Plant Cell Rep DOI 10.1007/s00299-014-1624-5

ORIGINAL PAPER

Uptake and cellular distribution, in four plant species, of fluorescently labeled mesoporous silica nanoparticles Dequan Sun • Hashmath I. Hussain • Zhifeng Yi • Rainer Siegele • Tom Cresswell Lingxue Kong • David M. Cahill

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Received: 8 February 2014 / Revised: 13 April 2014 / Accepted: 16 April 2014 Ó Springer-Verlag Berlin Heidelberg 2014

Abstract Key message We report the uptake of MSNs into the roots and their movement to the aerial parts of four plant species and their quantification using fluorescence, TEM and proton-induced x-ray emission (microPIXE) elemental analysis. Abstract Monodispersed mesoporous silica nanoparticles (MSNs) of optimal size and configuration were synthesized for uptake by plant organs, tissues and cells. These monodispersed nanoparticles have a size of 20 nm with interconnected pores with an approximate diameter of 2.58 nm. Communicated by K. Wang.

Electronic supplementary material The online version of this article (doi:10.1007/s00299-014-1624-5) contains supplementary material, which is available to authorized users. D. Sun
H. I. Hussain (&)
D. M. Cahill School of Life and Environmental Sciences, Deakin University, Geelong Campus at Waurn Ponds, Victoria 3217, Australia e-mail: [email protected] URL: http://www.deakin.edu.au/scitech/les/staff/cahilld/ http://www.deakin.edu.au/research/ifm/research/micro-nano.php D. Sun South Subtropical Crop Research Institute, Chinese Academy of Tropical Agricultural Sciences, Zhanjiang 524091, Guangdong Province, People’s Republic of China Z. Yi
L. Kong Institute for Frontier Materials (IFM), Geelong Technology Precinct, Deakin University, Geelong Campus at Waurn Ponds, Victoria 3216, Australia R. Siegele
T. Cresswell Australian Nuclear Science and Technology Organisation (ANSTO), New Illawarra Road, Lucas Heights, NSW 2234, Australia

There were no negative effects of MSNs on seed germination or when transported to different organs of the four plant species tested in this study. Most importantly, for the first time, a combination of confocal laser scanning microscopy, transmission electron microscopy and protoninduced X-ray emission (micro-PIXE) elemental analysis allowed the location and quantification MSNs in tissues and in cellular and sub-cellular locations. Our results show that MSNs penetrated into the roots via symplastic and apoplastic pathways and then via the conducting tissues of the xylem to the aerial parts of the plants including the stems and leaves. The translocation and widescale distribution of MSNs in plants will enable them to be used as a new delivery means for the transport of different sized biomolecules into plants. Keywords Mesoporous silica nanoparticles
Fluorescein isothiocyanate
Rhodamine isothiocyanate
Transmission electron microscopy Abbreviations MSNs Mesoporous silica nanoparticles FITC Fluorescein isothiocyanate RITC Rhodamine isothiocyanate TEM Transmission electron microscopy

Introduction Nanotechnology is a rapidly emerging field and the application of nanoparticles in medical and biological research has attracted several research groups to focus on important research topics such as targeted drug delivery, diagnostics, tissue engineering and environmental remediation (Fan

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et al. 2014; Koo et al. 2013; Machado et al. 2013; Marangoni et al. 2013). There are different types of inorganic nanoparticles with unique features, including metals, metal oxides, semiconductors, silica and carbon-based materials reported for delivery and tracking purposes (Kunzmann et al. 2011; Soenen et al. 2013). Among them, mesoporous silica nanoparticles (MSNs) have dominated as an efficient biomolecule delivery vehicle in the mammalian system (Barkalina et al. 2013; Li et al. 2011; Lu et al. 2012; Vivero-Escoto et al. 2010; Wang et al. 2013a, b). For example, mesoporous silica nanoparticles (MSNs) have attracted attention for their use as carrier systems, especially for drug delivery in the treatment of cancer in animal models (Gary-Bobo et al. 2012). MSNs are non-toxic to cells and are taken up by cells into acidic lysosomes by endocytosis, thus making them a popular candidate for drug delivery (Popat et al. 2012). In addition, suitable molecular gatekeepers are used to cap MSN pores so that an internalized cargo can effectively reach its specific target. A controlled and sustained release system is achieved through the gatekeeper’s response to internal or external stimuli, such as redox activation, competitive binding, pH and temperature changes, and light initiation (Colilla et al. 2013). In contrast to the several reports on the uptake mechanism and application of MSNs as a biomolecule delivery vehicle in mammalian system their application and knowledge on uptake pathway, sub-cellular localization and quantification in intact plants have been reported infrequently (Chang et al. 2013; Hussain et al. 2013). Several non-porous nanoparticles have been tested for their toxicity and uptake in both plants and mammalian systems but they lack the versatility that MSNs possess (Martin-Ortigosa et al. 2013; Nair et al. 2011; Navarro et al. 2012; Soenen et al. 2013). The majority of the studies were carried out with wall-less protoplasts, calli and isolated cells grown in liquid suspension (Martin-Ortigosa et al. 2012, 2013; Silva et al. 2010; Torney et al. 2007). However, Chang and colleagues showed the delivery of DNA using 100 nm MSNs to the cortical cells and endodermis of intact roots of Arabidopsis thaliana (Arabidopsis), but there was no evidence of gene expression reported in the aerial parts of the plant (Chang et al. 2013). MSNs possess many unique characteristics, such as a large volume of tunable pores, high surface area, ease of surface functionalization, physical and chemical stability, high biocompatibility and low degradability under physiological conditions, that make them ideal reservoirs for small molecules (Li et al. 2011; Nandiyanto et al. 2009; Nooney et al. 2002). Our ability to utilize the unique properties of MSNs as delivery system in plants has been restricted by a lack of understanding of how these particles are taken up and

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translocate within plants. Prior to conducting experiments for long distance transportation of MSNs, protoplasts and other in vitro grown plant cultures were considered as a good model system for testing and visualizing the uptake of MSNs functionalized with proteins and DNA as shown convincingly by other reports (Chang et al. 2013; MartinOrtigosa et al. 2012, 2013; Torney et al. 2007). Protoplasts are plant cells without a cell wall and thus macromolecules, such as nanoparticles, could be internalized by endocytosis. However, the isolation of protoplasts is a tedious process and transformed protoplasts and suspension culture cells are hard to regenerate into intact plants. Existing nanoparticle-mediated methods to deliver biomolecules into plants with intact cell walls are mainly based on mechanical forces, such as gene guns or ultrasound to penetrate the cell wall barriers (Liu et al. 2008; Martin-Ortigosa et al. 2012). An ultrasonic method has lower cost and is an easier operation, but is primarily applied on culture cells. Recently, gold-loaded MSNs have been demonstrated to deliver DNA, protein, and chemicals by the gene gun method to isolated or cultured cells (Martin-Ortigosa et al. 2012, 2013; Torney et al. 2007). Yet, bombardment is costly and the carriers with the biomolecules merely target to the surface of plant tissues. In this study, we report for the first time long distance transport, sub-cellular location and quantification of *20 nm monodispersed MSNs in three important crop species via both apoplastic and symplastic pathways and distributed through plants via the xylem.

Experimental section Synthesis and functionalization of MSNs The synthesis process for MSNs was based on our previous report with modification (Hussain et al. 2013). In brief, our results show that the ratio of CTAB: TEOS combined at high temperature provided MSNs of the appropriate size (*20 nm) to conduct plant uptake experiments (Refer online source 1). Characterization of MSN-FITC and MSN-RITC The functionalized MSNs were characterized for their shape and size using TEM (JEM-2100, JEOL, Japan). The images of MSNs were obtained at an accelerating voltage of 200 kV. Nitrogen adsorption–desorption isotherms were determined (Micromeritics Tristar 3000 analyzer, Particle & Surface Science, UK) at 77 K under a continuous adsorption condition. All samples were degassed at 150 °C for 1 h under nitrogen before measurement. The pore size distribution was calculated from adsorption branches of

Plant Cell Rep

isotherms by the Barrett–Joyner–Halenda (BJH) method. Pore volume and specific surface area were calculated using Brunauer–Emmett–Teller and Barrett–Joyner–Halenda (BET–BJH) methods. Seed germination assay Mesoporous silica nanoparticles (MSNs) were tested for any effects they may have had on seed germination of lupin, wheat and maize. MSNs at densities of 0.2, 0.5, 1, 2, 4, 10 and 20 mg mL-1 were used. For this study, Lupin seeds were commercially purchased (Naracoorte seeds, Naracoorte, South Australia, Australia), maize and wheat seeds were commercially purchased (Greenpatch Organic Seeds, Taree, New South Wales, Australia). The seeds tested for the seed germination assay were from the same lot. The MSNs were prepared in sterile distilled water prior to the exposure. The three seed types to be tested were germinated in water as a control. Prior to testing the effect of MSNs on seed germination, seeds were tested for their viability by immersing them in deionized water. Seeds that sank to the bottom were deemed viable and were then removed and were surface sterilized for 5 min in a solution containing sterile distilled water 45 % (v/v), ethanol 50 % (v/v) and hydrogen peroxide 5 % (v/v). The seeds were removed from the sterilization solution and were rinsed thoroughly three times with sterilized distilled water. The seeds were then placed into 9-cm-diameter glass Petri dishes containing moistened filter paper. The seeds were soaked with 6 mL of 20 nm MSNs at the desired densities. The Petri dishes were sealed (Parafilm MÒ, Pecheney Plastics Packaging, Chicago, USA), covered with aluminum foil and were set for germination at 21 °C in a temperature-controlled plant growth cabinet. The percentage of seeds that had germinated was calculated 5 days post seed plating. Statistical analysis was performed using GraphPad software (GraphPad, San Diego, USA). Plant material and growth condition for protoplast isolation Arabidopsis plant line Col-0 was used for protoplast isolation. The seeds were first surface sterilized and then spread onto MS plant growth medium (2.15 g of MS powder, 10 g of sucrose and 7 g of agar, pH 5.7). The seeds (100 mg) were placed into eppendorf microcentrifuge tubes and 1 ml of 70 % ethanol was added. The tube was mixed well and incubated for 2 min. The eppendorf tubes were spun down at 5,000 rpm for 2 min and the supernatant was carefully removed using a pipette. To the aspirated tube, 1 ml of 1.8 % bleach solution containing 0.1 % Tween-20 was added. The tube was

mixed for 10 min and spun down at 5,000 rpm. The supernatant containing the bleach solution was removed using a pipette. All the steps were performed in a sterile laminar flow cabinet. The seeds were washed with sterile distilled water five times and were then resuspended in 0.5 % sterile agar solution. The seeds were spread uniformly in a Petri dish containing the MS plant growth medium and were stratified for 48 h at 4 °C in the dark wrapped with aluminum foil. The Petri dish was shifted to a temperature-controlled plant growth cabinets with 8 h light/16 h dark photoperiod (at a photosynthetic photon flux density of 250 lmol m-2s-1) at 23/19 °C light/dark temperature regime. The internalization of MSNs within Arabidopsis protoplasts Fourteen days old Arabidopsis leaves were removed from the MS plates. Three grams of leaf tissue were weighed and were finely chopped in 15 ml of filter-sterilized TVL solution (0.3 M sorbitol and 50 mM CaCl2). The finely chopped tissues were then transferred into a beaker containing a cocktail of 20 mL filter-sterilized enzyme solution (0.5 M sucrose, 10 mM MES-KOH, 20 mM CaCl2, 40 mM KCl, 1 % cellulase and 1 % macerozyme). The beaker was then sealed with parafilm and covered by aluminum foil. The beaker was kept in an orbital shaker at 35 rpm for 18 h in the dark at room temperature (RT). The enzyme-digested plant material was filtered through eight layers of cheese cloth twice followed by the addition of 25 mL W5 solution (0.1 % (w/v) glucose, 0.08 % (w/v) KCl, 0.9 % (w/v) NaCl, 1.84 % (w/v) CaCl2, 2 mM MESKOH pH 5.7). The enzyme-digested plant material in W5 solution was centrifuged for 10 min at 102 rpm and 10 mL of protoplasts were collected from the interface of the enzyme and W5 solution. To the 10 mL of protoplasts collected, an additional 15 mL of W5 solution was added and centrifuged for a further 5 min at 61 rpm. The supernatant was removed and pellet containing the protoplasts was washed twice with W5 solution. The protoplasts were finally suspended in 2 mL W5 solution and their density was calculated using a hemocytometer. Arabidopsis protoplasts were diluted to 105 cells mL-1 with W5 solution and incubated with 20 nm MSNs and MSN-FITC at 50 lg mL-1 for 12 h at RT. The treated cells were washed with W5 solution and the cellular uptake was determined under the FITC channel (CLSM, 490–525 nm wavelength width). The uptake and localization of MSNs in intact plants To study the uptake and localization of MSN-FITC by the wheat and lupin plants, seeds from both the plants were

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grown in a hydroponic system. Wheat and lupin seeds were soaked in sterile distilled water for 2 h post surface sterilization. The seeds were allowed to germinate in a Petri dish containing moist filter paper for a period of 1 week. The young seedlings were transferred into a soil-free plant growth system (SPS) (Gunning and Cahill 2009). The SPS system consisted of two vertical plates that held the germinated seed at the top and allowed roots to grow vertically down between the plates on a sheet of moist filter paper supported through a reservoir of nutrient solution (Total Horticultural Concentrate; Excel Distributors, Reservoir, Australia) at the base of a box that supports the plates. The terminal part of the roots (5 mm from the root tip) of 14-day-old wheat seedlings was submerged in 1 mL of MSN-FITC solution held in a 1.5-mL eppendorf tube. Uptake experiments were performed at a density of 0.2 mg mL-1. The eppendorf tubes were attached to the plates using an adhesive tape. The roots of lupin and maize plants were exposed to MSN-FITC or RITC suspended in a 50-mL falcon tube. More specifically, MSN-FITC was tested on lupin and RITC was tested on maize. The 50-mL falcon tube contained 3 mL of nutrient solution with 0.2 mg mL-1 MSN-FITC or MSN-RITC. The seedlings were then placed in an environment-controlled plant growth cabinet (Thermoline, Coburg North, Australia) with 70 % relative humidity at 21 °C, and a 14 h light and 10 h dark cycle. The root, stem and leaves from the nanoparticle-exposed plants were embedded in optimal cutting temperature compound (O.C.T, Tissue-TekÒ) and snap frozen in liquid nitrogen. The embedded tissue samples were cut into 10 lm cross-sections using a cryotome (Zeiss). The sections were placed on a glass slide covered with anti-fading agent 1,4-diazobicyclo-(2.2.2) octane (DABCOTM, Sigma Aldrich) that prevents photo-bleaching of FITC and RITC. The cut sections were observed and images were taken using CLSM. The cross-sections were screened for FITC (490–525 nm wavelength width) and RITC (560–610 nm wavelength width) fluorescence.

tetroxide in PBS for 2 h. The blocks containing the plant specimens were washed again thrice for 10 min with PBS to remove excess osmium tetroxide. The specimens were dehydrated step by step in a series of graded ethanol solutions [30, 50, 70, 85, 95 and 100 % (v/v)] with 20 min incubation at each step. The specimens were then immersed in a mixture of propylene oxide and ERL 4206 resin (Spurr’s original formulation) at ratios of 1:1, 1:2, and 1:3 consecutively with each step lasting for 1 h. The specimens were soaked in 100 % Spurr resin overnight. After overnight incubation in 100 % Spurr’s resin, the specimens were replenished with fresh 100 % resin and were further incubated at 70 °C for 8 h. The ultrathin sections (*80 nm thick) were obtained using an ultramicrotome and post-stained with 3 % (v/v) uranyl acetate and 0.5 % (v/v) lead citrate. TEM was obtained with a JEM-2100 (JEOL, Japan) operating at an accelerating voltage of 100 kV.

Plant sample preparation for transmission electron microscopy

Root and leaf samples were analyzed with a spot size of 10 lm using a beam of 3 MeV and a beam current of 0.5–1 nA. The detection of X-rays used a 100 mm2 Ge detector with an angle of 90 msr. To prevent the scattered protons from entering the detector and to reduce the low energy X-ray yield from light elements, a 25-lm Be window and a 80-lm thick Al foil with a 0.8-mm diameter pinhole was placed in front of the detector. The data were collected using the Data Acquisition System mpsys4 from Melbourne University together with a Canberra Model 2060 digital signal processor. The quantification of the MSN from the elemental maps was accomplished using GeoPIXE 2 software (Ryan 2001).

Wheat and lupin plants grown as described above were exposed to 20 nm MSNs at a density of 0.2 mg mL-1 and the nutrient solution containing no MSNs was used as the control. The root and stem of both the plant species were cut into thin slices of 1 mm or blocks of 0.5–1 mm3. The blocks were fixed with 4 % (v/v) EMgrade glutaraldehyde prepared in phosphate buffered saline (PBS, pH 7.0) for 2 h at RT and incubated overnight at 4 °C. The next day, samples were rinsed thrice for 10 min and were post-fixed with 1 % (v/v) osmium

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Micro-PIXE instrumentation and analytical methods Sample preparation In this study, MSN distribution in the maize plant was investigated following the procedure reported by Mitani et al. (2009). Prior to elemental analysis, a preparatory confirmation test was conducted using maize as one of the major model plant systems for silica accumulation using CLSM. The maize seeds were germinated and the uptake of MSN-RITC was conducted as described previously. However, the density tested for this preparatory experiment and quantitative elemental distribution was higher (2 mg mL-1) than those tested for wheat and lupin. Plants were grown in controlled plant growth cabinets as described previously. Following 5 days of MSN exposure, excised leaf tissues were prepared using standard freezedrying reported by Kachenko et al. (2008). Microprobe analysis

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Results

Table 1 Surface area, pore volume, pore diameter and zeta potential of MSN, MSN-FITC and RITC

Synthesis of *20 nm MSNs

Samples

Surface area (m2/g)

Pore volume (cm3/g)

Pore diameter (nm)

Zeta potential (mV)

MSNs

481.2 ± 1.44

0.33

2.58

-25.9 ± 4.9

MSN-FITC

184.8 ± 2.43

0.11

-

-15.2 ± 3.8

MSN-RITC

241.2 ± 3.31

0.13

-

-8.8 ± 3.6

The synthesis of 20 nm MSNs (Fig. 1) was performed with slight modifications to the procedure previously reported by (Hussain et al. 2013) (Refer to online source 1 and 2). The synthesized monodispersed MSNs were found to exhibit type IV isotherm adsorption curve according to the IUPAC classification. The hysteresis loops in the isotherm demonstrated a relative pressure between 0.9 and 1 resulting in bi-continuous MSNs (Brinker et al. 1999)

(Fig. 2). The pore size of the MSNs was investigated by BET-BJH method and showed an average pore size of 2.58 nm for MSNs and as expected no pore size could be measured for MSN functionalized with FITC and RITC (Fig. 2; Table 1). No phytotoxic effects on seed germination

Fig. 1 TEM of monodispersed and circular shaped *20 nm MSNs with numerous interconnected pores (inset). Scale bar 50 lm

The synthesized MSNs were tested for any effects on lupin, wheat and maize seed germination. No negative effects were observed when tested with 2 mg mL-1 MSNs. However, when high densities of MSNs (20 mg mL-1) were tested, seed germination was drastically reduced in all the plant species tested. For instance, maize seed germination was found to be 6.6 % when treated with high MSN density (20 mg mL-1) (Refer to online source 3). Furthermore, MSNs at 2 mg mL-1, showed a slight, onefold increase in maize seed germination. Localization of MSNs in the protoplasts As an initial test experiment, MSNs functionalized with FITC were incubated with the protoplasts. Intact protoplasts with an ideal density of 105 cells per mL were successfully isolated from the leaves of Arabidopsis. The protoplasts incubated with appropriate density of MSNFITC resulted in the accumulation of green fluorescing nanoparticles in the plasma membrane and chloroplasts (Fig. 3A, h). The figure inset clearly shows greater accumulation of the MSNs in the chloroplasts. The protoplasts were examined under the green channel of CLSM. As expected, no fluorescence was observed when the protoplasts were treated with MSNs (Fig. 3A, c). Furthermore, the chloroplasts from the MSN and MSN-FITC treated protoplasts exhibited red autofluorescence when observed under the red channel of the CLSM.

Fig. 2 N2 adsorption and desorption isotherms of MSN (black curve), MSN-FITC (green curve) and RITC (red curve) and BJH pore size distribution in MSNs (inset). Nitrogen sorption method was used to determine the pore size distribution of MSNs. The inset figure shows the pore size distribution of 2.58 nm in MSNs alone (black curve) (color figure online)

Cellular and sub-cellular localization of MSNs in root, stem and leaves The major part of this study was to investigate the uptake of MSN-FITC in the intact cells of the whole plants tested

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Fig. 3 Uptake and quantification of MSN-FITC by the protoplasts. The protoplasts isolated from 14-day-old arabidopsis leaves were incubated with MSN and MSN-FITC *20 nm for 12 h. The protoplasts treated with MSNs and MSN-FITC were examined using CLSM and corrected total cell fluorescence was quantified using image analysis. A Representative images showing optical sections of the protoplasts (a, e) intact protoplast under the bright field, (b, f) autofluorescence from the chloroplasts under red channel, (c) protoplasts showing no FITC fluorescence as expected when treated with MSN under green channel, (d) merged images of the protoplasts showing no green fluorescence when treated with MSNs (g) fluorescence from MSN-FITC was observed in the protoplast in association with chloroplasts (arrow) under green channel and (h) the inset clearly shows the accumulation of MSN-APTES-FITC in the

chloroplast and in the cytoplasm. B The quantification of the fluorescence was performed by selecting the area showing fluorescence and the background fluorescence was subtracted from an area showing no green fluorescence. No green fluorescence could be detected and quantified in the chloroplasts treated with MSNs (cchloroplasts). However, fluorescence in the chloroplasts was significantly greater when the protoplasts were treated with MSN-FITC. Slight fluorescence was detected in the plasma membrane (p-plasma membrane) of the protoplast treated with MSN-FITC. As expected, no fluorescence could be detected in the plasma membrane treated with only MSNs. The values for each data point represent the mean ± SE of three replicate measurements. Significant differences were indicated using an asterisk. Scale bar 7.5 lm (color figure online)

in this study. To study the uptake and translocation of MSN-FITC in the intact plant cells, wheat and lupin plants were used. Two methods were used in this study. The first method was a modified hydroponic system to examine the uptake of nanoparticles and the second through a liquid solution containing the nanoparticles in a 50-mL falcon tube. In the first method, wheat plants were grown in a hydroponic system with MSN-FITC and MSNs separately in a 1.5-mL eppendorf tube (Fig. 4). The setup was designed in such a way that the main root and a few lateral roots had access to the nanoparticle solution. The crosssection of wheat root, stem and leaves was examined under the green channel of CLSM to locate the accumulation of

MSN-FITC in different cell types and associated cells. The fluorescent nanoparticles were observed in the casparian band and associated cells in the root cross-section. The examination of stem and leaf cross-sections resulted in the accumulation of fluorescent nanoparticles in the intercellular region, endodermis of the stem and in the xylem of the leaf tissue (Fig. 5). As expected, no green fluorescence was observed in the cross-sections of root, stem and leaves of wheat treated with MSNs (Refer to online source 4). Similarly, the cross-sections of lupin were examined following the same procedure that was used for wheat. The cross-section examined under the confocal microscope revealed green fluorescent nanoparticles in the epidermis

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comparison to the root cortex and leaf parenchyma cells (Fig. 8a). This gives an indication of the movement of MSNs in different plant species tested. As a further investigation, the sub-cellular compartments were quantified using the electron micrographs by counting the number of MSNs. An area of 1.10 lm2 was considered for quantifying the number of MSNs in the cell wall, intercellular space and vacuole. This resulted in *50 MSNs in the cell wall, 20 in the intercellular region and 30 in the vacuole of the lupin stem. The number of MSNs in the cell wall was significantly greater than the accumulations found in the intercellular space and vacuole of the lupin stem (Fig. 8b). Quantification of MSNs in the aerial parts of the plant

Fig. 4 Schematic representation of the modified hydroponic system. The diagram shows the experimental setup for uptake of MSNs from the 1.5-mL tube to various parts of the plant. Constant supply of nutrient solution was provided from the reservoir

and associated cells around the casparian band of the wheat root. Nanoparticles were also found in the apoplast and ground tissue of the stem and were transported through the vascular tissue to the palisade and spongy tissue of the lupin leaves (Fig. 6). As expected, no green fluorescence was observed when tested with MSNs (Refer to online source 5). To further investigate the localization of MSNs at sub-cellular level, transmission electron microscopy (TEM) was used. Wheat and lupin plant roots were exposed to 0.2 mg mL-1 MSNs. Five days post treatment, cross-sections of the root and stem were examined under the TEM. Representative images of the lupin plants have been shown to accumulate *20 nm MSNs in the cell wall, intercellular region and vacuole of the lupin root (Fig. 7). The fluorescence from the MSN-FITC in the root, stem and leaves of wheat and lupin plants was quantified using image analysis. The corrected total cell fluorescence (CTCF) was calculated using imageJ software program. The amount of fluorescence quantified was correlated to the amount of fluorescent nanoparticles present in that region. For instance, the amount of fluorescence found in the casparian band of the wheat root was found to be significantly greater than that found in the endodermis and xylem. However, the amount of fluorescence was found to be significantly greater in the cortex of the lupin stem in

To confirm the presence of MSNs and their chemical composition, micro-PIXE elemental analysis was performed on the roots and leaves of maize plants. Prior to micro- PIXE analysis, a test was conducted to confirm the uptake of MSNs in the root tissue post 5 days of nanoparticle exposure to maize plants. MSN-RITC was exposed to the roots of a 3-week-old maize plant in a tube containing 3 mL of the treatment solution. The uptake of nanoparticles in the root cross-sections was revealed using CLSM by the accumulation of red fluorescing MSNs in the xylem (Fig. 9). The preliminary results with microPIXE elemental detection confirmed the uptake of MSNs through the roots of the maize plant (Fig. 10b). The MSN accumulation was found to be greater than the plants treated with water (Fig. 10a). The elemental profiles were determined for silica (Si), calcium (Ca) and potassium (K) (Fig. 10 and refer to online source 6). Representative images of the enlarged elemental maps are shown (Fig. 10). The profiles in Fig. 11 show the elemental concentrations along the thin green line shown in Fig. 10. For example, maize roots exposed with MSNs showed silica concentrations between 250–750 ppm (Figs. 10f, 11b). Elemental silica in water-only treated plants was measured to determine the background contribution of the element and was found to be below the minimum detection level (MDL) (Fig. 11a). We also found MSNs to be co-localized with potassium that is important for normal plant growth (Fig. 11b).

Discussion In this study, we report the synthesis of *20 nm monodispersed MSNs and their uptake and quantification in four plant species. The mesoporous nature of the MSNs and their ready uptake and movement within the tested plant species gives rise to their potential as a molecular delivery vehicle. We have shown that fluorescently labeled

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Fig. 5 Accumulation of MSN-FITC by wheat. Three-week-old wheat plants were exposed to MSN and MSN-FITC solution for 5 days in a 50-mL falcon tube. The uptake of fluorescent MSN-FITC by wheat root, stem and leaves was analyzed under the bright field and green channel using CLSM. a–c Bright field image showing cross section of wheat root, stem and leaf treated with MSN-FITC, d–f images of wheat root, stem and leaf cross-sections observed under the green channel suitable for FITC detection, d aggregates of fluorescent MSN-FITC in the casparian band of the developed root and smooth

flow of MSN-FITC towards the xylem were also observed, e fluorescent nanoparticles were in the intercellular spaces and in the endodermis of the stem and f showing accumulation of the fluorescent aggregates in the xylem and associated cells of the vascular tissue. d– f The magnified inset in these images clearly shows the accumulation of fluorescent nanoparticles. Images g–i are the corresponding merged images of (a, d), (b, e) and (c, f). Where x xylem, c cortex, ep epidermis and en endodermis. Scale bar 25 lm (color figure online)

MSNs of this size were efficiently absorbed by the roots and were able to be detected in the aerial parts of the plants. The movement of MSN within the cellular and subcellular regions of the plant root, stem and leaves was confirmed using confocal microscopy, TEM and a microPIXE elemental detection system. More interestingly, for the first time we found that MSNs exposed to the roots of the maize plant were transported to the aerial parts of the plants. The percentage of MSN accumulation in the root was found to range between 25 and 37.5 % using microPIXE as a sensitive and reliable technique. The nanoparticle synthesis procedure to achieve spherical shaped *20 nm monodispersed MSNs was fine-tuned

for the plant uptake study. Our results show that the concentration of the surfactant and optimum pH are critical parameters in obtaining monodispersed MSNs. We were successful in achieving monodispersed nanoparticles to show the uptake and widespread distribution of MSNs in plants. These MSNs are unique because they have interconnected pores with uniform pore volume. The ratio of CTAB:TEOS combined at higher temperature provided MSNs of the appropriate size (*20 nm) for conducting plant uptake studies. It was important to synthesize MSNs of this type and size to ensure that the MSNs were taken up through the pores within the microfibrillar matrix of the root cell walls. Even though the methodology used to determine

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Fig. 6 Accumulation of MSN-FITC by lupin. Lupin plants were exposed to MSN and MSN-FITC solution for 5 days in a 50-mL falcon tube. The lupin plants were 3 weeks old. The uptake of fluorescent MSN-FITC by the root, stem and leaves of lupin was analyzed under green channel using CLSM. a–c Showing crosssections of lupin root, stem and leaf treated with MSN-FITC under the bright field of CLSM, d showing aggregates of fluorescent MSN-

FITC in the epidermis and endodermis of the root, e fluorescent nanoparticles accumulated in the ground tissue of the stem and, f showing accumulation of the fluorescent nanoparticles in the pallisade and surrounding cells. d–f The magnified inset in these images clearly shows the accumulation of fluorescent nanoparticles. Where x xylem, ep epidermis, en endodermis, pm palisade mesophyll and sm spongy mesophyll. Scale bar 25 lm (color figure online)

the pore sizes of the plant cell wall has varied, reports have shown that the average pore size is within the range 5–20 nm in diameter although there are reports of much larger pores within the cell walls (Carpita et al. 1979; Carpita and Gibeaut 1993; He et al. 2009; Nooney et al. 2002; Pereira et al. 2011; Taiz 1984). We used a fluorescent marker for tracking MSNs in the cells and tissues and found it to be a sensitive and reliable method for their detection, confirming its usefulness in nanoparticle localization in living cells (Hussain et al. 2013). In addition, we found that RITC could be used as an alternative to FITC. The autofluorescence emitted by endogenous proteins and metabolites in some cells were similar in wavelength to that of FITC. For example, the use of RITC as the fluorophore in the plant root is more useful than in leaves because the red fluorescence from the RITC is indistinguishable from the red autofluorescence emitted by chloroplasts. Furthermore, because RITC and FITC labeled MSNs can potentially be loaded with biomolecules and encapsulated, they can be followed to sites where release of the biomolecule takes place. We chose to use FITC or RITC as the reporter molecule in our study but other similar, or

more brightly fluorescent species, could also be used, for example Cy5.5 dye (Ma et al. 2012). There are a number of reports of the uptake by plants of non-porous nanoparticles but majority of these particles have been used with plant suspension cultures and protoplasts (Cifuentes et al. 2010; Lee et al. 2012; Nair et al. 2011; Navarro et al. 2012; Sabo-Attwood et al. 2012; Slomberg and Schoenfisch 2012; Torney et al. 2007; Wang et al. 2012). Therefore, for the first time, we analyzed the uptake, distribution and quantification of MSNs in whole plants, in the roots and in the aerial parts of three different plant species that were grown in a modified hydroponic system and the roots exposed to an MSN suspension. We also tested the effect of the MSNs on seed germination to determine whether there are any adverse effects of these particles on plant growth. To test the effects of MSNs on seed germination, we used a broader concentration range than used previously (Hussain et al. 2013; Rico et al. 2011; Stampoulis et al. 2009) and at lower concentrations found no significant effect on germination (2 mg mL-1). However, complete inhibition of seed germination was observed when MSNs were tested at high concentrations (20 mg mL-1). This inhibition of seed germination by

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Fig. 7 TEM of lupin root. The 3-week-old lupin plants were exposed to MSNs for 5 days and cross-sections of the roots were examined with TEM. (a–e) water treated control and (b–f) MSN treated. MSNs were detected in the cell wall (cw, inset) (b), intercellular space (is, arrow) (d) and vacuole (v, inset) (f) but no MSNs were found in the corresponding parts of the lupin root treated with water. Scale bar (a– d) 100 nm and (e–f) 2 lm

silica nanoparticles may be caused by a decrease in pH in the solution surrounding the seeds because of the formation of silicon colloids (Cauda et al. 2010; Lin et al. 2011). Silicic acid is formed when Si–O binds to water molecules and this decreases the pH (Cauda et al. 2010; Lin et al. 2011). Therefore, at the higher concentration of MSNs tested, a decrease in the pH of the water resulted in the inhibition of seed germination. Recently, MSNs with an external coating of a hydrophilic polymer shell (polyethylene glycol) have been shown to have slower biodegradation kinetics and a more neutral pH is maintained in comparison with uncoated MSNs (Cauda et al. 2010; Lin et al. 2011). In the present study, we were able to transport fluorescing MSNs to cellular and specific sub-cellular locations of the

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Fig. 8 Quantification of fluorescence and MSN accumulation in a number of sub-cellular locations of wheat and lupin from the confocal and TEM micrographs. A The fluorescence was measured using the imageJ software in the casparian strip (cs), endodermis (en) and xylem (x) of the entire wheat (white bars). The fluorescence measured cs was found to be significantly greater than the fluorescence measured in en or x. Similarly, fluorescence measured in the ground tissue (gt) of the lupin stem was greater than the fluorescence measured in cortex (c) or parenchyma (p) (grey bars). Significant differences were indicated using an asterisk. Where w-r, s and l correspond to root, stem and leaf of wheat and l-r, s and l correspond to root, stem and leaf of lupin. The values for each data point represent the mean ± SE of three replicate measurements. B The number of MSNs was counted in the cell wall, intercellular space and vacuole of the lupin from the TEM. The number of MSNs counted in the cell wall was significantly greater than the accumulations in intercellular space or vacuole

entire plant. Prior to the plant uptake experiments, we showed that MSNs labeled with a reporter molecule were taken up by protoplasts and were localized in the chloroplasts. Our results were quite similar to the results obtained by Torney et al. (2007). It is interesting to note the affinity of silica to the chloroplast, but why this might be so is not clear at this stage. The results from the direct uptake through the roots of the plants tested showed that MSNs moved upwards through the xylem and downwards through the phloem using the source sink pressure gradient (Oparka and Cruz 2000). In our results, we found that MSNs were embedded in the plant cell walls and this clearly indicates the interaction between MSNs and cell wall components. In addition, previous reports have confirmed that the nanoparticles move radially between cells through the plasmodesmata

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Fig. 9 Distribution of MSN-RITC in maize roots. Three-week-old maize plants were exposed to MSN-RITC for 5 days. a–c Maize root cross-sections treated with MSNs under bright field and red channel using CLSM. b As expected, no fluorescence was observed from the roots treated with MSNs. c Merged image of a and b. d–f Maize root

cross-section treated with MSN-RITC under bright field and red channel using CLSM. e Bright red fluorescence was observed in the xylem indicating root uptake of MSN-RITC (arrow). Scale bar 25 lm (color figure online)

Fig. 10 Elemental maps of cryo-fixed freeze-dried MSN treated roots and leaf of maize. a Roots treated with water showed no silica accumulation and the silica limit was considered below minimum detection level (MDL), b MSN treated root showing heavy accumulation of silica in the maize roots and other essential elements for normal plant growth such as, c potassium and d calcium were also detected. More interestingly, e silica levels were found to be below MDL in maize leaf when the roots were exposed only with water and f leaves showed bright stripes of silica nanoparticle in the xylem of the leaf. Scale bars for (a– d 300 lm and e–f 500 lm

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et al. 2007; Ma and Yamaji 2008; Ma 2004; Ma and Yamaji 2006; Richmond and Sussman 2003).

Conclusions

Fig. 11 Quantitative elemental profile for MSNs (Si), Potassium (K) and Calcium (Ca) across the freeze-dried maize leaf section. A Elemental silica in water-only treated plants was measured to determine the background contribution of the element. The accumulation of background silica was found to be below the minimum detection level (MDL, black curve), and the accumulation of K (red curve) and Ca (blue curve) was also measured. B The accumulation levels of MSNs were measured in the leaves and were found to be between 100 and 800 ppm (black curve), and the accumulation of K (red curve) and Ca (blue curve) were also measured (color figure online)

We have successfully controlled the particle size to *20 nm by controlling the amount of TEOS, CTAB, reaction time, and pH value. Our study demonstrates that MSNs can penetrate the cell wall, enter the endodermis and intercellular spaces and to the vascular tissue to be transported to the aerial parts of the plants. In addition, we could locate the accumulation of MSNs in the chloroplast when MSNs were taken up by the protoplasts. This is a great advantage because it opens up an avenue for transforming chloroplast with plant transformation vectors using MSNs as delivery vehicles. From this study, we could say that no external force or biochemical treatment is required for MSNs to penetrate the cell wall. As the results provide an indication of where these MSNs become localized, an efficient quantitative data could be derived for prolonged exposure of MSNs and other elements important for plant growth could be quantified simultaneously. Finally, MSNs could potentially be loaded with small biomolecule of interest such as plant growth regulators, pathogen effector molecules and large molecules such as DNA and siRNA to be used as a gene delivery system. Acknowledgments This work was funded by the Centre for Chemistry and Biotechnology, Deakin University. Dequan Sun and Zhifeng Yi were supported by a Deakin University postgraduate scholarship. Conflict of interest of interest.

(Corredor et al. 2009). There are proteins closer to the plasmodesmata region involved in the process and direct the nanoparticles to plasmodesmata (Oparka 2004). Once the nanoparticles pass through the plasmodesmata, they accumulate in the cytoplasm and move to the endodermis and the casparian strip. However, the initial transport is through apoplastic or symplastic pathway once they reach the endodermis and the casparian strip. The localization of MSNs in the cell wall, plasmodesmata, cortical cells, endodermis, intercellular space, vascular tissue and the quantification of MSNs from CLSM, TEM and micro-PIXE elemental analyses are well documented in this study. Silica is an important element for plant growth. The use of silicabased nanoparticles for transporting biomolecules of interest would not only act as a delivery vehicle but plants could also incur potential benefits such as enhancement of pest and pathogen resistance, drought and heavy metal tolerance and improvement of agricultural crop quality and yield, in a wide variety of plant species (Fauteux et al. 2005; Liang

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The authors declare that they have no conflict

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