 0.05 % Tween 20. Only data for 1000 mg Zn L ZnSO4 are shown here, with the images for nano-ZnO and bulk-ZnO being similar.

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-1 Fig. S6 Comparison of the contact of droplets of 1000 mg Zn L ZnSO4 (5 µL droplets with 0.05 % Tween 20) with the vein and the interveinal tissue of a soybean leaf. The image was captured immediately after the droplets had been applied.

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Fig. S7 Scanning electron micrograph showing opening stomatal size of P1 L62-1385 (one of the four soybean near-isogenic lines, with the other three having similar stomatal size).

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Table S1 Results of analyses using dynamic light scattering (DLS) for the nano- and bulk- ZnO. The numbers in brackets are the relative abundance of particles of each size (%). Mean diameter (nm) Fit Product Residual Volume Number Error Peak 1 Peak 2 Peak 1 Peak 2 Bulk-ZnO 2.13 5.52 154 555 143 506 (15.9) (84.1) (89.7) (10.3) Nano-ZnO 2.59 0.00 59.4 173 55.9 161 (72.1) (27.9) (98.4) (1.60)

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Chapter 5 Absorption of foliar-applied Zn in sunflower (Helianthus annuus): Importance of the cuticle, stomata, and trichomes

5.1 Introduction

Evidence collected and presented in the previous two Chapters indicated that the cuticle plays an important role for the foliar absorption of Zn, while the trichomes appear to not be of importance for this process in soybean and tomato. However, as discussed in Chapter 4, trichomes are highly diverse, and differ markedly among plant species and the ecological and environmental conditions. In addition, whether stomata play an important role in foliar Zn absorption also remains unclear. In this study, sunflower was investigated as it is known that this plant accumulates metals in its trichomes when they are supplied in the rooting environment. The distribution of the foliar absorbed Zn was examined in both intact leaves and cross sections using µ-XRF in order to examine the potential role of trichomes. Next, the Zn distribution within the leaf cross sections was examined using NanoSIMS, with this providing a subcellular analysis of Zn distribution. Overall this study aimed to explore the importance of cuticle, trichomes, and stomata in foliar Zn absorption in sunflower. In addition, the absorption of nano-ZnO and ZnSO4 was compared using µ-XRF, with the chemical speciation of the absorbed Zn examined using XAS.

5.2 Paper 4

Li C, Wang P, Van der Ent A, Cheng M, Jiang H, Read TL, Lombi E, Tang C, de Jonge M, Menzies NW, Kopittke PM (2018) Absorption of foliar-applied Zn in sunflower (Helianthus annuus): Importance of the cuticle, stomata, and trichomes. Annals of Botany. DOI: 10.1093/aob/mcy135.

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ORIGINAL ARTICLE Absorption of foliar-applied Zn in sunflower (Helianthus annuus): Importance of the cuticle, stomata, and trichomes

Running Title: The pathways of foliar Zn absorption in sunflower

Cui Li1*, Peng Wang2,3, Antony van der Ent4,5, Miaomiao Cheng6, Haibo Jiang7, Thea Lund Read8, Enzo Lombi8, Caixian Tang6, Martin D. de Jonge9, Neal W. Menzies1, Peter M. Kopittke1

1The University of Queensland, School of Agriculture and Food Sciences, St Lucia, Queensland, 4072, Australia; 2Nanjing Agricultural University, College of Resources and Environmental Sciences, Nanjing, 210095, China; 3The University of Queensland, Centre for Soil and Environmental Research, School of Agriculture and Food Sciences, St Lucia, Queensland, 4072, Australia; 4Centre for Mined Land Rehabilitation, Sustainable Minerals Institute, The University of Queensland, Australia, 5Laboratoire Sols et Environnement, Université de Lorraine, France; 6La Trobe University, Centre for AgriBioscience, Bundoora, Victoria, 3086, Australia; 7University of Western Australia, Centre for Microscopy, Characterization and Analysis, Crawley, WA 6009, Australia; 8University of South Australia, Future Industries Institute, Mawson Lakes, South Australia, 5095, Australia; 9ANSTO, Australian Synchrotron, Clayton, Victoria, 3168, Australia

* For correspondence. Email: [email protected]; Phone: +61 424662660

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Abstract

 Background and Aims The pathways whereby foliar-applied nutrients move across the leaf surface remain unclear. The aim of the present study was to examine the pathways whereby foliar-applied Zn moves across the sunflower leaf surface, considering the potential importance of the cuticle, stomata, and trichomes.

 Methods Using synchrotron-based X-ray florescence microscopy and nanoscale secondary ion mass spectrometry (NanoSIMS), the absorption of foliar-applied

ZnSO4 and nano-ZnO were studied in sunflower (Helianthus annuus). The speciation of Zn was also examined using synchrotron-based X-ray absorption spectroscopy.

 Key Results It was found that the non-glandular trichomes (NGTs) were particularly important for foliar Zn absorption, with Zn preferentially accumulating within trichomes in ≤ 15 min. The cuticle was also found to have a role, with Zn appearing to move across the cuticle before accumulating in the walls of the epidermal cells. After 6 h, the total Zn that accumulated in the NGTs was ca. 1.9 times higher than in the cuticular tissues. No marked accumulation of Zn was found within the stomatal cavity, likely indicating a limited contribution of the stomatal pathway. Once absorbed, the Zn accumulated in the walls of the epidermal and the vascular cells, and trichome bases of both leaf sides, with the bundle sheath extensions that connected to the trichomes seemingly facilitating this translocation. Finally, the absorption of nano-

ZnO was substantially lower than for ZnSO4, with Zn likely moving across the leaf surface as soluble Zn rather than nanoparticles.

 Conclusions In sunflower, both the trichomes and cuticle appear to be important for foliar Zn absorption.

Keywords: bundle sheath extension; cuticle; foliar absorption; stomata; trichomes; Zn foliar fertiliser; sunflower

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INTRODUCTION It has been known since the 19th Century that nutrients can be absorbed by plant leaves (Mayer, 1874). Since this time, foliar fertilisation has become increasingly widespread in its usage (Alexander, 1986, Fernández et al., 2009, Fernández et al., 2013). Although the movement of nutrients into root systems is well understood, comparatively little is known about nutrient absorption by above-ground tissues. Indeed, the cuticle, stomata, and trichomes have all been proposed as potential pathways for the movement of foliar-applied nutrients, but their relative contributions to overall foliar absorption are uncertain (Fernández et al., 2013), with this being the focus of the present study.

For the cuticle, it is proposed that there are two different cuticular penetration pathways: (i) the lipophilic pathway for non-ionic, apolar, and lipophilic compounds (e.g. pesticides and herbicides); and (ii) the hydrophilic pathway for ionic, polar, hydrophilic substances (e.g. mineral nutrients) (Schönherr, 2006, Stagnari, 2007, Schreiber and Schönherr, 2009). In the present study, the hydrophilic pathway is of particular interest, although comparatively little is known regarding this pathway (Schönherr, 1976, Schönherr, 2006). Fernández et al. (2017) proposed the ‘dynamic aqueous continuum’ model which suggested that the cuticle has highly tortuous, relative humidity dependent, dynamic sorption and desorption of water connections associated with hydrophilic domains existing between the leaf surface and interior tissues. However, due to the complex chemical composition and structure of the cuticle, the mechanisms and importance of the cuticle in the foliar absorption of mineral nutrients remains unclear (Fernández et al., 2017).

Turrell (1947) reported that water could not penetrate the stomata of citrus (Citrus reticulatus) while oil could. Subsequently, Schönherr and Bokovac (1972) suggested that spontaneous infiltration of stomata cannot occur unless pressure is applied or surfactants are added in order to reduce surface tension. Interestingly, Eichert et al. (1998) used leaf epidermal strips found that the penetration of uranine through stomata happened without the addition of either surfactants or pressure. Furthermore, Eichert et al. (2008) observed that hydrophilic polystyrene particles with a diameter of 43 nm suspended in water can enter leaves through stomata, although only < 10 % of the stomata beneath the droplet were penetrated. In addition, Burkhardt et al., (2012) found that the hygroscopic leaf surface particles potentially promote the stomatal penetration. Thus, while it has been shown that the movement of foliar-applied nutrients through the stomata is possible, both the mechanisms by 146 which this occurs and the overall contribution of stomata to foliar nutrient absorption are unclear.

The role of trichomes in foliar nutrients absorption has received comparatively little attention (Fernández et al., 2013). Schlegel et al. (2001) used Ag to visualize foliar absorption through the precipitation of AgCl, finding that there was a substantial accumulation of Ag in trichomes of broad bean (Vicia faba) and maize (Zea mays). Absorption of water from dew (Pina et al., 2016) or fog (Ju et al., 2012) through trichomes has been reported in some species as a drought resistance mechanism, thereby indicating the possibility that absorption of foliar-applied nutrients may potentially occur through trichomes in some plants. However, trichomes are markedly diverse in their shape, structure, and function (Huchelmann et al., 2017), and hence it is possible that the potential contribution of trichomes to absorption of foliar-applied nutrients vary amongst plant species. Of particular interest in the present study is sunflower (Helianthus annuus) which has been reported to accumulate excess Mn in the non-glandular trichomes (NGTs) when the Mn is supplied in the rooting medium (Blamey et al., 2015, Blamey et al., 1986).

The aim of the present study was to examine the pathways whereby foliar-applied Zn moves across the sunflower leaf surface, considering the potential importance of the cuticle, stomata, and trichomes. In addition, given the increasing interest in using nanotechnology in plant production systems (De Rosa et al., 2010, Wang et al., 2016), we compared absorption of a traditional Zn foliar fertiliser (ZnSO4) with a nano foliar fertiliser (nano-ZnO). To do this, we first examined leaf structure and surface morphology using scanning electron microscopy (SEM) and light microscopy. Next, we examined the lateral distribution of foliar absorbed Zn in situ within hydrated, intact leaves using synchrotron-based X-ray fluorescence microscopy (µ-XRF). Then, we examined the distribution of the foliar absorbed Zn within leaf cross sections at tissue and subcellular tissue levels using µ-XRF and nanoscale secondary ion mass spectrometry (NanoSIMS), respectively. Finally, the speciation of foliar absorbed Zn within the leaves was examined using synchrotron-based X-ray absorption spectroscopy (XAS). It was hypothesized that the cuticle, stomata, and trichomes would all contribute as pathways for the movement of foliar-applied nutrients across the leaf surface. It was hoped that this study would help to identify the mechanisms of foliar nutrient absorption, and hence provide important information for improving the efficacy of foliar fertilisers.

147

MATERIALS AND METHODS Plant growth and examination of leaf morphology Sunflower plants were grown in nutrient solution at The University of Queensland (St Lucia, Australia) as described by Li et al. (2017). Briefly, seeds (cv. Hyoleic) were germinated for 3 d using paper towel moistened with tap water. Seedlings were transplanted to 11 L black containers, with one plant placed in each of the four holes in the lids. The containers were - + filled with nutrient solution which contained (µM): 910 N (94 % NO3 and 6 % NH4 ), 20 P, 475 K, 1126 Ca, 227 Mg, 1251 Cl, 556 S, 25 Fe(III)EDTA, 3 B, 0.5 Mn, 0.5 Zn, 0.2 Cu and 0.01 Mo (Blamey et al., 2015). The nutrient solution was continuously aerated, the room temperature maintained at 25 °C, and light was provided using high-pressure sodium lamps (photon flux density of 1500 μmol m–2 s–1) for 12 h d-1. Nutrient solutions were changed after the first 7 d then every 4 d thereafter. After 10 d, 5 mL of 44 mM KH2PO4 was added to each 11 L container every other day to replenish P. Nutrient solution pH was unadjusted and was ca. 5.5.

After 16 d of growth, the youngest fully-expanded leaves (YFELs) were used to examine leaf morphology and structure using SEM following processing as described by Li et al. (2017). The adaxial leaf surface was examined using an SEM (JEOL NEOSCOPE, Japan) at 10 kV accelerating voltage. For examining leaf structure, the fresh YFELs were cut into pieces ca. 1 mm × 4 mm and fixed, dehydrated, and embedded in Epon resin as described by Li et al. (2017). Leaf sections (1 µm thick) were cut using an ultramicrotome (LEICA EM UC6), stained by Toluidine blue (0.5 % in 1 % borax) and observed using light microscopy (O'Brien et al., 1964). Leaf structure was also examined using fresh leaf tissues. For this, fresh leaf segments were embedded in a 4 % agar solution at 30 °C. Once the agar solidified, the block was glued to the sample holder of a vibratome (Leica VT 1000s) using super glue, then 80 µm leaf sections prepared without any buffering solution. Sections were immediately imaged using an Aperio XT Slide Scanner without staining.

Laterally-resolved examination of Zn distribution in leaf tissues using µ-XRF The 16 d old sunflower plants were transported to the Australian Synchrotron (Clayton, Australia) where they continued to be grown under the same conditions as at The University of Queensland. Two types of Zn fertilisers were prepared at 1000 mg Zn L-1 (15.4 mM), being ZnSO4.7H2O (pH 5.2) and nano-ZnO (ca. 30 nm diameter, pH 7.1) [see Li et al. (2018) for scanning electron micrograph and the dynamic light scattering results of the nano-ZnO]. 148

To prepare 50 ml of 1000 mg Zn L-1 nano-ZnO, nano-ZnO was placed into a 50 ml Falcon tube, 30 ml water was first added and mixed for 15 min using ultrasonic processor, then water was added to 50 ml. The suspension was shaken and mixed thoroughly before use. The solutions of both types of Zn compounds contained 0.05 % Tween 20 (Sigma-Aldrich) as a surfactant.

Firstly, we did a time series experiment to investigate the Zn absorption process using intact, hydrated leaves for which only ZnSO4 was used. Specifically, we investigated six foliar application times, being 0 (control), 0.25, 0.5, 1, 3, and 6 h. Two 5 µL droplets of the ZnSO4 solution were applied to the adaxial surface of the YFELs. The whole leaf (still attached to the plant) was then placed inside a Petri dish (the dish had a hole in the side wall to allow the petiole to pass through) with moistened filter paper in the bottom of the dish to maintain humidity. The temperature within the Petri dish was 30 °C and the relative humidity increased rapidly to 98 %. The growth lamps remained switched on during the incubation period and the droplets remained as a liquid (i.e. did not dry out) during the experimental period. After incubation for the appropriate length of time, the leaves were excised and rinsed using 2 % HNO3, 3 % ethanol, and deionized water (Vu et al., 2013). For each specimen, the rinsed leaf was blotted dry and mounted on a sample holder between two layers of Ultralene film (4 µm thick). Samples were then immediately analysed using µ-XRF at the XFM beamline of the Australian Synchrotron. This beamline has been described previously (Paterson et al., 2011), including for the in situ analysis of hydrated plant tissues (Kopittke et al., 2011). For these scans, a 384-element Maia detector system in a backscatter position was used to collect the X-ray fluorescence emitted by the specimen. We used an incident energy of 12.9 keV with a total photon flux of ca. 1.5 × 109 photon s-1. Initially, a ‘survey scan’ was used to obtain a comparatively rapid image of the leaf from which the area for a detailed scan could be identified. For this initial survey scan, the transit time per virtual pixel was 7.5 ms and the horizontal stage velocity was 4 mm s-1, resulting in a virtual pixel size of 30 µm. The scan area was often ca. 10 × 10 mm, with these scans taking ca. 15 min. After the survey scan, an area entirely underneath where the Zn droplet had been applied was selected, and a detailed scan was conducted. For the detailed scan, the transit time was 1 ms and the horizontal stage velocity was 1 mm s-1, resulting in a virtual pixel size of 1 µm. The detailed scans were often ca. 4 × 5 mm in size and took ca. 75 min to complete. Next, we compared the absorption of ZnSO4 and nano-ZnO. For this experiment, 5 µL droplets of ZnSO4 and

149 nano-ZnO were applied next to each other on the adaxial surface of the same YFEL for either 3 h or 3 d. The leaf was rinsed as described above before being analysed using µ-XRF.

For the next experiment, we examined the distribution of Zn within cross sections of leaf tissues. All tissues examined within the cross sections were obtained from tissues that had been entirely underneath the Zn-containing droplets. This was done by applying a larger droplet (40 µL) and only excising those tissues that had been underneath these droplets. A time series was prepared, being 0 (control), 0.25, 0.5, 1, 3, and 6 h from the time of application. Cross sections of each sample were then prepared from tissues located beneath where the Zn droplets had been applied. Two types of cross sections were prepared. Firstly, freeze dried specimens were prepared in advance at The University of Queensland. For these samples, leaf tissues were excised with a razor blade and leaf segments sectioned (150 µm thick) using a vibratome as outlined earlier. The sections were then carefully sealed in Ultralene film held onto XRF sample cups (SC-8047, Premier Lab Supply) and freeze dried after small holes had been punctured in the film. Secondly, we prepared fresh (hydrated) cross sections at the Australian Synchrotron in order to allow comparison with the freeze dried samples. Specifically, using plants growing at the Australian Synchrotron (the plants were grown under the same conditions as at The University of Queensland), the tissues underneath where the droplet had been applied were excised and placed in 4 % agar solution, transported to Monash University (Clayton, Australia) whilst being held on ice (the transfer from the Australian Synchrotron to Monash University took ca. 10 min), and sectioned (150 µm) using a vibratome as outlined earlier. These hydrated (fresh) sections were then sealed between two layers of Ultralene film held using XRF sample cups and immediately transported back to Australian Synchrotron (on ice) for examination.

The leaf cross sections were then examined using µ-XRF. We first examined samples that had been incubated for either 3 or 6 h, comparing the fresh and freeze dried samples. It was found that freeze drying did not alter the distribution of Zn in the leaf tissues (see below). Thus, hereafter, unless otherwise noted, all cross sections that were examined were freeze dried. To analyse these samples using µ-XRF, the sections were held between two layers of Ultralene film stretched over a Perspex frame magnetically mounted on the motion stage under a cryo-stream (operated at -100 °C, while the actual temperature on the sample is estimated below -50 °C). A beam of 15.8 keV X-rays was focussed to a spot of around 2 µm diameter, and a silicon drift-diode detector (Vortex) located in the 90° geometry was used to 150 collect X-ray fluorescence emitted by the specimen. The photon flux was ca. 2 × 109 photon s-1. Firstly, a quick ‘survey scan’ was conducted, with the transit time being 50 ms and the horizontal stage velocity being 1 mm s-1, resulting in a virtual pixel size of 50 µm. The area scanned was generally ca. 5 × 0.15 mm, taking ca. 3 min to complete. Next, a ‘detailed scan’ was conducted, with the transit time being 3.33 ms and the horizontal stage velocity being 0.6 mm s-1, yielding a virtual pixel size of 2 µm pixel. The detailed scans took ca. 100 min. In order to compare the foliar absorption of ZnSO4 and nano-ZnO, a 6 h nano-ZnO exposure cross section was also examined using µ-XRF. All µ-XRF data was analysed using the CSIRO Dynamic Analysis method in GeoPIXE (http://www.nmp.csiro.au/dynamic.html) (Ryan et al., 1993, Ryan, 2000).

To obtain measurements of bulk Zn concentrations, bulk leaves (four replicates) were analysed by inductively coupled plasma mass spectrometry (ICP-MS) to compare foliar absorption of ZnSO4 and nano-ZnO. A half-leaf loading method was used (Vu et al., 2013), with half of each leaf receiving either 60 droplets (5 μL per droplet) of 1000 mg Zn L-1 of

ZnSO4 or nano-ZnO and the other half receiving 60 droplets (5 μL per droplet) of deionized water for 6 or 24 h (both the Zn solution and the deionized water both contained 0.05 % Tween 20). The leaves were cut in half along the mid-vein after incubation in the Petri dishes, then rinsed, oven dried (60 °C), digested using a 5:1 mixture of nitric acid and perchloric acid, and analysed using ICP-MS. Blanks and reference materials were included to ensure accuracy.

Examining subcellular distribution of foliar absorbed Zn in leaf tissues using NanoSIMS To supplement the µ-XRF scans of the leaf cross sections, we used NanoSIMS due to its higher resolution (ca. 100 nm). For NanoSIMS analyses, Zn distribution was examined only in leaf cross sections from two samples: 0 (control) and 6 h application of 1000 mg Zn L-1 -1 ZnSO4. Using the 16 d old sunflower, a 40 µL droplet of 1000 mg Zn L ZnSO4 with 0.05 % Tween 20 was applied on the adaxial surface of a YFEL. Another YFEL had a 40 µL droplet of deionized water (also containing 0.05 % Tween 20) applied the adaxial leaf surface as the control. The leaves were incubated in Petri dishes for 6 h with the growth lamps switched on. Afterwards, the leaves were rinsed and the area beneath the droplets excised with a steel razor blade and cut into 2 × 3 mm segments. These segments were placed in planchettes filled with hexadecane, and frozen in a high-pressure freezer (Bal-Tec HPM010), and processed as

151 described Kopittke et al. (2015). Next, 1 μm thick sections were cut using an ultramicrotome (LEICA EM UC6) and placed on substrates for NanoSIMS analysis.

NanoSIMS analysis were done similarly as described before (Blamey et al., 2018). Areas of interest were identified by optical microscopy in the NanoSIMS instrument. Signals of 12C14N and 64Zn16O secondary ions and secondary electron (SE) signal were collected to give morphological information and show the subcellular distribution of Zn. A large aperture (D1 = 1) and high primary current (~1nA) was used to presputter the areas of interest to achieve the steady state for secondary ions. Imaging was done with a smaller aperture (D1 = 2) with a primary current of 10 pA with total dwell time 56 ms per pixel for 256 × 256 pixels. Image analysis used the ImageJ plugin OpenMIMS (MIMS, Harvard University, Cambridge, MA, USA; https://nano.bwh.harvard.edu).

Examining the speciation of foliar absorbed Zn in bulk leaf tissues using XAS We used synchrotron-based XAS analysis to examine the chemical speciation of Zn in -1 YFELs to which 1000 mg Zn L as ZnSO4 and nano-ZnO had been applied for 6 h. The XAS analyses were performed at the XAS beamline at the Australian Synchrotron as described by Kopittke et al. (2011). To prepare these samples, 60 droplets (5 µL) of 1000 mg Zn L-1

ZnSO4 or nano-ZnO were placed evenly across the entire whole leaf, using separate YFELs (four replicates) of 16 d old sunflower plants. Leaves were rinsed after 6 h (as described earlier), frozen in liquid nitrogen, and ground using an agate mortar and pestle maintained under liquid nitrogen. The homogeneous specimen was then placed in a sample holder, sealed with Kapton tape, and immediately transferred to the cryostat (15 K) for analysis. At no time were the frozen (hydrated) samples allowed to thaw. In addition to the samples, we also prepared 11 standard reference compounds. First, we prepared three solid compounds, being

ZnO (Sigma Aldrich, 93632), Zn3(PO4)2 (Sigma Aldrich, 587583), and [ZnCO3]2·[Zn(OH)2]3 (Sigma Aldrich, 96466). These three solid compounds were ground and then diluted to a Zn concentration of 100 mg Zn kg-1 using cellulose. We also examined eight aqueous compounds, being Zn oxalate, Zn pectin, Zn citrate, Zn histidine, Zn(NO3)2, Zn phytate, Zn cysteine, and Zn glutathione. The Zn(NO3)2 standard had a final Zn concentration of 4 mM, while the other seven standards were made by mixing of 4 mM Zn(NO3)2 with 20 mM of oxalate acid, pectic acid, citric acid, histidine, phytic acid, cysteine, and glutathione. Where appropriate, pH was then adjusted to ca. 6 using 0.1 M NaOH. The eight aqueous standards

152 were then analysed following mixing with 30 % glycerol solutions (except Zn glutathione) which limits ice-crystal formation during cooling. Modelling using GEOCHEM-EZ (Shaff et al., 2010) indicated that > 99% of Zn was complexed with citric acid, oxalic acid, and histidine, and > 96% was with cysteine. Furthermore, for the Zn(NO3)2 standard, > 99.9% of 2+ the Zn was present as free Zn . Thus, hereafter, the Zn(NO3)2 standard is referred to as ‘Zn2+’.

The Zn Kα-edge X-ray absorption near edge structure (XANES) and extended X-ray absorption fine structure (EXAFS) spectra were obtained in fluorescence mode using a 100- element solid-state Ge detector. Two replicate spectra were collected for all samples and standards. Each spectrum was calibrated by simultaneous measurement in transmission mode of a metallic Zn foil. The two replicates were merged and normalized using Athena (version 0.8.056), and linear combination fitting (LCF) was performed with a maximum of three standards.

RESULTS Leaf properties of sunflower Three types of trichomes were observed on the surface of sunflower leaves (Fig. 1), being a type of NGT, linear glandular trichome (LGT), and capitate glandular trichome (CGT) according to the classification by Aschenbrenner et al. (2013). The NGTs were ca. 0.1-0.8 mm in length, 0.05-0.1 mm in width at the base and consisted of multi-stalk cells and base cells. The LGTs were ca. 0.05 mm in length and consisted of 6-11 linearly arranged cells (Fig. 1A,B,D). Finally, the CGTs were ca. < 0.05 mm in length and consisted of one short stalk cell and a round glandular cell on the tip (Fig. 1A,B,F). The total trichome density on sunflower adaxial leaf surface was 1,600 cm-2, with the NGTs accounting for ca. 55 % of the total trichomes (hence it can be estimated that the NGTs accounted for ca. 17 % of the leaf area), LNGs for 33 %, and CGTs for 12 % (Fig. 1). The stomatal density of the adaxial leaf side was 22,900 cm-2, with the size of the stomatal opening being ca. 14 × 3 µm (Fig. 1B,C). Secondly, as expected, the leaf consisted of upper epidermal cells, palisade mesophyll cells, spongy mesophyll cells, and a layer of lower epidermal cells. However, it was also noted that the NGTs and LGTs were generally connected to the bundle sheaths extensions (BSEs) (Wylie, 1952), with the BSEs either directly beneath the NGTs and LGTs or connecting with the edge of trichome bases (Fig. 1D,E,G,H). It was also noted that the BSEs extended

153 horizontally in the leaf tissues (Fig. 1G) and that they connected with vascular tissues (Fig. 1I).

The distribution of Zn in intact leaves using µ-XRF The distribution of Zn in the leaves was examined in situ using synchrotron-based µ-XRF after foliar application for 15 min, 30 min, 1 h, 3 h, and 6 h. First, we conducted survey scans which examined larger areas of the leaves (Fig. 2), with Zn detected in the leaf tissues underneath the droplets within only 15 min (Fig. 2F), and with the tissue Zn concentration increasing as the exposure time increased. However, for all treatments, the Zn was observed to accumulate primarily underneath the Zn droplets, suggesting the Zn that had moved across the leaf surface had only limited translocation away from the site of application. Surprisingly, it was also observed that there were some bright spots (i.e. high Zn concentrations), ca. 0.05- 0.1 mm in size, with the frequency and the intensity of these spots increasing with exposure time (Fig. 2).

Based on these survey scans, we conducted detailed scans to examine smaller areas entirely underneath the droplets (Fig. 3). Using tri-colour maps of Ca-Mn-Zn, it was found that the ‘bright spots’ identified from the survey scans were bases of NGTs (Supplementary Fig. S1). Indeed, within only 15 min, Zn was observed to accumulate at the base of some of the NGTs, with some Zn also found to accumulate within the veins (Fig. 2F and Fig. 3A). As the exposure time increased, the number of NGTs that had high concentrations of Zn in their basal cells also increased. Indeed, at the final period of exposure (6 h), the highest Zn concentrations were found in the basal cells of the NGT, with Zn also accumulating (to a lesser extent) in the vascular tissues (Fig. 3).

The distribution of Zn in cross sections examined using NanoSIMS and µ-XRF Next, the distribution of Zn was examined in cross sections (1 µm thick) using NanoSIMS in order to obtain information at a subcellular level. We examined both a control and a leaf exposed to ZnSO4 for 6 h. By comparing with the control (Supplementary Fig. S2), for the leaf exposed to ZnSO4 for 6 h, it was found that the entire abaxial surface contained elevated concentrations of Zn. It appeared that Zn had moved across the entire leaf surface rather than only at specific locations (such as stomata). Indeed, Zn appeared to have penetrated through the entire cuticle area underneath the Zn droplet (Fig. 4). Of particular importance, there was not any marked accumulation of Zn within the stomatal cavity and surrounding cells, 154 indicating that there did not appear to be preferential movement of Zn through the stomata (compare Fig. 4C,D and Fig. 4E,F). In addition, where the epidermis connected with the palisade cells, Zn accumulated primarily within the epidermal cell walls (Fig. 4A,B). In contrast, where the epidermis connected with vascular tissues, Zn was observed to not only accumulate in the cell walls of the epidermal cells, but also in the cell walls of the vascular cells (Fig. 4A,B,I,J). Unfortunately, we were only able to examine a small area using NanoSIMS, and the analyses did not include any NGTs.

Then, we examined the distribution of Zn using µ-XRF in cross sections (150 µm thick) of leaves that had been exposed to ZnSO4 for 15 min, 30 min, 1 h, 3 h, and 6 h. Initially, we compared fresh and freeze dried cross sections that had been exposed for 3 or 6 h of ZnSO4, finding that the freeze drying process did not appear to alter Zn distribution (Fig. 5 and Supplementary Fig. S3). Hence, hereafter we examined freeze dried samples (Fig. 5). Firstly, as observed from the µ-XRF analyses of the intact leaves (Fig. 2 and Fig. 3), Zn was again found to accumulate rapidly (in ≤ 15-30 min) in the bases of the NGTs on the adaxial surface (Fig. 5A and Supplementary Fig. S4). After exposure for ≥ 1 h, not only did the Zn concentration in the basal cells of the NGTs of the adaxial surface increase, but Zn was also detected in the inner leaf tissues, as well as in epidermal cells and NGTs on both leaf surfaces (adaxial and abaxial) (Fig. 5A).

From these µ-XRF scans, it is possible to compare the amount of Zn within the various tissues. Analysis of the data from the µ-XRF scans showed that normalised counts for Zn in the NGTs was averagely ca. five times higher than the cuticular areas (including cuticle, stomata, and the rest of the trichome areas) after 15 min and ca. nine times higher than the cuticular areas after 6 h (Supplementary Fig. S5). As noted earlier, the NGTs accounted for ca. 17 % of the leaf area, hence after exposure for 6 h, it is estimated that the total Zn accumulation in the NGTs was 1.9-fold higher than the Zn that accumulated in the cuticular tissues between the NGTs.

Of particular interest, it is known that BSEs generally connect the NGTs on opposite sides of the leaf (Fig. 1), with the µ-XRF analyses of cross sections appearing to show that Zn moved readily from the NGT through the BSEs to the opposite leaf surface (Fig. 5B). Given (i) the penetrating nature of X-rays, and (ii) that the sections examined by µ-XRF were 150 µm

155 thick (ca. 10 cellular layers, see Fig. 1), the exact cellular and subcellular distribution of Zn is not clear from the µ-XRF analyses.

Comparing the foliar absorption of nano-ZnO and ZnSO4 -1 Firstly, droplets of 1000 mg Zn L nano-ZnO and ZnSO4 were applied to the same leaf before being scanned using µ-XRF after either 3 h or 3 d (Fig. 6A,B). It was found that the absorption of nano-ZnO was substantially lower than that of ZnSO4 regardless of exposure time, with this result agreeing with measurements of bulk Zn (Supplementary Fig. S6). For example, foliar absorption of ZnSO4 was 92-fold higher than that of nano-ZnO after 6 h exposure (Supplementary Fig. S6). However, the localization of the foliar absorbed Zn was highly limited for both forms of Zn, with the Zn accumulating almost exclusively in the tissues underlying where the droplet had been applied, even after 3 d (Fig. 6B). Although the concentration of Zn in the tissues underlying the nano-ZnO droplet was substantially lower than the ZnSO4 exposure leaf, the distribution of Zn within the tissues was similar, with the highest Zn concentrations again found in the basal cells of the NGTs (Fig. 5 and Fig. 6C).

Speciation of Zn within leaf tissues To compare the speciation of the foliar-absorbed Zn, leaves exposed to 1000 mg Zn L-1 nano-

ZnO or ZnSO4 for 6 h were examined using XAS (Fig. 7). First, we compared the spectra of the 11 standard compounds (black lines in Fig. 7). There were clear differences among the various spectra, differing not only in the energy of the white line peak and the magnitude of the peak, but also differing in other spectral features. For example, compounds in which Zn is complexed with carboxyl groups (oxalate, pectin, citrate) or is present as free Zn2+

[Zn(NO3)2] tended to have a white line peak at 9,669 eV, while Zn3(PO4)2 had a peak at 9,665 eV and ZnO at 9,671 eV. Other differences were also evident among the standard compounds, with Zn phytate and Zn cysteine tending to have broader peaks compared to compounds in which Zn is complexed with carboxyl groups.

Next, we compared the spectra from the two leaf tissue samples which had been exposed to either ZnSO4 or nano-ZnO for 6 h (red lines in Fig. 7). Although the standard spectra for the two fertilisers differed markedly (compare black lines for Zn2+ and ZnO), the spectra for leaf tissues were visually similar, despite they being exposed to the two different fertilisers (red lines in Fig. 7). Of particular importance, it was noted that the spectra of Zn in leaves exposed to nano-ZnO differed markedly from the spectra of ZnO (compare black line for ZnO with 156 red line for 6 h nano-ZnO, Fig. 7). Using LCF, it was modelled that 40-50 % Zn within the leaf tissues of both treatments was present as Zn phytate, with 30-40 % present as Zn oxalate (Supplementary Table S1). However, given the similarity in the spectra of many of the standard compounds (for example, see Supplementary Fig. S7), it was not possible to unequivocally identify the exact forms of Zn within the leaf tissues.

DISCUSSION In the present study, we examined the importance of the cuticle, stomata, and trichomes in foliar absorption of Zn. We found that the trichomes appear to be of particular importance, with Zn rapidly (≤ 15 min) accumulating within the basal cells of NGTs (Fig. 3, Fig. 5, and Supplementary Fig. S1). The cuticle was also found to be important, with Zn found to accumulate in the walls of all epidermal cells underlying the Zn-containing droplet (Fig. 4). Interestingly, there was no apparent accumulation of Zn in the stomatal cavity or the surrounding cells (Fig. 4), indicating that the stomata were likely not an important pathway for the foliar absorption of Zn in sunflower. We also compared nano-ZnO and ZnSO4, finding that the absorption of nano-ZnO was substantially lower than for ZnSO4 (Fig. 6 and Supplementary Fig. S6) despite the pattern of absorption being similar (Fig. 6 and Fig. 7). Finally, although the absorbed Zn had only limited subsequent transportation from the application site (Fig. 2 and Fig. 6), our analyses of Zn distribution appeared to indicate that the Zn moved through the BSEs and other vascular tissues to the opposite leaf side (Fig. 4 and Fig. 5).

Role of trichomes in foliar Zn absorption In the present study investigating sunflower, we have shown that trichomes appear to be an important organ for foliar-applied Zn across the leaf surface, with the NGTs containing the highest Zn concentrations within the tissues to which the Zn was applied (Figs. 2-3, 5-6, and Supplementary Figs. S2 and S5). Specifically, Zn accumulated rapidly (≤ 15 min) in the basal cells of the NGTs (Fig. 3 and Fig. 5) and the total Zn accumulation amount in the NGTs for 6 h exposed leaf was ca. 1.9 times higher than that in the cuticular area. There are several possible reasons why Zn moved rapidly into the base of NGTs. Firstly, it is possible that the area surrounding the NGTs had a higher wettability than the surrounding interveinal tissues. Secondly, it is possible that the chemical composition and structure of the NGTs’ cuticle differs from the cuticle covering the main portion of the leaf surface, and it could be more permeable to solutes. For example, Bahamonde et al. (2018) showed that the major veins of 157 the adaxial beech (Fagus sylvatica) leaf surface are more permeable than the other leaf areas. Thirdly, it is also possible that sunflower has a specialised mechanism for the absorption of water through trichomes, as has been noted for other plant species adapted to drought stress (Ohrui et al., 2007, Winkler et al., 2010, Vitarelli et al., 2016,), with this resulting in the concomitant absorption of Zn in the present experiment. However, it is known that trichomes are highly diverse (Ohrui et al., 2007, Werker, 2000), and it is likely that not all plant species would have trichomes that are important for foliar nutrient absorption. For example, Li et al. (2018) found that trichomes of tomato (Solanum lycopersicum) and soybean (Glycine max) are not important in foliar Zn absorption. Thus, the role of trichomes in foliar nutrients absorption depends upon the ecological conditions, species, and the property of the specific trichome.

The observation that the NGTs contained elevated concentrations of Zn following its foliar application is similar to that observed in other studies where elevated levels of metals are supplied in the rooting medium. For example, Blamey et al. (2015) found that high concentrations of Mn accumulated in these NGTs of sunflower when exposed to elevated levels of Mn in the rooting medium [also see Sarret et al. (2009) in Arabidopsis halleri and Arabidopsis lyrata, Broadhurst et al. (2009) in Alyssum murale and Alyssum corsicum, McNear et al. (2014) in Alyssum murale, and Tomasi et al. (2014) in Cucumis sativus]. Furthermore, the prediction that 30-40 % of the foliar absorbed Zn is associated with oxalate (Supplementary Table S1) is presumably associated with the accumulation of Zn in the NGTs, with Zn potentially substituting for Ca in the Ca-containing compounds of the NGTs. This is similar to that reported by Sarret et al. (2006) in tobacco (Nicotiana tabacum), with these authors finding that when exposed to toxic levels of Zn, the Zn preferentially accumulated in trichomes, with Zn substituted calcite excreted as a mechanism of detoxification. However, whether the Zn that accumulates within the NGTs can subsequently be translocated to other parts of the plant requires further study. In a similar manner, it is necessary to better clarify the accumulation of Zn within trichomes that we have observed in the present study. In particular, it is important to understand whether the rapid movement of Zn into the NGT is a specific absorption mechanism, or whether it occurs indirectly due to absorption of water as a drought stress mechanism. In a related manner, it is necessary to understand differences among plant species, including why the trichomes of some plant species do not absorb foliar-applied nutrients (Li et al., 2018) but others do. Finally, it would

158 also be useful to assess the impact of growth conditions (such as water stress) on the absorption of nutrients by trichomes.

Role of the cuticle and stomata in foliar Zn absorption Apart from the important role of trichomes in the movement of Zn across the leaf surface, we also found that Zn appeared to move across the leaf cuticle. The importance of the cuticle was demonstrated by the subcellular distribution of Zn examined using NanoSIMS (Fig. 4). To the best of our knowledge, this is the first time that NanoSIMS has been used for the examination of foliar nutrient absorption. We found that Zn concentrations were increased and appeared to be comparatively uniform along the epidermal cells of the adaxial surface (Fig. 4 and Supplementary Fig. S2), indicating the movement of Zn across the cuticle. But it needs to note that the relative humidity during foliar Zn application in the present study was 98 %, this could promote the cuticular penetration (Fernández et al., 2009, Schonherr, 2001). Furthermore, cuticular permeability differs among the various plant species (Schreiber et al., 1996; Schreiber and Schönherr, 2009), depending upon the chemical composition of the cuticle, including cutin (Sadler et al., 2016, Huang et al., 2017) and intracuticular wax (Zeisler-Diehl et al., 2018), with the permeability enhanced greatly by adjuvants such as Tween 20 (Arand et al., 2018). However, much remains unknown regarding the structure, composition, formation, and the physicochemical properties of the cuticle (Fernández et al., 2017, Khanal et al., 2017, Ingram et al., 2017, Schuster et al., 2017), as do mechanisms by which Zn moves across the cuticle.

We found no evidence that the stomata played an important role in the foliar absorption of Zn. Rather, concentrations of Zn in the stomatal cells were similar to those observed for other epidermal cells on the adaxial surface, with no distinct accumulation of Zn in the stomata or the stomatal cavity (Fig. 4A-F). This observation is consistent with that of Eichert et al. (2008) who visually observed that hydrophilic polystyrene particles with a diameter of 43 nm suspended in water entered leaves through the stomata by diffusion along the walls of the pore other than from the opening cavity, but less than 10 % of all stomata participated in this pathway. The role of hygroscopic leaf surface particles on promoting stomatal uptake can also not be neglected (Burkhardt et al., 2012). Despite the potential hydrophobility of the stomatal cavity and its geometry which limit solution infiltration (Schönherr and Bukovac, 1972), the accumulation of hygroscopic particles in the pore may lead to the formation of a continuous liquid film connecting along the cuticular walls of the stomata between the 159 interior and the outer surface, with this process called “hydraulic activation of stomata” (Burkhardt et al., 2009, Burkhardt, 2010). The activation of this process is required separately for each stomata and could be relevant to the composition or structure of the cuticle above the guard cells or the pore wall cells (Burkhardt et al., 2012), modifications of the pore such as deposition of hygroscopic particles, and microbes growing in the pore which increased wettability (Eichert et al., 2012, Burkhardt et al., 2012). However, major differences in the chemical composition and structure of the pore walls and stomatal surfaces can be found among species and this is likely to affect the process of stomatal penetration and the significance and contribution of this pathway to the overall process of foliar absorption as noted by Fernández et al. (2017).

Role of bundle sheath extensions in movement of foliar absorbed Zn In the present study, we have identified an important role of BSEs, with little currently known regarding these tissues (Wylie, 1952, Buckley et al., 2011). In sunflower, the BSEs extend vertically and horizontally within the leaf (Fig. 1). We noted that following the accumulation of Zn at the NGT bases, the Zn appeared to move comparatively rapidly into BSEs, and as a result, although Zn was applied to the adaxial surface, it also moved rapidly to the abaxial surface (Fig. 5). Thus, we propose that in sunflower, BSEs are involved in the movement of foliar-applied Zn from the trichome into other leaf tissues. However, in the present study, we examined changes in Zn distribution rather than Zn movement per se. Although the changes in distribution reflect the underlying movement of Zn, further studies are required to examine the redistribution of the absorbed Zn.

Comparing the absorption of nano-ZnO and ZnSO4 There is increasing interest in the use of nanomaterials in agricultural production systems (De Rosa et al., 2010, Wang et al., 2016). We found that the foliar absorption of nano-ZnO was substantially lower than that of ZnSO4 regardless of the exposure time, but the two types of Zn had a similar pattern of absorption (Fig. 6 and Supplementary Fig. S6). Furthermore, we found that the Zn within the leaf tissues that had moved across the leaf surface was present as the same chemical species, regardless of whether the Zn was supplied as nano-ZnO or ZnSO4 (Fig. 7). These findings indicate that nano-ZnO was likely absorbed in the same form as

ZnSO4 – soluble Zn ions rather than as intact ZnO particles. This is in agreement with previous studies, including a similar studies of Li et al. (2018) in soybean and Alexander et al. (2016) found that absorption of insoluble foliar applied nutrients was limited (even for

160 nano-scale particles). Therefore, solubility is an important factor to consider when designing foliar fertilisers. However, this slower release of Zn from nano-ZnO is potentially advantageous, as it may reduce any potential ‘burning’ observed from highly soluble Zn foliar fertilisers (Drissi et al., 2015). In this regard, it would be useful to identify how sparingly soluble forms of Zn may be retained on the leaf surface as a slow release fertiliser.

CONCLUSIONS We have demonstrated that trichomes are important for the absorption of foliar-applied Zn for leaves of sunflower. The cuticle was also found to be important, with Zn appearing to move across the cuticle before accumulating in the walls of underlying cells. No marked accumulation of Zn was found within the stomatal cavity, indicating that movement of Zn through the stomata was not likely to be important. After absorption, Zn was located in the epidermal cells on both sides of the leaf. In this regard, the BSEs which connected to the trichomes appear to be important for the movement of Zn away from the overlying trichomes.

Finally, the absorption of nano-ZnO was substantially lower than for ZnSO4, with Zn likely moving across the leaf surface as soluble Zn rather than as nanoparticles. It is hoped that the information from this study will assist in improving the efficacy of foliar fertilisers as required to improve the nutrition of both crop plants and humans.

SUPPLEMENTARY DATA Fig. S1 Tri-colour image from µ-XRF analyses showing the distribution of Ca, Zn, and Mn in a sunflower leaf directly beneath where a ZnSO4 droplet had been applied for 6 h. Fig. S2 NanoSIMS analyses showing Zn distribution in leaves of sunflower exposed to

ZnSO4 for 6 h and a control leaf. Fig. S3 µ-XRF analyses showing the distribution of Zn in fresh (hydrated) cross sections of sunflower leaves, to which ZnSO4 had been applied for either 3 or 6 h. Fig. S4 Tri-colour image from µ-XRF analyses showing the distribution of Ca, Zn, and Mn in cross sections of sunflower leaf.

Fig. S5 µ-XRF image of leaf treated with ZnSO4 for 15 min and 6 h, with the upper µ-XRF images showing Zn distribution and the bottom graphs showing the normalised concentration of Zn along the transect (white lines) shown in the upper images.

Fig. S6 Comparison of foliar Zn absorption of ZnSO4 and nano-ZnO in sunflower using inductively coupled plasma mass spectrometry.

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Fig. S7 Normalized K-edge XANES spectra illustrating the close similarity of four Zn compounds which prevented differentiation during linear combination fitting. Table S1 Results of linear combination fitting of K-edge XANES data for sunflower leaves to which Zn had been applied for 6 h.

ACKNOWLEDGEMENTS This work was supported by Sonic Essentials. Funding was also provided by the Australian Research Council (ARC) through the Linkage Projects funding scheme (LP130100741), the Future Fellowship scheme (FT120100277, Peter Kopittke), and the Discovery Early Career Researcher Award scheme (DE160100429, Antony van der Ent). Cui Li is supported by the China Scholarship Council and The University of Queensland (UQ). The authors acknowledge use of the facilities and technical assistance of both the Monash Histology Platform (Department of Anatomy and Developmental Biology, Monash University), the Queensland Brain Institute (UQ) and the Centre for Microscopy and Microanalysis (UQ). Much of this research was undertaken on the XFM and XAS beamlines at the Australian Synchrotron, part of ANSTO. The assistance of both Chris Glover (XAS beamline, Australian Synchrotron) and Victoria Fernández is gratefully acknowledged.

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FIGURES

Fig. 1 Trichomes on the sunflower leaf surface, with the NGTs (non-glandular trichomes), LGTs (linear glandular trichomes), and CGTs (capitate glandular trichomes) labelled accordingly. The bundle sheath extensions (BSEs) are also labelled. A-C, images of the leaf surface from scanning electron microscopy. D-F, light micrographs showing leaf cross sections (1 µm thick), with the sections stained using toluidine blue. G-I, light micrographs (80 µm thick) of fresh leaf cross sections without staining.

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Fig. 2 A-E, light micrographs showing the Zn-containing droplets (ZnSO4) on the sunflower leaf surfaces before being removed and rinsed. F-J, images from the µ-XRF survey scans showing the distribution of Zn. All Zn maps are presented on the same concentration scale (mg/kg), with brighter colours corresponding to higher Zn concentrations. Droplets were left on the leaf surface for 15 min, 30 min, 1 h, 3 h, or 6 h. The scale bar in A applies to A-E, and the scale bar in F applies to F-J.

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Fig. 3 Detailed µ-XRF scans showing the distribution of Zn in hydrated leaves of sunflower to which ZnSO4 had been applied for 15 min (A), 30 min (B), 1 h (C), 3 h (D), or 6 h (E). F, high magnification of the white box on D. All Zn maps are presented on the same concentration scale (mg/kg), with brighter colours corresponding to higher Zn concentrations. All of the areas imaged were entirely underlying where the Zn-containing droplet had been applied. The scale bar in A applies to A-E.

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Fig. 4 NanoSIMS images showing the distribution of Zn in a cross section of a sunflower leaf 12 14 that had been exposed to ZnSO4 for 6 h. The C N images are in Grayscale for identification of the cellular and subcellular structures (A, C, E, G and I). The 64Zn16O images are given as a 16 colour scale showing Zn distribution (B, D, F, H, and J). C-D, the Zn distribution for a stomata, which correspond to the yellow rectangle in A. E-F, the Zn distribution in the epidermis, which correspond to the purple rectangle in A. G-J, the Zn distribution in vascular tissues, which correspond to the green rectangle (G-H) and orange rectangle (I-L) in A. The scale bar in A applies to B, and the scale bar in C applies to D-J.

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Fig. 5 A, µ-XRF scans showing Zn distribution in cross sections of sunflower leaves (150

µm, freeze dried) underlying ZnSO4 droplets that had been applied for 0 min (control), 15 min, 30 min, 1 h, 3 h, or 6 h. B, a close-up of a portion of a cross section (3 h exposure) showing the movement of Zn through the bundle sheaths extensions from the adaxial (Zn applied side) to the inner leaf tissues and abaxial side. Note that the µ-XRF image and light micrograph in B are not from the same sample. Rather, the light micrograph is simply provided for reference. Brighter colours correspond to higher Zn concentrations, but colours are not comparable among these Zn maps as their Zn concentrations differed too much to show using the same colour threshold.

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Fig. 6 Comparison the foliar absorption of nano-ZnO and ZnSO4 in leaves of sunflower after exposure for either 3 h (A) or 3 d (B). In A and B, images on the left are light micrographs showing the Zn droplets on leaves (before being removed), while the images on the right are µ-XRF survey scans showing the distribution of Zn. C, a µ-XRF scan showing distribution of Zn a cross section of a sunflower leaf (150 µm thick, freeze dried) underlying the Zn droplets after 6 h exposure of nano-ZnO. Brighter colours correspond to higher Zn concentrations, but colours are not comparable among these Zn maps as their Zn concentrations differed too much to show using the same colour threshold.

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Fig. 7 Normalized K-edge XANES spectra of 11 standard Zn compounds (black lines) and sunflower leaf tissues exposed to two different forms of Zn (red lines).

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Absorption of foliar-applied Zn in sunflower (Helianthus annuus): Importance of the cuticle, stomata, and trichomes

Cui Li1*, Peng Wang2,3, Antony van der Ent4,5, Miaomiao Cheng6, Haibo Jiang7, Thea Lund

Read8, Enzo Lombi8, Caixian Tang6, Martin D. de Jonge9, Neal W. Menzies1, Peter M.

Kopittke1

1The University of Queensland, School of Agriculture and Food Sciences, St Lucia, Queensland, 4072, Australia; 2Nanjing Agricultural University, College of Resources and Environmental Sciences, Nanjing, 210095, China; 3The University of Queensland, Centre for Soil and Environmental Research, School of Agriculture and Food Sciences, St Lucia, Queensland, 4072, Australia; 4Centre for Mined Land Rehabilitation, Sustainable Minerals Institute, The University of Queensland, Australia, 5Laboratoire Sols et Environnement, Université de Lorraine, France; 6La Trobe University, Centre for AgriBioscience, Bundoora, Victoria, 3086, Australia; 7University of Western Australia, Centre for Microscopy, Characterization and Analysis, Crawley, WA 6009, Australia; 8University of South Australia, Future Industries Institute, Mawson Lakes, South Australia, 5095, Australia; 9ANSTO, Australian Synchrotron, Clayton, Victoria, 3168, Australia

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Fig. S1 Tri-colour image from µ-XRF analyses showing the distribution of Ca (red), Zn

(green), and Mn (blue) in a sunflower leaf directly beneath where a ZnSO4 droplet had been applied for 6 h. The red stalks (Ca) and the blue bases (Mn) are the non-glandular trichomes (NGTs), with the Zn (green) accumulating at the NGT bases.

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Fig. S2 Comparison of Zn distribution in leaves of sunflower exposed to ZnSO4 for 6 h (A-C) and a control leaf (D-F). Analyses were performed using NanoSIMS, with red colour being Zn and blue colour being S. The scale in A applies to all the images.

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Fig. S3 Images from µ-XRF analyses showing the distribution of Zn in fresh (hydrated) cross sections (150 µm thickness) of sunflower leaves, to which ZnSO4 had been applied for either 3 (left) or 6 h (right). By comparison of these images of hydrated tissues with the freeze-dried tissues of Fig. 5, we conclude that Zn does not appear to be redistributed during the freeze- drying process. Brighter colours correspond to higher Zn concentrations. Colours are not comparable between the two Zn leaf sections (3 and 6 h) as their Zn concentration differed too greatly to show in the same colour threshold.

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Fig. S4 Tri-colour image from µ-XRF analyses showing the distribution of Ca (red), Zn

(green), and Mn (blue) in freeze dried cross sections of a control (left) and a 3 h ZnSO4 treated (right) sunflower leaf. The red stalks (Ca) and the blue bases (Mn) are the non- glandular trichomes (NGTs), with the Zn (green) accumulating at the NGT bases and the inner leaf tissues.

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Fig. S5 µ-XRF image of leaf treated with ZnSO4 for 15 min and 6 h, with the upper µ-XRF images showing Zn distribution (note that the two µ-XRF images are not comparable) and the bottom graphs showing the normalised concentration of Zn along the transect (white lines) shown in the upper images.

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Fig. S6 Comparison of foliar Zn absorption of ZnSO4 and nano-ZnO in sunflower using inductively coupled plasma mass spectrometry (ICP-MS). Data are mean ± SD, n = 4.

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Fig. S7 Normalized K-edge XANES spectra illustrating the close similarity of four Zn compounds which prevented differentiation during linear combination fitting.

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Table S1 Results of linear combination fitting of K-edge XANES data for sunflower leaves to which Zn had been applied for 6 h. Values within both columns do not add to 100 %. This is because of the similarity in the spectra of many standard compounds (Fig. S7), which meant that it was not possible to unequivocally identify the remaining forms of Zn within the leaf tissues. XANES Standards (%) ZnSO4 Nano-ZnO Phytate 37 ± 2 51 ± 2 Oxalate 34 ± 2 38 ± 2

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Chapter 6 Absorption of foliar applied Zn is decreased in Zn deficient sunflower (Helianthus annuus) due to changes in leaf properties

6.1 Introduction

The foliar nutrient absorption rate, for example for B and P, is known to be affected by the plant nutrient status (Will et al., 2011, Fernández et al., 2014a). However, previous studies in tomato (Du et al., 2015) and wheat (Erenoglu et al., 2002) reported that the plant Zn status did not influence the foliar Zn absorption. In this study, using µ-XRF and ICP-MS, the absorption of foliar applied nano-ZnO and ZnSO4 was compared between Zn deficient and Zn sufficient sunflower plants. In addition, the chemical speciation of the absorbed Zn in the leaves was also compared using XAS. The aim was to examine whether plant Zn status affects foliar Zn absorption and the speciation of the foliar absorbed Zn.

6.2 Paper 5

Li C, Wang P, Lombi E, Wu J, Blamey FPC, Fernández V, Daryl HL, Menzies NW, Kopittke PM (2018) Absorption of foliar applied Zn reduced in Zn deficient sunflower (Helianthus annuus) due to the changing of leaf properties. Plant and Soil 433: 309-322.

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Absorption of foliar applied Zn is decreased in Zn deficient sunflower (Helianthus annuus) due to changes in leaf properties

Cui Li1, 2*, Peng Wang2, 3, Enzo Lombi4, Jingtao Wu2, F. Pax C. Blamey2, Victoria Fernández5, Daryl L. Howard6, Neal W. Menzies2, Peter M. Kopittke2

1 Northwestern Polytechnical University, Research Centre for Ecology and Environmental Sciences, Xi’an, China 2 The University of Queensland, School of Agriculture and Food Sciences, St Lucia, Queensland, Australia 3 Nanjing Agricultural University, College of Resources and Environmental Sciences, Nanjing, China 4 University of South Australia, Future Industries Institute, Mawson Lakes, South Australia, Australia 5 Technical University of Madrid, School of Forest Engineering, Forest Genetics and Ecophysiology Research Group, Madrid, Spain 6 Australian Synchrotron, Clayton, Victoria, Australia

Author for correspondence: Cui Li The University of Queensland School of Agriculture and Food Sciences St Lucia, Queensland, 4072 Australia Email: [email protected]; Tel: +61 424 662 660

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Abstract Aims Despite the importance of foliar Zn fertilisation, it remains unclear how the Zn status of the plant itself influences the efficacy of foliar applied Zn, with this forming the focus of the present study.

Methods Sunflower (Helianthus annuus) was grown in nutrient solutions with either 0.5 (sufficient) or 0 µM Zn (deficient) to relate the absorption of foliar applied Zn to leaf properties. The distribution and speciation of Zn within the leaf were examined in situ using synchrotron-based X-ray fluorescence microscopy and X-ray absorption spectroscopy.

Results Zinc deficiency decreased foliar absorption of Zn as ZnSO4 by 50-66% compared to Zn sufficient sunflower, despite µ-XRF analysis showing the pattern of Zn absorption was similar for both plants. Rather, Zn deficiency decreased the leaf trichome density and likely altered the adaxial leaf surface composition and structure, with these presumably being the main causes for the reduced Zn absorption. The Zn status of the plant also influenced the speciation of the absorbed Zn, with 37-53% as Zn phytate in Zn sufficient leaves but 55% as Zn phosphate in Zn deficient leaves.

Conclusions Zinc status of the plants influenced leaf surface properties, with Zn deficiency leading to decreased absorption of foliar applied Zn.

Keywords Cuticle; Foliar absorption; Nano-Zn; Zn foliar fertiliser; Zn speciation

Abbreviations ICP-MS Inductively coupled plasma mass spectrometry ICP-OES Inductively coupled plasma optical emission spectroscopy LCF Linear combination fitting XAS Synchrotron-based X-ray absorption spectroscopy XANES K-edge X-ray absorption near edge structure spectra YFELs Youngest fully-expanded leaves µ-XRF Synchrotron-based X-ray fluorescence microscopy

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Introduction Zinc (Zn) deficiency is a common physiological disorder affecting plants and is associated with low Zn availability, occurring for example in alkaline, saline, weathered tropical or waterlogged soils (Cakmak and Kutman 2018). Indeed, Zn deficiency in plants is a global problem, with an estimated 49% of the world’s important agricultural soils having low levels of plant-available Zn (Sillanpää 1990). As a result, both the yield and nutritional quality (i.e. Zn concentration) of food produced from these soils are reduced (Sadeghzadeh 2013). Furthermore, Zn deficiency in humans is often caused by low dietary intake that is associated with a large consumption of cereal-based foods which contain very low Zn concentrations (Cakmak and Kutman 2018). It is estimated that approximately one third of the world’s human population suffers from Zn deficiency (Cakmak et al. 2010). Foliar Zn fertilisation is being increasingly used (Fernández et al. 2013), and it has proven to be a promising Zn biofortification approach for increasing Zn bioavailability in agricultural commodities (Velu et al. 2014; Joy et al. 2015; Zhang et al. 2018). Foliar fertilisation is particularly useful when soil conditions limit availability of soil applied Zn, for example, in soils with free CaCO3 or soils with high pH (Alloway 2009). However, much is still unknown about the mechanisms of application, absorption, and translocation of foliar Zn fertilisers. Hence, improving our understanding of such foliar absorption aspects will allow for increasing the effectiveness of Zn foliar fertilisers as tool for controlling Zn deficiency and for biofortification of crop plants.

The effects of the nutritional status on plant surface traits and absorption of foliar applied nutrients have been examined in only a few studies. In this regard, it was found the amount of wax per unit surface was decreased in Fe-deficient leaves of peach (Prunus persica) (Fernández et al. 2008) and P-deficient leaves of wheat (Triticum aestivum) (Fernández et al. 2014a). It was also found that the cuticle weight per unit surface and the size of stomatal pores decreased in Fe-deficient pear (Pyrus communis) leaves (Fernández et al. 2008). In a similar manner, the leaf trichome and stomatal densities were decreased under P deficiency (Fernández et al. 2014a). With these effects on the leaf surface traits, Will et al. (2011) found that the foliar absorption of B in soybean (Glycine max) that was B deficient was significantly lower than B sufficient soybean. Similarly, Fernández et al. (2014a) found that absorption of foliar-applied P was lower in P deficient wheat than in P sufficient wheat. In contrast to these studies, however, Erenoglu et al. (2002) reported that foliar absorption of Zn

187 in wheat was not influenced by Zn nutritional status, but more Zn translocated out of Zn deficient leaves than from leaves with sufficient Zn.

Given the lack of knowledge on the mechanisms of foliar absorption in relation to plant nutritional status and the significance of Zn deficiency in agricultural crops, the present study analysed the foliar absorption of Zn as affected by the Zn status of the plants itself. Furthermore, with increasing interest in the potential use of engineered nanomaterials within agricultural production systems (DeRosa et al. 2010; Wang et al. 2016), and with sunflower (Helianthus annuus) being an important agricultural crop, we compared the rates of foliar absorption of ZnSO4 and nano-ZnO in Zn sufficient and Zn deficient sunflower, and attempted to relate potential foliar Zn absorption differences to changes in leaf surface properties, including trichome density, stomatal density, gross adaxial surface composition, and fertiliser droplet contact angles. In addition, we used synchrotron-based X-ray fluorescence microscopy (µ-XRF) for in situ analyses of changes in Zn accumulation within the leaf tissue. We also used synchrotron-based X-ray absorption spectroscopy (XAS) for in situ analyses of changes in Zn speciation within the leaf tissue. The results of the present study will assist understanding the processes involved in foliar Zn absorption and provide information for improving the efficiency of foliar Zn fertilisation.

Materials and Methods Plant growth and preparation of Zn foliar fertilisers Sunflower (cv. Hyoleic 41) seeds were germinated in paper towel suspended vertically in tap water for 3 d in a laboratory at The University of Queensland (St Lucia, Australia). Then, seedlings were transferred to 11 L black containers (the lid of each container had four holes - and each hole had one plant), with nutrient solutions that contained (µM): 910 N (94 % NO3 + and 6 % NH4 ), 20 P, 475 K, 1126 Ca, 227 Mg, 1251 Cl, 556 S, 25 Fe(III)-EDTA, 3 B, 0.5 Mn, 0.2 Cu and 0.01 Mo (Blamey et al. 2015). There were two treatments, 0 and 0.5 µM Zn, supplied as ZnSO4, designed to induce Zn deficiency and supply sufficient Zn. Nutrient solution pH was unadjusted and was ca. 5.5 for both treatments. Light was provided using high-pressure sodium lamps (photon flux density of 1500 μmol m–2 s–1) for 12 h d-1 and the temperature maintained at 25 °C. Nutrient solutions were replaced after the first 7 d then every 4 d thereafter. After 10 d, 5 mL of 44 mM KH2PO4 was added to each 11 L container every other day to replenish P that had been taken up by the plants (Blamey et al. 2015).

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After growing for 40 d, the youngest fully-expanded leaves (YFELs) were used for the measurements outlined below.

-1 Two types of Zn foliar fertilisers were prepared: 1000 mg Zn L (15.4 mM) as ZnSO4.7H2O (pH 5.2) and 1000 mg Zn L-1 as nano-ZnO (ca. 30 nm diameter, pH 7.1, sparingly soluble, none commercial product) [see Li et al. (2018a) for details of the nano-ZnO]. The nano-ZnO suspension was mixed for 15 min using an ultrasonic processor before use. Both Zn solutions contained 0.05% Tween 20 (Sigma-Aldrich) as a surfactant.

Bulk Zn foliar absorption Foliar absorption of Zn was analysed by measuring the bulk Zn concentration in the leaves using inductively coupled plasma mass spectrometry (ICP-MS) after the plants had been growing in the nutrient solutions for 16, 28, and 40 d. Both ZnSO4.7H2O and nano-ZnO were used at 40 d while only ZnSO4.7H2O was used at 16 and 28 d. A half leaf method was used as described by Li et al. (2018a). Briefly, at each time interval, 70 droplets (5 µl) of the appropriate Zn-containing solution were applied to half of the adaxial leaf, while 70 droplets (5 µl) deionised water were applied to the other half (both contained 0.05% Tween 20). The leaves were then incubated for 6 h in a Petri dish with moistened filter paper in the base, and a small hole in the side to allow the petiole to pass through. The leaf remained attached to the plant during the incubation period. The relative humidity inside the dish rapidly rose to 98% and the droplets remained as a liquid for the entire time. As the movement of the foliar absorbed Zn in the leaf tissues was highly limited over 6 h (Li et al. 2017), we disregarded the amount of Zn translocation out of the leaves. After the 6 h incubation period, the leaf was cut in half along the mid-vein and rinsed separately using 2% HNO3, 3% ethanol, and deionised water to remove the Zn fertiliser remaining on the leaf surface (Vu et al. 2013).

The leaves were oven dried (60°C) and digested using a 5:1 mixture of nitric acid and perchloric acid. The Zn concentration was determined using ICP-MS and P concentration was determined using inductively coupled plasma optical emission spectroscopy (ICP-OES), with blanks and reference materials included to ensure accuracy. Four replicates were examined for each treatment. Zinc foliar absorption was calculated as Zn absorption (μg half leaf-1) = (elemental concentration in the treated half – elemental concentration in the control half) × dry weight of the treated half of the leaf. Data were analysed using GenStat version 18

189 and IBM SPSS version 24. Comparisons between means were made using the two-way or one-way analysis of variance with least significant difference (LSD) at 5%.

Examining Zn foliar absorption using µ-XRF Living plants that had been grown at 0 and 0.5 µM Zn for 40 d were transported to the XFM beamline of the Australian Synchrotron (Clayton, Australia) for in situ analysis of Zn distribution within YFELs using µ-XRF. The plants were grown under the same conditions as at The University of Queensland. Two droplets, each 5 µL, were applied next to each other on the adaxial surface of YFELs, with one droplet being ZnSO4 and the other being nano- ZnO. The leaves were immediately sealed in Petri dishes as described earlier and incubated for 3 h, during which time the leaves remained attached to the plant and the growth lamps remained switched on. Thereafter, the droplets on the leaf surface (which remained as a liquid and had not dried out) were blotted dry, with the leaves then cut off and rinsed as described earlier. The rinsed leaf was blotted dry and mounted on a sample holder between two layers of 4 µm Ultralene film to hold the sample in place and limit dehydration. The sample was then analysed, with scanning generally commencing within 5 min of the leaf being excised.

Details of the XFM beamline at the Australian Synchrotron have been provided by Paterson et al. (2011), with Kopittke et al. (2011) providing details for the analysis of plant tissues. In the present experiment, a beam of 15.8 keV X-rays was focussed to a spot of around 2 µm × 2 µm, with a photon flux of 2 × 109 photons s-1. The secondary X-rays (fluorescence and scattering) were collected using a silicon drift-diode detector (Vortex) located at 90° to the incident beam using a FalconX (XIA) digital signal processor. All elemental mapping was conducted with on-the-fly scanning in the horizontal direction (x-axis), with discrete steps in the vertical direction (y-axis). For the scans of the intact leaves, the step size of 25 µm was used with a horizontal stage velocity of 1.5 mm s-1, yielding a dwell of 16.7 ms pixel-1. All µ- XRF data was analysed using the CSIRO Dynamic Analysis method in GeoPIXE (http://www.nmp.csiro.au/dynamic.html) (Ryan and Jamieson 1993; Ryan 2000).

In addition to scanning intact leaves, we also scanned leaf cross sections of 40-d-old plants to obtain information regarding the distribution of Zn within the inner leaf tissues. For this purpose, we used cross sections from the area where the droplets had been applied, with the cross sections prepared in advance at The University of Queensland. Four samples were prepared, being cross sections from YFELs from Zn sufficient and deficient sunflower with 190

ZnSO4 applied for 6 h or nano-ZnO applied for 5 d. The Zn application method was the same as mentioned earlier, but 40 µl droplets were applied to allow for sections to be obtained from tissues entirely under Zn the droplets. After the appropriate application time, leaves were cut off and rinsed as discussed earlier. After rinsing, the area where Zn had been applied was excised using a razor blade and the leaf segment was embedded in a 4% agar solution at ca. 30 °C. Once the agar solidified, the block was glued to the sample holder of a vibratome (Leica VT 1000s) using super glue, then 150 µm sections were cut without any buffering solution. The sections were then carefully sealed in Ultralene film held onto XRF sample cups (SC-8047, Premier Lab Supply) and freeze dried after small holes were punctured in the film. The samples were then transported to Australian Synchrotron with the XRF sample cups. At the XFM beamline, the samples were removed from the sample cups and mounted on a sample holder between two layers of Ultralene film. For the µ-XRF analysis of the cross sections, the step size was 2 µm and the horizontal stage velocity was 0.4 mm s-1, yielding a dwell of 5 ms pixel-1.

Trichome and stomatal density, rough cuticle composition of the adaxial leaf surface, and contact angle of the Zn droplets Leaf trichome density and stomatal density were assessed for the adaxial surface of 40-d-old YFELs, using scanning electron microscopy (SEM) (Li et al. 2017). Specifically, the fresh leaves were removed from both treatments and immediately cut into 1 × 4 mm leaf segments, and fixed with 3% glutaraldehyde in 0.1 M sodium cacodylate using microwave processing

(Li et al. 2017). The samples were then post-fixed with 1% OsO4, dehydrated through a graded ethanol series, dried using a critical point dryer (Autosamdri-815 CPD, USA), and coated with Au (30 nm). The adaxial leaf surface of four replicates was examined using SEM (JEOL NEOSCOPE, Japan) at 10 kV accelerating voltage.

The gross composition of the adaxial surface of YFELs (40-d-old plants) was analysed using Fourier transform infrared (FTIR) spectroscopy as described in Li et al. (2017). Firstly, a scalpel blade was used to gently scrape the abaxial side of the fresh leaves that left the adaxial layer with some residual tissues. These were then placed in 2% pectinase and 0.1% cellulase enzyme solution in 50 mM sodium acetate at 25°C overnight to remove the residual cell debris. Next, they were washed thoroughly with deionized water. Using light microscopy (Olympus, SZX16), it was found that the trichomes were still on the partially digested leaf surface. After being air-dried, the isolated tissues (containing the cuticle and likely additional 191 epidermal cell wall material and trichomes) of Zn-deficient and Zn-sufficient sunflower plants were mixed evenly before being analysed by FTIR with the spectra then normalized.

The contact angles of Zn droplets with the adaxial leaf surface of Zn-sufficient and Zn- deficient leaves were assessed using the drop-shape analysis method (Woodward 1999). -1 Briefly, 5 µl droplets of 1000 mg Zn L ZnSO4.7H2O were placed on the adaxial surface of YFELs of Zn-sufficient and Zn-deficient (40-d-old plants). Images were captured immediately after the droplets were applied using a light microscope (Olympus, SZX16), with six replicates examined. The images were then analysed using ImageJ (available free-of- charge at https://imagej.nih.gov/ij/) for determining the contact angles.

Examining the speciation of Zn in bulk leaf tissues using XAS The chemical speciation of the absorbed Zn in the YFELs of 40-d-old sunflower was analysed in situ at the XAS beamline of the Australian Synchrotron. Four samples were examined, being leaves of Zn sufficient and Zn deficient sunflower to which ZnSO4 and nano-Zn had been applied for 6 h. To prepare these samples, 60 droplets (5 µL) of either

ZnSO4 or nano-ZnO were applied across the whole leaf, incubated in Petri dishes, and then rinsed as described earlier. There were four replicates of each treatment. The rinsed leaves were immediately frozen in liquid nitrogen before being transported to Australian Synchrotron in a dry shipper cooled with liquid nitrogen. At the XAS beamline, the leaves were ground together under liquid nitrogen using an agate mortar and pestle, mixed evenly and placed in a cooled sample holder, sealed with Kapton tape, and immediately transferred to a liquid helium cryostat (ca. 10 K) for analysis. At no time were the frozen (hydrated) samples allowed to thaw.

Ten standard Zn compounds were prepared as references. There were three solid compounds, being ZnO (Sigma Aldrich, 93632), Zn3(PO4)2 (Sigma Aldrich, 587583), and

[ZnCO3]2·[Zn(OH)2]3 (Sigma Aldrich, 96466). The three solid compounds were diluted using cellulose such that the final Zn concentration was 100 mg Zn kg-1. Another seven aqueous compounds were prepared, being Zn oxalate, Zn pectin, Zn citrate, Zn histidine, Zn(NO3)2,

Zn phytate, and Zn cysteine. These compounds were prepared by mixing of 4 mM Zn(NO3)2 with 20 mM of oxalate acid, pectic acid, citric acid, histidine, phytic acid, or cysteine. Where appropriate, pH was then adjusted to ca. 6 using 0.1 M NaOH. Modelling using GEOCHEM- EZ (Shaff et al. 2010) indicated that > 99% of Zn was complexed withu citric acid, oxalic 192 acid, and histidine, and > 96% with cysteine. Furthermore, for the Zn(NO3)2 standard, > 2+ 2+ 99.9% of the Zn was present as free Zn , thus, the Zn(NO3)2 standard is referred to as ‘Zn ’ hereafter. The seven aqueous compounds were analysed following mixing with glycerol to form a final solution with 30% glycerol (to limit ice-crystal formation during cooling).

The Zn K-edge X-ray absorption near edge structure (XANES) spectra were obtained in fluorescence mode using a 100-element solid-state Ge detector. Two replicate spectra were collected for each sample and standard. Each spectrum was calibrated by simultaneous measurement in transmission mode of a metallic Zn foil. The two replicates were merged and normalized using Athena (version 0.8.056) (Ravel and Newville 2005), and linear combination fitting (LCF) was performed with a maximum of three standards.

Results Effects of Zn deficiency on plant growth No visible differences were evident between the Zn sufficient and Zn deficient plants in the first month of growth in the nutrient solutions. Thereafter, the growth rate of the Zn deficient sunflower plants gradually decreased and the older leaves became chlorotic around the leaf margins (Supplementary Fig. S1a). After 40 d, the old leaves of the Zn deficient sunflower plants had died (Supplementary Fig. S1b) and the younger leaves developed a crinkled appearance and became chlorotic with necrotic margins (Supplementary Fig. S1c). The YFELs and the emerging leaves of the Zn deficient sunflower were noticeably smaller, although there were no other visible symptoms (Supplementary Fig. S1d).

Bulk foliar analysis of Zn absorption Bulk analyses of the foliar absorption of Zn were conducted using ICP-MS for plants that had been growing for 16, 28, and 40 d. Firstly, the leaf Zn concentration was analysed. It was found that the leaf Zn concentration in the Zn deficient plants was ca. 11 to 20 µg g-1 dry weight (Fig. 1a), which were about half of the values for the Zn sufficient plants (in a range from 33 to 39 µg g-1 dry weight). Hence, the leaf Zn concentrations recorded for Zn deficient plants are similar to the critical values reported for Zn deficiency in sunflower (13-20 µg g-1 dry weight ) (Reuter et al. 1997), confirming that plants growing at 0 µM Zn were indeed Zn deficient. The leaf Zn concentrations in the Zn deficient or sufficient plants did not differ significantly over time (Fig. 1a).

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Next, we investigated the foliar Zn absorption of the YFELs after applying droplets of a 15.4 mM ZnSO4 solution on the adaxial leaf surface for 6 h (Fig. 1b). The magnitude of absorption in the Zn deficient leaves was significantly lower than that in Zn sufficient leaves at both 28 and 40 d (Fig. 1b). For example, absorption for the Zn deficient sunflower was ca. 50 µg half leaf-1 (with only a slight change over time), while in the Zn sufficient sunflower, foliar Zn absorption increased from 70 µg half leaf-1 at 16 d to 120 µg half leaf-1 at 40 d (Fig. 1b).

Finally, we also compared absorption of ZnSO4 and nano-ZnO (6 h application) at 40 d growth. The absorption of ZnSO4 was 5-16 times higher than the absorption of nano-ZnO, with no differences being found between Zn deficient and sufficient sunflower when supplied with nano-ZnO (Fig. 1c).

Distribution of Zn in YFELs analysed using µ-XRF The µ-XRF scans of intact, hydrated YFELs showed marked differences in Zn foliar absorption rates, with differences between plant Zn status and between the forms of Zn fertiliser. From the µ-XRF scans, it was apparent that the addition of Zn droplets to the leaf surface markedly increased Zn concentrations in the underlying tissue, as indicated by the brighter colours which correspond to higher Zn concentrations (Fig. 2). Firstly, examining the background Zn concentrations away from the droplets, it was apparent that the Zn deficient leaf had lower background Zn concentrations than did the Zn sufficient leaf. Secondly, the Zn concentrations underneath where the droplets were applied were higher (i.e. brighter colours) for the Zn sufficient leaf than for the Zn deficient leaf, confirming that the foliar absorption of Zn was higher in the Zn sufficient sunflower than in Zn deficient sunflower. Thirdly, comparing the two forms of Zn, concentrations underneath where the droplets were applied were higher (i.e. brighter colours) for ZnSO4 than for the nano-ZnO, again confirming that the absorption of ZnSO4 was higher than for nano-ZnO. Finally, across both forms of Zn fertiliser, although Zn concentrations in the leaf tissue were lower for the Zn deficient leaf than for the Zn sufficient leaf, the pattern of the distribution of Zn was similar, with the Zn found almost entirely underneath the applied droplet and limited accumulation away from the site of application. It was also noted that there were small spots of ca. 100 µm that had the highest Zn concentrations. These corresponded to the non-glandular trichomes of sunflower in which foliar-applied Zn accumulated to high concentrations (Li et al. 2018).

Next, Zn distribution was examined in leaf cross sections that had been sampled from tissues entirely underlying the Zn-containing droplet (Fig. 3). As observed after treatment with both 194 forms of Zn to intact leaves, the foliar absorption of Zn was higher in the Zn sufficient plants than Zn deficient plants, as shown by the brighter colours in the Zn deficient leaves (Fig. 3).

Furthermore, the absorption of Zn from the ZnSO4 was again noted to be higher than that from the nano-ZnO, despite the ZnSO4 only being applied for 6 h and the nano-ZnO being applied for 5 d. The distribution of Zn across the leaf tissues was similar regardless of the foliar Zn treatment. Indeed, after 6 h application of ZnSO4, the highest Zn concentrations was in the trichomes on both the adaxial and abaxial leaf surfaces (Fig. 3).

Leaf trichome and stomatal density, leaf adaxial surface composition, and contact angle of the Zn droplets with the YFELs Leaf anatomical variables were measured to further investigate possible reasons for the differences observed in foliar absorption between the Zn sufficient and deficient sunflower. Firstly, trichome density was 14% lower and stomatal density was 17% lower for the Zn deficient YFELs than those with sufficient Zn (Fig. 4 and Fig. 5). The FTIR spectra of the isolated, adaxial surfaces of Zn sufficient and deficient sunflower leaves followed a similar pattern as shown in Fig. 6. Based on previous cuticular material assignment data (Fernández et al. 2014b; Heredia-Guerrero et al. 2014; Li et al. 2017), the following bands associated with different chemical functional groups and hence chemical constituents were identified for Zn deficient and sufficient adaxial cuticle layer (Fig. 6): (1) long-chain aliphatic waxes and -1 cutin (asymmetric and symmetric CH2 stretching absorptions at 2919 and 2849 cm and the -1 CH2 bending one at 1450 cm ); (2) the presence of cutin can also be ascribed to C-O-C stretching absorption at 1160 cm-1 and to the (OH) band at 1240 cm-1 which is also associated with polysaccharides; (3) a major presence of polysaccharides is derived from the high intensity band associated with the glycosydic bond at 1020 cm-1; (4) the occurrence of unsaturated/aromatic compounds is derived from the two strong bands at 1600 cm-1 and 1550 cm-1 are related to aromatic C=C and C-C functional groups; and (5) the broad and very strong band at 3350 cm-1, assigned to the O-H stretching vibration, may be related to tissue hydration. Despite the semi-quantitative character of FTIR analyses, comparison of the spectra obtained for isolated adaxial surfaces of Zn sufficient versus Zn deficient leaves suggests a higher presence of polysaccharides in Zn sufficient tissues and of phenolics, aromatic compounds and hydroxyl groups (possibly linked to cutin or other lipids) in Zn deficient leaves. Finally, we found no differences in the contact angles of the Zn droplets

195 between the Zn sufficient and deficient leaves, with both having values of ca. 60° (Supplementary Fig. S2).

Speciation of Zn in leaf tissues The chemical speciation of the Zn within hydrated leaf tissues was examined in situ at the XAS beamline using XANES. Firstly, 10 Zn standard compounds were examined (Fig. 7). For forms of Zn associated with carboxyl groups or present as free Zn2+, the peak was found to correspond to an energy of 9,670 eV (Fig. 7). In contrast, for Zn associated with P, spectra were observed to have a peak at 9,667 eV (Zn phosphate) or in the case of Zn phytate, at 9,669 eV (Fig. 7). When associated with S, Zn cysteine had a peak at 9,665.5 eV, while ZnO had a peak at ca. 9,671 eV and a shoulder at ca. 9,665 eV. In addition to shifts in the energy values corresponding to the white line peaks, there were also pronounced differences in heights of the main peaks (for example, compare Zn phosphate and Zn phytate) as well as differences in other spectral features among the various forms of Zn (Fig. 7).

Analysis of Zn speciation in frozen, hydrated leaf tissues revealed differences in the XANES spectra between the Zn sufficient and deficient sunflower (Fig. 7 and Supplementary Fig. S2). Although the XANES spectra for both the Zn sufficient and Zn deficient leaves exposed to

ZnSO4 had peaks at 9,669 eV, the peaks differed in their height, and the spectrum for the Zn deficient leaves was flatter and broader than for the Zn sufficient leaves (Fig. 7 and

Supplementary Fig. S3). Interestingly, however, a comparison of the two forms of Zn (ZnSO4 and nano-ZnO) revealed that the form of Zn supplied to the leaves did not appear to alter the speciation of Zn within the leaves, with the XANES spectra appearing visually similar regardless of the form of Zn supplied to the leaves (Fig. 7). In Zn sufficient sunflower (regardless of the form of Zn supplied), LCF predicted that 37-53% of the absorbed Zn was present as Zn phytate, 29-34% as Zn oxalate, and 18-29% as Zn histidine. However, for the Zn deficient sunflower, LCF predicted that Zn was present as Zn phosphate (55%), Zn pectin (24%), and Zn2+ (21%) (Table 1). The Zn deficient leaves to which nano-ZnO had been applied for 6 h could not be analysed using XAS as the Zn concentration in the leaf tissue was below the detection limit.

Discussion The main objective of this study was to characterise the absorption rate of foliar-applied Zn by Zn sufficient versus Zn deficient plants using sunflower as a model plant species. While 196 some studies have shown a lower foliar absorption rate of nutrient deficient plants, for example, P deficiency in wheat (Fernández et al., 2014a) and B deficiency in soybean (Will et al. 2011), this is the first study in which the rate of absorption, the absorption pattern, and speciation of Zn within the leaf tissues has been examined. The results suggest that the process of absorption of foliar applied ZnSO4 and ZnO nanoparticles is similar for Zn sufficient and Zn deficient leaves, with the Zn foliar penetration being quantitatively lower in Zn deficient plants and the speciation of the absorbed Zn in the leaves being different. The methods employed provide new insight into the mechanisms of foliar Zn absorption and transport and can be useful for improving our understanding of the mechanisms of absorption and bioactivity of foliar applied nutrients.

Foliar absorption of Zn is lower in Zn deficient sunflower Bulk analyses indicated that absorption of the foliar-applied Zn in Zn deficient sunflower was significantly lower than in Zn sufficient sunflower, being an average of 66% lower for 40 d old plants (Fig. 1). Similarly, using µ-XRF for the in situ analyses of leaves from plants that were 40 d old, we observed that Zn absorption was substantially lower for the Zn deficient leaves than for the Zn sufficient leaves (Fig. 2). This is in general agreement with previous studies, for example, Fernández et al. (2014a) observed decreased foliar absorption of P in P deficient wheat and Will et al. (2011) found decreased foliar absorption of B in B deficient soybean. Interestingly, however, the distribution of Zn was similar between Zn deficient and Zn sufficient leaves (Fig. 2 and Fig. 3), indicating that the absorption and accumulation of foliar-applied Zn follow similar mechanisms regardless of the plants’ Zn status.

Given the marked differences in absorption rates between the Zn sufficient and deficient leaves (which did not appear to be due to differences in the mechanisms of absorption), we examined whether Zn deficiency induced changes in leaf surface properties. For sunflower leaves treated with Zn-containing solutions, it was found that penetration via the trichomes and cuticle are the main foliar absorption pathways (Li et al. 2018). Therefore, the decreased trichome density in Zn deficient sunflower (ca. 15% lower than for the Zn sufficient leaves) could be a reason for the lower rate of Zn absorption in Zn deficient sunflower (Fig. 4 and Fig. 5).

We also examined the gross composition of adaxial leaf surface of Zn sufficient leaves compared to Zn deficient leaves, finding that the relative proportion of polysaccharides was 197 lower in Zn deficient tissues while the proportion of cutin and phenolics increased (Fig. 6). The plant cuticle has recently been proposed to be considered as a lipid-rich part of the epidermal cell, with chemical and structural changes potentially occurring in the cuticle of among. e.g., different species, organs, environmental conditions or plant physiological status (Fernández et al. 2016). However, it must be noted that the adaxial surface enzymatic digestion method employed was very mild compared to other studies [e.g., cuticle isolation in 2% cellulose and 5% pectinase solutions for 1 month (Guzmán et al., 2014)]. This was because the cuticle would break at higher enzyme concentrations as experienced in preliminary trials (data not shown). Hence, the isolated tissues in the present study contained the cuticle, trichomes and possibly, additional epidermal cell wall material. Several hypotheses can be suggested for the observed gross chemical composition differences. Firstly, the structural and chemical composition of the adaxial surface might be different between the Zn deficient and Zn sufficient leaves, and the Zn deficient tissues may have indeed contained a relatively lower proportion of cell wall material and a higher proportion of lipid impregnation compared to the healthy leaves. Secondly, given that trichomes were present in the isolated cuticle layers and Zn sufficient sunflower leaves had higher trichome density (Fig. 4A), the relative higher presence of polysaccharides in the Zn sufficient cuticles can also be attributed to the trichomes as they contain significant amount of cellulose (Fernández et al. 2011; Fernández et al. 2014b). Nevertheless, the reduced relative proportion of polysaccharides and the increased proportion of cutin and phenolics in the Zn deficient cuticles imply that the permeability of the Zn deficient cuticle might be decreased. However, the mechanisms of penetration of solutes through the cuticle, for example, the so-called ‘polar pathway’ (Schönherr 2006) are still not fully understood, although it has been related to cuticle hydration (Schreiber and Schönherr 2009) and the formation of the ‘dynamic aqueous continuum’ (under conditions favouring the deposition of water, for example, high air relative humidity, foliar spray, rain or fog) (Fernández et al. 2017). This would require the development of more detailed microscopic and chemical analyses.

Form of Zn applied to the leaf surface

It was found that the short-term absorption of nano-ZnO was substantially lower than ZnSO4 (Fig. 1C and Fig. 2). This is presumably due to the observation that nano-ZnO has a high molecular size and low solubility: a suspension of 1000 mg Zn L-1 results in a soluble Zn concentration of only 3 mg Zn L-1 (Li et al. 2018a). The process of cuticular penetration has been shown to be very much size limiting, with molecular diameters being restricted in the 198 range of 0.3 to 2.4 nm (e.g., Schönherr 1976; Popp et al. 2005; Schönherr 2006; Eichert and Goldbach 2008). Thereby, the 30 nm diameter size of nano-ZnO particles will restrict their rate of cuticular diffusion. Apart from the solubility, the point of deliquescence is also important for the process of foliar absorption (Fernández and Eichert 2009). A lower point of deliquescence will prolong the drying time of fertiliser drops and hence facilitate the penetration of the active ingredient for a longer period (Fernández et al. 2017). ZnSO4 has a high point of deliquescence of 90% (Hadrami 2011) and it seems that the point of deliquescence of nano-ZnO is also high, therefore limiting the significance of this parameter when examining the potential penetration mechanisms of both Zn sources. However, the use of nano-ZnO may be useful as a slow release form of Zn if it can be retained on the leaf surface, particularly given that more soluble forms of Zn often result in leaf scorching when foliar-applied (Drissi et al. 2015).

Why does the speciation of Zn change in Zn deficient leaves? Not only did the foliar absorption of Zn decrease in Zn deficient sunflower, but the speciation of Zn within the leaf tissues also changed (Fig. 7). Changes in the speciation of Zn within the leaf tissue following its absorption would potentially influence its subsequent translocation. In the present study, LCF suggests that around 37-53% of the Zn is associated with phytate in Zn sufficient leaves (Fig. 7 and Table 1). However, the XANES spectrum was much flatter and broader for the Zn deficient leaves (Fig. 7, Supplementary Fig. S2 and Table 1), with LCF indicating that 55% Zn was associated with phosphate in Zn deficient leaves. In this regard, it is known that there is an interaction between P and Zn within plants (Lei 2018). For example, it has been reported that high levels of available P can induce Zn deficiency (Soltangheisi et al. 2013) while Zn deficient plants can over-accumulate P in shoot tissues (Khan et al. 2014). Thus, in the present study, we also examined changes in bulk leaf P concentrations, finding that the tissue P concentrations were ca. twice as high in Zn deficient leaves than in Zn sufficient leaves (Supplementary Fig. S4). These results suggest that Zn deficiency resulted in increased accumulation of P within the leaf tissues (Supplementary Fig.S3) and the speciation of the P also changed (phytate in Zn sufficient leaves while phosphate in Zn deficient leaves) (Table 1), with this influencing the speciation of Zn after its absorption through the leaf surface (Fig. 7 and Table 1). However, the increased P concentration in the leaves of Zn deficient plant could potentially be associated with the decreased biomass of the Zn deficient plants, but this parameter was not measured in the present study and will have to be assessed in future trials. 199

In addition, it was also indicated that in leaves of Zn sufficient sunflower, 29-34% Zn was as Zn oxalate and 18-29% as Zn histidine, but in Zn deficient leaves 24% was present as Zn pectin and 21% as Zn2+ (Table 1). It is proposed that Zn could be transported through the phloem as Zn2+, Zn-nicotianamine (Nishiyama et al. 2012), Zn-malate, Zn-histidine complexes (Álvarez-Fernández et al. 2014; Gupta et al. 2016) or Zn-DMA (2’- Deoxymugineic acid) (Bashir et al. 2012). However, the form in which Zn is translocated within plants remains unclear.

Conclusions The Zn status of sunflower plants was found to influence the absorption of foliar-applied Zn. Specifically, absorption of Zn was significantly lower for Zn deficient sunflower than for Zn sufficient sunflower. Despite this difference, in situ analyses using µ-XRF showed that the pattern of Zn accumulation was similar in both Zn deficient and Zn sufficient leaves, suggesting that the difference in absorption was not due to differences in the mechanisms of foliar absorption. In addition to the decreased leaf trichome density, Zn deficiency also altered the chemical composition of the adaxial leaf surfaces which had relatively higher proportions of cutin, wax, and phenolics and a lower presence of polysaccharides. These chemical and structural changes in Zn deficient leaves presumably reduced the permeability of the cuticle to solutes, although further investigation is required in this regard. For example, it could potentially be worth investigating the effectiveness of using foliar fertilisation at plant early growth stages (i.e. before Zn deficiency influences leaf properties) to overcome Zn deficiency. Following the process of Zn foliar absorption, the speciation of Zn in the leaves was also affected by plant Zn status, with Zn phytate dominating in Zn sufficient leaves and Zn phosphate dominating in Zn deficient leaves. Finally, absorption of nano-ZnO was substantially lower than of ZnSO4, suggesting that nano fertilisers could potentially be used as a slow release foliar fertiliser provided they can be retained on the leaf surface during plant growth.

Acknowledgements The authors acknowledge use of the facilities and technical assistance of both the Queensland Brain Institute and the Australian Microscopy & Microanalysis Research Facility at the Centre for Microscopy and Microanalysis, The University of Queensland. Parts of the research were undertaken on the XFM and XAS beamlines at the Australian Synchrotron, 200 part of Australian Nuclear Science and Technology Organisation (ANSTO). This work was supported by Sonic Essentials, as well as by the Australian Research Council (ARC) through the Linkage Projects funding scheme (LP130100741). Support was also provided to Peter Kopittke by the ARC Future Fellowship funding scheme (FT120100277) and to Cui Li through the China Scholarship Council and The University of Queensland International Scholarship award.

Supplementary Information Supplementary Fig. S1 Comparison of Zn deficient (0 µM Zn) and sufficient sunflower (0.5 µM Zn). Supplementary Fig. S2 Contact angle of Zn droplets with the leaf surface. Supplementary Fig. S3 Detailed K-edge XANES spectra (from energy of 9,660eV to 9,680eV) of four standard Zn compounds and two sunflower leaf samples. Supplementary Fig. S4 Comparison of the leaf P concentration between Zn deficient (0 µM Zn) and sufficient sunflower (0.5 µM Zn).

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Figures

Fig. 1 Changes in (a) leaf Zn concentration and (b) the foliar Zn absorption (6 h ZnSO4.7H2O application) for Zn deficient (0 µM Zn) and sufficient (0.5 µM Zn) sunflower leaves over time. (c) Influence of the form of Zn on foliar absorption, with Zn supplied as either

ZnSO4.7H2O or nano-ZnO to adaixal leaves surface of Zn deficient and sufficient sunflower grown for 40 d. In (a) and (b), the LSD values either allow comparison between the leaves differing in Zn status (‘Zn’) or differing in period of growth (‘time’). D.W. = dry weight. In (c), the different letters represent significant differences (P < 0.05).

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Fig. 2 µ-XRF scans showing the distribution of Zn in hydrated leaves of Zn sufficient (top,

0.5 µM Zn) and deficient sunflower (bottom, 0 µM Zn) to which either ZnSO4 or nano-ZnO had been applied for 3 h. All images are shown using the same concentration scale (and hence images are comparable), with brighter colours corresponding to higher Zn concentrations.

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Fig. 3 µ-XRF scans showing the distribution of Zn in cross sections of Zn sufficient (0.5 µM

Zn) and deficient (0 µM Zn) sunflower leaves to which either ZnSO4 or nano-ZnO droplets had been applied. The ZnSO4 was applied for 6 h and the nano-ZnO applied for 5 d. All cross sections were taken from tissues entirely underlying droplets. All images are shown using the same concentration scale, with brighter colours corresponding to higher Zn concentrations.

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Fig. 4 Changes in (a) leaf trichome density and (b) stomatal density, in Zn sufficient (0.5 µM Zn) and Zn deficient (0 µM Zn) sunflower. Data are the arithmetic mean of four replicates, with standard deviations shown. Different letters represent significant differences (P < 0.05).

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Fig. 5 Scanning electron micrographs showing trichomes (a and b) and stomata (c and d) on the adaxial leaf surface of Zn sufficient (0.5 µM Zn) and deficient (0 µM Zn) sunflower. In (e) and (f), light micrographs of unstained leaf cross sections (80 µm thick) of Zn sufficient and deficient sunflower. The scales on the left images apply also to the right images on their right.

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Fig. 6 Spectra from Fourier transform infrared spectroscopy (FTIR) obtained from isolated cuticles from the adaxial leaf surfaces of Zn sufficient (0.5 µM Zn, green) and deficient (0 µM Zn, blue) sunflower.

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Fig. 7 Normalized K-edge XANES spectra of 10 standard Zn compounds (upper colour lines) and the three sunflower leaf samples (bottom black lines). For the sunflower leaf samples, the black lines corresponding to XANES spectra and the dotted red lines corresponding to the results of the linear combination fitting (LCF).

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Table 1 Results of linear combination fitting (LCF) of K-edge XANES data for sunflower leaves to which Zn had been applied for 6 h.

0.5 µM Zn 6 h 0 µM Zn 6 h Standards (%) ZnSO4 Nano-ZnO ZnSO4 Nano-ZnO Phytate 37 ± 2 53 ± 4 Oxalate 34 ± 1 29 ± 5 Histidine 29 ± 1 18 ± 3 Pectin 24 ± 2 Phosphate 55 ± 0.8 Zn2+ 21 ± 2 R factor a 0.0002 0.001 0.0002 NA a R factor = ∑(experimental-fit)2/∑(experimental)2, where the sums are over the data points in the fitting region.

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Supplementary Information

Absorption of foliar applied Zn is decreased in Zn deficient sunflower (Helianthus annuus) due to changes in leaf properties

Cui Li1*, Peng Wang1, 2, Enzo Lombi3, Jingtao Wu1, F. Pax C. Blamey1, Victoria Fernández4, Daryl L. Howard5, Neal W. Menzies1, Peter M. Kopittke1

1 The University of Queensland, School of Agriculture and Food Sciences, St Lucia, Queensland, 4072, Australia 2 Nanjing Agricultural University, College of Resources and Environmental Sciences, Nanjing, 210095, China 3 University of South Australia, Future Industries Institute, Mawson Lakes, South Australia, 5095, Australia 4 Technical University of Madrid, School of Forest Engineering, Forest Genetics and Ecophysiology Research Group, Madrid, 28040, Spain 5 Australian Synchrotron, Clayton, Victoria, 3168, Australia

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Supplementary Fig. S1 Comparison of the Zn deficient (0 µM Zn) and Zn sufficient (0.5 µM Zn) sunflower. (a) Plants that are 33 d old, with the old leaves of Zn deficient sunflower displaying chlorotic and necrotic symptoms. (b) Plants that are 40 d old, with the Zn deficient plant being smaller and showing chlorosis and necrosis in their old leaves. (c) Older leaves (third from the base of the plant) from plants that are 40 d old. (d) The youngest (upper) leaves of plants that are 40 d old, with the leaves of Zn deficient plants being smaller in size with but no other visual symptoms.

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Supplementary Fig. S2 Light micrographs showing the contact angle of Zn droplets (5 µl of ZnSO4.7H2O with 0.05 % Tween 20) applied to leaves of Zn sufficient (0.5 µM Zn) and Zn deficient (0 µM Zn) sunflower. Photos were taken immediately after the droplets had been applied.

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Supplementary Fig. S3 Detailed K-edge XANES spectra from 9,660 eV to 9,680 eV of four standard Zn compounds and two sunflower leaf samples, showing the differences between Zn sufficient (0.5 µM Zn) and Zn deficient (0 µM Zn) sunflower leaves to which ZnSO4 had been applied for 6 h.

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Supplementary Fig. S4 Comparison of the leaf P concentration between Zn deficient (0 µM

Zn) and sufficient (0.5 µM Zn) sunflower, with the LSDZn (5%) value allowing comparison between the Zn deficient and sufficient leaves at any given value of time.

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Chapter 7 General discussion and conclusions

Zinc is an essential element for both plants and humans. However, the presence of inadequate levels of Zn in soils is a widespread problem, resulting in crop Zn deficiency around the world (Cakmak, 2000, Alloway, 2008,). As a result, not only is the growth of the crop reduced, but also the quality of the agricultural commodities is impaired. Consequently, one third of the world’s human population suffers from Zn deficiency (Cakmak et al., 2010). Therefore, growing plants that produce food with sufficient levels of Zn is of great significance for both improving crop production and improving human health.

Deficiencies of Zn occur not only due to the level of Zn in soil being low, but it also occurs when soil properties reduce its availability, such as in soils with high pH and high carbonate content. In these soils, addition of Zn fertilizer to the soil is ineffective, for example, due to the precipitation of Zn(OH)2 (Fernández et al., 2013a, Prasad et al., 2014). Foliar Zn fertilisation has been shown to be an efficient approach to overcome this problem (Velu et al., 2014, Joy et al., 2015, Zhang et al., 2018). However, much remains unknown regarding the absorption and translocation of the foliar applied Zn. Using ZnSO4 as a traditional soluble Zn fertilizer and comparing with the sparingly soluble nano-ZnO, the absorption and translocation of foliar applied Zn was examined in soybean, sunflower, and tomato.

7.1 Absorption of foliar applied Zn

7.1.1 The cuticle is an important pathway for foliar Zn absorption In Chapter 3, MeJA was used to induce changes in leaf properties of soybean, sunflower, and tomato. These changes were then related to changes in the foliar absorption of Zn, Mn, and Fe in order to identify which leaf properties influence foliar nutrient absorption. It was found that after treatment with 1 mM MeJA, the absorption of the foliar applied Zn, Mn, and Fe all increased in sunflower, but decreased in tomato, and did not differ in soybean. Interestingly, examination of the leaf properties showed that the thickness of the cuticle and the epidermal cell wall decreased in sunflower but increased in tomato, and did not differ in soybean (Paper 2). These results suggest that the thickness of the cuticle and the epidermal cell wall is important for foliar nutrient absorption in these three plant species.

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The importance of the cuticle was confirmed in Chapters 4 and 5. Using synchrotron-based µ-XRF to examine the distribution of the absorbed Zn in the leaf tissues of soybean, sunflower, and tomato, it was found that Zn accumulated in the leaf cuticular areas even after only 15 min, with the Zn concentration in the cuticular area increasing with the application time (Paper 3 and 4). In addition, in Chapter 5, using NanoSIMS, it was found that Zn appeared to cross the cuticle and accumulate in the epidermal cell walls.

In Chapter 6, the foliar absorption in Zn deficient and Zn sufficient sunflower plantswas compared. It was found that Zn foliar absorption decreased in Zn deficient sunflower. The data suggest that the alteration of the cuticle composition and structure was one of the reasons for the decreased foliar Zn absorption in Zn deficient plants. Specifically, Zn deficiency increased the proportions of cutin, wax, and phenolics in the cuticle while decreased the proportions of polysaccharides (Paper 5). This suggests that the cuticle structure and composition are related to foliar Zn absorption, and also confirmed that uptake through the cuticle is an important pathway for absorption of foliar applied Zn.

Overall, all five experiments support the notion that the cuticle is an important pathway for foliar Zn absorption, with the thickness, chemical composition, and structure of the cuticle altering foliar Zn absorption. Specifically, thicker cuticles tend to reduce foliar Zn absorption, while higher proportions of cutin, wax, and phenolics and lower polysaccharides tend to reduce foliar Zn absorption. Therefore, when designing foliar fertilisers, the addition of a suitable surfactant will likely increase the absorption rate by increasing the hydrophilcity of the nutrient solution. In this regard, different types of foliar Zn fertilisers could potentially be designed depending upon the properties of the plant cuticle. For example, for a plant species that has a thinner leaf cuticle, a lower ionic concentration may be considered. However, the physical structure and chemical heterogeneity of the cuticle are not fully understood (Yeats et al., 2013b, Guzmán et al., 2014b, Fernández et al., 2017), and the mechanisms by which nutrient penetration occurs remain unresolved (Fernández et al., 2017).

7.1.2 Some trichomes could be important for foliar Zn absorption In Chapter 4, consideration was particularly given to the role of trichomes in foliar Zn absorption. Two type of trichomes were identified on the leaf surface of soybean and seven types for tomato. µ-XRF was used for in situ analysis of the distribution of the foliar

220 absorbed Zn was examined after 15 min, 30 min, 1 h, 3 h, and 6 h. Other than the observation that Zn accumulated in the cuticular areas, no trichomes of tomato were found to accumulate Zn, while for soybean it was found that about 29 % of the non-glandular trichomes accumulated Zn. Next, bulk foliar Zn absorption was compared in four soybean NILs that differed up to 10-fold in trichome density. However, it was found that the differences in foliar Zn absorption were not related to the leaf trichome densities, but rather, were related to the different cuticle properties of the NILs (Paper 3). Therefore, the data demonstrate that the trichomes are not a primary pathway for foliar absorption of Zn in tomato and soybean. However, the accumulation of Zn in some of the trichomes of soybean is interesting, and possibly occurred due to a higher wettability at the trichome bases (Brewer et al., 1991, Grammatikopoulos et al., 1994, Fernández et al., 2014b). However, trichomes are highly diverse, and it is possible that trichomes are important for foliar Zn absorption in some species. For example, it was found in cactus (Ju et al., 2012) and Croton (Vitarelli et al., 2016) that trichomes were involved in leaf water absorption.

To explore more of this issue, sunflower was used as model plant, as it is known to accumulate metals in its trichomes when supplied in the rooting environment. In Chapter 5, the distribution of Zn in intact hydrated leaves was investigated using µ-XRF. It was found that Zn accumulated rapidly in the non-glandular trichomes even after only 15 min. Indeed, the highest Zn concentrations within the leaves was always found in the non-glandular trichomes, with this also being confirmed by examining leaf cross sections. It was estimated that after 6 h, the total Zn that accumulated in the NGTs was ca. 1.9 times higher than in the cuticular tissues. Hence, the non-glandular trichomes of sunflower were found to be important for foliar Zn absorption (Paper 4). The possible reasons are that the cuticle above this trichome was different and had a higher wettability (Bahamonde et al., 2018), or because of the special structure and function of this trichome, for example, a previous investigation reported that high concentrations of Mn can accumulate in this trichome (Blamey et al., 2015). Therefore, depending upon the plant species and the climatic and ecological conditions, some trichomes could be important for foliar Zn absorption. However, whether the Zn in the trichome bases can be translocated to other parts of the plant and whether the Zn was directly absorbed through the trichome bases themselves require further testing.

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7.1.3 Stomata appear to not be an important pathway for foliar Zn absorption The role of stomata for the foliar absorption of Zn was considered in Chapter 5. Using NanoSIMS, the distribution of foliar absorbed Zn was examined at a subcellular scale in leaf cross sections of sunflower. The Zn concentration around the stomata was not higher than the surrounding cuticular areas, suggesting that the movement of Zn through the stomata does not appear to be an important pathway for foliar Zn absorption (Paper 4). However, it must be noted that the analysis in the present study was considered a leaf cross section rather than as a three-dimensional analysis, and further research is required in this area. It has previously been proposed that the absorption of foliar applied nutrients through stomata is limited by the geometry and hydrophobility of the stomata (Burkhardt, 2010), and that wetting of the stomata is a necessary process for foliar nutrients to enter the leaf through stomata, with the properties of the applied chemical and the stomata itself (e.g. the cuticle above the stomata) influencing this process (Burkhardt et al., 2012, Eichert et al., 2012b).

Finally, although Zn foliar absorption through the abaxial leaf side was not considered in the present study, a comparison of the absorption of foliar applied nutrients through the abaxial and adaxial leaf surfaces has been examined previously, with these previous studies generally finding that the abaxial surface has a higher absorption rate than the adaxial surface. For example, the absorption of foliar applied P (in wheat) (Peirce et al., 2014), Zn (in tomato) (Du et al., 2015), and Na (in apple) (Burkhardt et al., 2012) was higher in the adaxial leaf surface than from the abaxial leaf surface, due to the adaxial side had higher density of stomata or trichome. Based on the results from the present study, the differences observed regarding the foliar nutrient absorption could also related to differences in the composition and thickness of the cuticle on the adaxial and abaxial leaf surfaces, with further study required in this regard.

7.1.4 Foliar Zn absorption is decreased in Zn deficient sunflower In Chapter 6, Zn foliar absorption was compared in Zn sufficient and deficient sunflower using µ-XRF, XAS, and ICP-MS. Surprisingly, foliar Zn absorption was decreased in Zn deficient sunflower. However, using µ-XRF, it was found that the Zn absorption pattern was similar between Zn deficient and Zn sufficient sunflower. Examination of the leaf properties revealed that the decreased absorption of Zn in the deficient plant possibly occurred due to a decrease in the leaf trichome density and an altered adaxial leaf surface composition and

222 structure. In addition, the interaction of Zn and P should be considered, as it was found that Zn deficiency increased root P uptake and affected the speciation of the absorbed Zn (Paper 5). Given that the gap of the foliar Zn absorption between the Zn deficient and sufficient sunflower increased over time, Zn fertilisation should be applied early to have a higher agronomic efficiency.

Finally, consideration should be given to the concentration of the Zn solutions used in the present study. Although foliar fertilisers have been used for more than 100 years (Alexander, 1986, Fernández et al., 2009, Fernández et al., 2013), there is still no agreement on the best approaches for the use of these fertilisers, including their concentration. In this study, 1000 -1 mg Zn L (either ZnSO4 or nano-ZnO) was used to investigate the process of foliar Zn absorption. The concentration of 1000 mg Zn L-1 is in the range of those used previously, including 40 mg Zn L-1 (ZnEDTA) was used in cotton (Sawan et al., 2001), 1000 mg Zn L-1 -1 (ZnSO4 and ZnEDTA) (Doolette et al., 2018) and 2000 to 5000 mg Zn L (ZnSO4) in wheat (Zhang et al., 2012). Furthermore, Zn concentrations of commercial foliar Zn fertilizer products are as high as 1500 mg Zn L-1 (Doolette et al. 2018). It has also been shown that absorption of foliar-applied Zn increases linearly with the solution Zn concentration (Du, 2014) although very high Zn concentrations can result in leaf scorch (Doolette et al. 2018). In practice, the concentration of the foliar Zn fertiliser solutions should be decided giving consideration to the solubility and the point of delinquency of the chemical, leaf properties (e.g. young or mature, hydrophilicity, etc.), and environmental conditions (especially humidity and temperature).

7.2 Comparison of the foliar absorption of ZnSO4 and nano-ZnO

In Chapters 4, 5, and 6, foliar absorption of ZnSO4 and nano-ZnO were compared in soybean and sunflower using µ-XRF, XAS, and ICP-MS. In Chapter 4, to test whether nano-ZnO (~30 nm diameter) was absorbed as nano particles, a homologous bulk-ZnO (~200–400 nm diameter) which ca. 10 times larger in size was also included in the comparison. In addition, to test whether nano-ZnO was absorbed as its dissolution products, the solubility of the nano- and bulk-ZnO was tested and found to be approximately 3‰. Therefore, 1000 mg Zn L-1 in -1 the form of ZnSO4, nano-ZnO, and bulk- ZnO, and a 3 mg Zn L as ZnSO4 (equal to the soluble Zn in 1000 mg Zn L-1 nano- and bulk-ZnO) were applied to leaves of soybean and the foliar absorption was measured using ICP-MS. After 6 h of application, the absorption of

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-1 1000 mg Zn L ZnSO4 was 10 times higher than from the nano- and bulk-ZnO. Furthermore, the absorption of Zn did not differ between the nano- and bulk-ZnO. These results suggest that the ZnO was unlikely absorbed as intact particles given the same results from products that differed in size particles by a factor of 10. Interestingly, however, the absorption of Zn -1 from 3 mg Zn L ZnSO4 was 6 times lower than its absorption from the nano- and bulk-ZnO. This is likely due to the ongoing dissolution of the ZnO while on the leaf surface (Paper 3).

In Chapter 5, foliar absorption of ZnSO4 and nano-ZnO were compared in sunflower using both ICP-MS and µ-XRF. It was found that the absorption of nano-ZnO was substantially lower than that of ZnSO4 after exposure for either 3 h or 3 d. However, regardless of the form in which the Zn was supplied, the distribution of the Zn within the underling leaf was similar. In addition, examining the chemical speciation of the absorbed Zn using XAS, it was found that the speciation of the Zn was similar within the leaf tissues, regardless of whether the Zn was supplied as ZnSO4 or as nano-ZnO. This further suggests that nano-ZnO and ZnSO4 were both absorbed in the same form, namely, as soluble Zn2+ rather than as intact nanoparticles (Paper 4).

In Chapter 6, the foliar absorption of ZnSO4 and nano-ZnO was compared in Zn sufficient and deficient sunflower. Again, it was found that foliar absorption of nano-ZnO was substantially lower than for ZnSO4 regardless of the Zn status of the plant itself (Paper 5).

Overall, the foliar absorption of nano-ZnO was substantially lower when compared with

ZnSO4. Although the agricultural (i.e. field) efficiency of the nano-ZnO as a foliar Zn fertiliser was not considered in the present study, previous studies in the field have found that the use of nano foliar fertilisers have increased the yield of pomegranate (Punica granatum cv. Ardestani) (Davarpanah et al., 2016) and snap beans (Phaseolus vulgaris L.) (Morsy et al., 2017). Therefore, this low absorption rate could potentially be advantageous, not only in avoiding the ‘burning’ that can occur with soluble forms (Drissi et al., 2015), but also in providing a slow release fertiliser if the ZnO nanoparticles can be retained on the leaf surface.

7.3 Translocation of the foliar absorbed Zn

To analyse the translocation of the foliar absorbed Zn, the distribution of Zn was analysed using µ-XRF (Chapters 3, 4, 5, and 6). Regardless of the form in which the Zn was supplied

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(ZnSO4 or nano-ZnO) and regardless of the plant species (soybean, sunflower, and tomato), the translocation of the majority of the foliar absorbed Zn was highly limited (< 1 mm after 6 h). The extent of translocation was slightly higher in the veins than in the interveinal tissues (Paper 2). In addition, in Chapter 6, it was found that the translocation of the absorbed Zn did not differ in Zn deficient and Zn sufficient sunflower plants (Paper 5).

Although lateral movement away from the site of application was low, in Chapters 4, 5, and 6, examination of the Zn distribution using leaf cross sections of soybean and sunflower, it was found that the foliar absorbed Zn did not accumulate solely on the adaixal leaf surface (i.e. the site of Zn application), but rather, the foliar absorbed Zn also accumulated at the abaxial leaf surface. Particularly, in sunflower, it appears that the Zn was rapidly translocated to the abaxial leaf surface from the adaxial surface, with the bundle sheaths extensions likely facilitating this process (Paper 4). Finally, in Chapter 5, analysis using NanoSIMS found that Zn seems translocated to the other leaf side through the apoplastic pathway through the veinal tissues (Paper 4). Nevertheless, this study has examined the translocation of Zn within the leaves in only comparatively short time periods (mostly 6 h) through examining the distribution of Zn mainly using µ-XRF, with µ-XRF being valuable for examining the distribution of Zn in situ in hydrated leaves. However, µ-XRF is not able to differentiate the background Zn from the foliar-absorbed Zn, and it did not directly show the Zn movement within the leaf tissues. Therefore, in the future studies, the translocation of the foliar absorbed Zn should be considered in the whole plant, in a longer time period (could consider to associate with the plant growth stages), and using other techniques such as isotopic tracing.

7.4 Conclusions

(1) Movement through the leaf cuticle seems to be an important pathway for absorption of the foliar absorbed Zn. The thickness of the cuticle and its chemical composition influence the magnitude of absorption. Some trichomes are important for the absorption of foliar applied Zn, depending upon the plant species and the climatic and ecological conditions. The movement of Zn through the stomata does not appear to be an important pathway for foliar Zn absorption. (2) It was found that the foliar absorption of the sparingly soluble nano-ZnO was

substantially lower than when supplied as ZnSO4. However, their absorption pattern was similar, as was the speciation of Zn within the leaf tissues, thereby suggesting

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that the nano-ZnO had been absorbed as soluble Zn2+, rather than as intact nano particles. Thus, nano-ZnO is potentially a valuable slow release form of Zn, provided it can be retained on the leaf surface. (3) In a short time period, the majority of the foliar absorbed Zn had a limited translocation in the leaf tissues. However, Zn was observed to move rapidly from the adaxial surface to the abaxial surface, perhaps through the bundle sheath extensions. (4) Absorption of foliar applied Zn was lower in Zn deficient sunflower than in Zn sufficient plants, with this being due to changes in leaf properties including decreasing in leaf trichome density and changes in cuticle composition.

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