Zein Nanoparticles Uptake and Translocation in Hydroponically

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Zein Nanoparticles Uptake and Translocation in Hydroponically Grown Sugarcane Plants Alisha Prasad, Carlos E Astete, Andreea Elena Bodoki, McKenzie Windham, Ede Bodoki, and Cristina M. Sabliov J. Agric. Food Chem., Just Accepted Manuscript • DOI: 10.1021/acs.jafc.7b02487 • Publication Date (Web): 02 Aug 2017 Downloaded from http://pubs.acs.org on August 3, 2017

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Journal of Agricultural and Food Chemistry

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Zein Nanoparticles Uptake and Translocation in Hydroponically

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Grown Sugarcane Plants

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Alisha Prasada, Carlos E. Astetea, Andreea E. Bodokib, McKenzie Windhama, Ede Bodokic*, and

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Cristina. M. Sabliova*

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a

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Agricultural and Mechanical College, and LSU AgCenter, Baton Rouge, LA 70803, USA

Department of Biological and Agricultural Engineering, Louisiana State University

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b

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University of Medicine and Pharmacy, 12, Ion Creanga St., 400010, Cluj-Napoca, Romania

General and Inorganic Chemistry Department, Faculty of Pharmacy, “Iuliu Hatieganu”

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c

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Medicine and Pharmacy, 4, Louis Pasteur St., 400349, Cluj-Napoca, Romania

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* Co-corresponding authors

Analytical Chemistry Department, Faculty of Pharmacy, “Iuliu Hatieganu” University of

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Key words: Zein nanoparticles, sugarcane, translocation, nanopesticides, zein-FITC

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Abstract

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The main objective of this study was to investigate the uptake and translocation of positively

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charged zein nanoparticles (ZNPs) in hydroponically grown sugarcane plants. Fluorescent ZNPs

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(spherical and measuring an average diameter 135±3 nm) were synthesized by emulsion-

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diffusion method from FITC-tagged zein. Fluorescent measurement following digestion of plant

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tissue indicated that sugarcane roots had a significant adhesion of ZNPs, 342.5±24.2 µg

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NPs/mg of dry matter, while sugarcane leaves contained a very limited amount, 12.9±1.2 µg

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NPs/mg dry matter for high dose after 12 hrs. Confocal microscopy studies confirmed

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presence of fluorescent ZNPs in the epidermis and endodermis of the root system. Given their

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ability to adhere to roots for extended periods of time, ZNPs are proposed as effective delivery

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systems for agrochemicals to sugarcane plants, but more studies are needed to identify effect of

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nanoparticle exposure to health of the plant.

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Until recently, nanomaterials were developed nearly exclusively for biomedical,

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electronics, and energy fields applications.1 As a result of significant achievements in

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these fields, research in the agricultural sector has begun to eagerly approach

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nanoparticles as pesticide vectors, while investigating their impact on the environment.2,3

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Current methods for administering pesticides such as manual backpack type spraying,

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self-propelled row-crop spraying, aerosol spraying etc. have several limitations most

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critically including the direct human and environment exposure, and uneven distribution

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of pesticides. Despite toxicity concerns, pesticides might undergo microbial degradation,

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degradation by sun exposure, and hydrolysis, such that a very low percentage of the

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sprayed chemical is delivered to the plant and insect.4,5 It has been reported that more

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than 90% of the conventional agrochemicals are lost and never reach the desired target

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due to application or environmental conditions. To maintain the agrochemical

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concentration over the minimum effective concentration, periodical applications are

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required. The process increases the cost, and excess of agrochemicals end up in the soil

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and water with negative environmental impacts.6,7

INTRODUCTION

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Nanomaterials offer a safer and more efficient method for pesticide delivery.8

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Pesticide concentration can be decreased by delivering it with engineered bio-adhesive


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nanodelivery systems and by decreasing degradation of the nano-entrapped pesticide

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during application.9,10,11 Some relevant examples of nanoparticles studied for

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agrochemical delivery are polymeric nanoparticles synthesized out of sodium alginate,10

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Eudragit S100,11 chitosan,12 poly(lactic acid),13 as well as solid lipid nanoparticles.14

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These nanoparticles showed promising results for agrochemical delivery relative to

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traditional formulations, but limited data about polymeric nanoparticle interaction with

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plants is available in the literature.

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Current research on nanoparticle-plant interaction has focused mainly on metallic

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nanoparticles such as TiO2, ZnO2, fullerenes, quantum dots, gold, silver, copper and iron

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oxide, showing evidence of nanomaterial uptake by plants through seeds, roots and

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leaves.15,16,17,18,19 Several factors affect uptake and translocation of metallic nanomaterials

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such as particle’s size, shape, dissolution rate, agglomeration state, surface chemistry,20

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but equally important, plant species has an enormous impact on whether or not the

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nanoparticle will translocate from the root system into the stems and leaves.21

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It has been established that metallic nanoparticles enter a plant through the roots.22 The

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soil is a place in which nanoparticles are likely to accumulate as a result of run-off,

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making the roots a potential primary organ of exposure, while some nanoparticles can be

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applied directly to the leaves by broadcasting.23 Through the roots, plants interact strongly

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with the environment, absorbing mainly water and nutrients, but also contaminants and

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other materials. Absorption of nanoparticles through the root is controlled based mainly

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on nanoparticle size, given the pore diameter of the root of 5 to 20 nm.1 The macro-

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structure of the root is responsible for the extent of nanoparticle absorption: plants with

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more lateral roots provide additional surface area for more absorption.24 Nanomaterials


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properties may also modulate interaction with the roots because of the possibility of

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mechanical adhesion, electrostatic adsorption, and hydrophobic affinity, which has been

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responsible for accumulation on the surface and in the epidermis of the root.25 Part of the

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reason nanoparticles are thought to adhere to the root surface is because of the release of

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exudates and mucilage, which is made up of amino acids and organic acids providing an

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overall negative surface charge.26, 27

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A subject of keen interest is nanoparticle translocation, which refers to the particle

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moving from the roots into the shoots. After the nanoparticles diffuse through the cell

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wall pores of the roots, there are two paths they can follow to move from the epidermis to

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the endodermis: the apoplastic and symplastic route, similar to ways in which water and

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nutrients are moved through the plant.28 To the authors’ knowledge, there is little-to-no

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literature available on the uptake and translocation of polymeric nanoparticles (NPs) in

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plants. The aim of this research was to study root nanoparticle uptake and translocation in

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sugarcane plants exposed to NPs hydroponically.

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The selected polymer for nanoparticle synthesis was zein, a protein derived from

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Zea mays L (corn). Zein is water insoluble, amphiphilic, renewable, and biodegradable29

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and hence preferred for agrochemical delivery. Zein nanoparticles (ZNPs) have shown

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potential applications as carrier vehicle for delivery of drugs,8,30 gene delivery,31 as well

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as surface coatings,32 etc. Various geometries of ZNPs have been explored such as

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spheres, hexagonal, sponge, lamellar phases, and nano-tubes etc. by tailoring the

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microphase behavior of zein based on ethanol: water ratio.33 This is possible since zein

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contains over 50% hydrophobic regions, and has the ability to form nanoparticles of

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specific surface chemistry due to its unique amino acid distribution.34 Various methods of


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ZNPs synthesis, their stability, and surface properties have already been well established.

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The main objective of this study was to synthesize ZNPs via emulsion-diffusion method

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using didodecyldimethylammonium bromide (ddMab) as a positively charged stabilizing

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surfactant and particles were fluorescently tagged to allow for qualitative and quantitative

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tracking in the plant. Fluorescence microscopy, fluorescence measurement, and

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transmission microscopy were employed to quantitatively and qualitatively assess

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nanoparticle association with sugarcane roots and their translocation to the leaves of the

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plant. The understanding of the ZNPs uptake, translocation, and distribution in plants

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would help in the development of zein nanoparticles as a potent carrier vehicle for

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delivery of agrochemicals with a broader impact both in food and agricultural

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applications. This work will advance the fields of controlled release materials, plant

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pathology, materials science/engineering, and agricultural chemistry.

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Materials. Zein powder, fluorescein isothiocyanate (FITC), dichloromethane (DCM), and

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didodecyldimethylammonium bromide (ddMab) were purchased from Sigma-Aldrich

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(Sigma

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dimethylformamide (DMF), triethylamine and acetone was purchased from Fisher

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Chemical (Fisher Scientific International, Fairlawn, NJ). Ultrapure (type 1, 18.2 MΩ)

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water was obtained using a NANOpure Diamond™ machine (Barnstead International, IA,

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USA). Hoagland medium was purchased from plant media bioWORLD (bioWORLD,

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Ohio, USA). Sugarcane plants (HoCP09-804 variety) were provided by Louisiana State

MATERIALS AND METHODS

Chemical

Co.

Ltd.,

St.

Louis,

MO).

Sodium


hydroxide

(NaOH),

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University’s Agricultural Center and collected from Sugar Research Station. All other

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chemicals used were of laboratory grade.

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Fluorescent tagging of zein via FITC. In order to track the ZNPs through the

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roots and leaves, prior to preparation of ZNPs, zein was conjugated with a fluorophore

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(FITC). Accurately weighed zein powder (3 g) was dissolved in 100 ml DMF, in a 250 ml

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round bottom flask completely covered with aluminum foil to avoid light exposure. Next,

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0.1 mL of triethylamine and 80 mg of FITC were slowly added under gentle stirring at

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room temperature. Stirring was stopped after 24 hours and the sample was concentrated

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(approximately 25 ml) under high vacuum conditions. Subsequently, the concentrated

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suspension was washed and precipitated with DCM (50 ml) and the cycle was performed

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for six times to ensure free dye removal. Finally, the precipitate was dried in an oven

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(40°C) under high vacuum for 24 hours and kept at 4oC in an amber container until

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further use. The absence of free FITC residue was further confirmed by HPLC analysis.

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ZNPs preparation. Emulsion-diffusion method was used to prepare fluorescent

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ZNPs. The organic phase was prepared by dissolving equal amounts of zein powder (200

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mg) and zein with covalently tagged FITC (zein-FITC, 200 mg) in 10 ml of an acetone:

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water solution (8:2, v/v) and kept under rigorous stirring in a closed container covered

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with aluminum foil until complete dissolution. The aqueous phase was prepared by

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mixing ddMab (57.8 mg) in 100 ml of water under stirring settings. Finally, the emulsion

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was prepared by adding the organic phase drop wise into the aqueous phase under

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rigorous stirring, and passing the resulting emulsion through a microfluidizer (M-110P,

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Microfluidics, MA, USA) at a pressure of 30,000 psi, thrice. Next, the acetone was

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evaporated with a rotavapor Buchi R-300 (Buchi Analytical Inc., DE, USA) under


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vacuum and the suspension was brought with additional water to a final volume of 110

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ml. For each experiment a freshly prepared fluorescent ZNPs suspension was used.

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Transmission electron microscopy. Transmission electron microscopy (TEM)

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was performed on a JEOL 1400, (Jeol USA Inc, Peabody, MA). A droplet of freshly

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prepared ZNPs suspension was placed over a carbon grid and allowed to dry. The dried

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patch was then stained with 2% uranyl acetate solution as a contrast agent and the excess

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droplet was removed with a filter paper. The dried film was then loaded into a TEM

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sample holder for visualization.

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Dynamic light scattering. Dynamic light scattering (DLS) was used to determine

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the mean particle size, zeta potential and polydispersity index (PDI) of the ZNPs using a

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ZetasizerNano (Malvern Instruments, Southborough, MA) at a pH of 6. The analysis was

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performed at 25 ºC on a freshly prepared ZNPs sample, and the values of size and

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polydispersity index were determined using mono-modal distribution.

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Exposure of sugarcane plants to ZNPs in a hydroponic system. To study the

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uptake and translocation of ZNPs in sugarcane plants of HoCP09-804 variety, the 2

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weeks old-plants were transferred in a hydroponic culture systems (1.6g/L) spiked with

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varying doses of freshly prepared fluorescent ZNPs. In this study, the plants were exposed

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to two doses containing 0.88 mg/mL low dose (LD), and 1.75mg/mL high dose (HD) of

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fluorescent ZNPs. The concentrations selected were correlated to relevant pesticide

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concentrations when entrapped in zein nanoparticles. Control plants were not exposed to

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ZNPs and contained only water and Hoagland hydroponic solution (1:1 ratio). All

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experiments were conducted in triplicates in beakers completely covered with aluminum

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foils to prevent evaporation and the photobleaching of the fluorescently labeled NPs. All


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plants were grown in a controlled environment chamber (Intellus Environmental

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Controller, Percival: Perry, IA) at 27°C under cool, white fluorescent light with 16/8 h

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light/dark cycle. Samples of roots and leaves were collected at 2, 4, 6, 8, 10, and 12 hours

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using separate batches of sugarcane plants for roots and leaves. The collected samples

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were freeze dried using a lyophilizer FreeZone 2.5 Plus (Labconco Corp., Kansas City,

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MO) for further study.

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Confocal laser-scanning microscopy. Confocal laser scanning microscopy

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(CLSM) performed on Leica TCS SP2 microscope (Leica Inc, Buffalo Grove, IL) was

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used to visualize the fluorescent ZNPs translocation in sugarcane root samples immersed

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into Hoagland solution spiked with two different doses of ZNPs. The accumulation of

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ZNPs in the sugarcane root system was detected based on the fluorescence imparted by

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FITC. CLSM images were obtained at Excitation/Emission: 480-495nm and 550/560nm.

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Images were taken under Leica 20x (NA 0·75) oil immersion objective lens. Argon laser

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were used for NR and GFP excitation. Pictures have brightness and contrast

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enhancements. Thin sections of the root samples were placed on a glass slide covered

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with a cover slip and few drops of distilled water was added to keep it hydrated. For

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imaging the internal root structure, a standard protocol was followed which involved four

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basic steps: tissue fixation, dehydration, rehydration and successive embedment of the

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root.35 Root samples from control, low dose and high dose were washed with PBS and

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subjected to 2.3 M sucrose gradient infiltration overnight at 4°C. The root samples where

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then placed on the cryostat and sectioned with optimum cutting temperature (OCT). The

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thinner cut sections were then placed on the glass slide with the sections facing upright.


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For each study three root tips were examined from one treatment group before

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representative images were selected.

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Uptake, biotransformation, and bioaccumulation of fluorescent ZNPs.

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Pulverized freeze dried root (3mg) and leaves (2mg) samples, originating from the low

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dose, high dose and control plant batches (5 samples/batch), were weighed in amber

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Eppendorf tubes. Next, 1 ml of 1N NaOH was added to each tube and placed into an

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agitated water bath at 70° C for 30 minutes. Then, the samples were centrifuged at

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50,000xg for 15 minutes at 16° C. After digestion, the obtained clear supernatant’s

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fluorescence was determined in triplicate on 100 µl sample aliquots in a dark 96-well

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plate using a Cell Imaging Multi-Mode reader Cytation 3 (BioTek Instruments, Inc.,

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Winooski, VT) equipped with Gen5 data analysis software. Three samples of 1N NaOH

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(100 µl) were used as blank. The nanoparticle concentration in each sample was

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calculated using a standard curve developed by measuring the fluorescence (λexc = 495

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nm; λem = 525 nm) of FITC-labeled ZNPs at known concentrations digested with 1N

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NaOH.

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Statistical analysis. One way analysis of means using all pairs Tukey Kramer

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HSD were performed using JMP Pro 13 software (SAS Institute, Cary NC) to

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demonstrate significant difference (p

Zein Nanoparticles Uptake and Translocation in Hydroponically

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