Uptake, Distribution, and Bioimaging Applications of Aggregation

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Uptake, Distribution and Bioimaging Applications of AggregationInduced Emission Saponin Nanoparticles in Arabidopsis thaliana Alexander William Nicol, Kai Wong, Ryan T. K. Kwok, Zhegang Song, Ning Li, and Ben Zhong Tang ACS Appl. Mater. Interfaces, Just Accepted Manuscript • DOI: 10.1021/acsami.7b09387 • Publication Date (Web): 11 Aug 2017 Downloaded from http://pubs.acs.org on August 13, 2017

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ACS Applied Materials & Interfaces

Uptake, Distribution and Bioimaging Applications of Aggregation-Induced Emission Saponin Nanoparticles in Arabidopsis thaliana Alexander Nicol,†,⊥ Kai Wong,∥ Ryan T. K. Kwok,†,⊥ Zhegang Song,†,⊥ Ning Li,∥ and Ben Zhong Tang*,†,⊥,§ †

Division of Biomedical Engineering, Department of Chemistry, Hong Kong Branch of Chinese

National Engineering Research Center for Tissue Restoration and Reconstruction, Institute for Advanced Study, Institute of Molecular Functional Materials and State Key Laboratory of Molecular Neuroscience, The Hong Kong University of Science and Technology (HKUST), Clear Water Bay, Kowloon, Hong Kong, China. ⊥HKUST-Shenzhen

Research Institute No. 9 Yuexing 1st RD, South Area, Hi-tech Park

Nanshan, Shenzhen 518057, China. ∥Department §

of Life Science, HKUST, Clear Water Bay, Kowloon, Hong Kong, China.

Guangdong Innovative Research Team, SCUT-HKUST Joint Research Laboratory, State Key

Laboratory of Luminescent Materials and Devices, South China University of Technology, Guangzhou 510640, China. *Corresponding author: B. Z. Tang (email: [email protected])

ACS Paragon Plus Environment

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Keywords: aggregation-induced emission (AIE), nanoparticles, saponin, Arabidopsis thaliana, plant, bioimaging. ABSTRACT The application of aggregation-induced emission luminogens (AIEgens) has heralded a new age in the analysis of subcellular events and has overcome many of the limitations of conventional fluorescent probes. Despite the extensive literature investigating AIEgens in mammalian cells, few reports exist of their bioimaging applications in plant cells. In this report, we describe the first systematic investigation of the uptake, distribution and bioimaging applications of AIEgens and AIE saponin nanoparticles in the plant model system Arabidopsis thaliana. We find that the superior photostability, high colocalization with fluorescent proteins and unique tissue specific turn-on emission properties make AIEgens well suited to tackle the emergent challenges faced in plant bioimaging. INTRODUCTION Fluorescent imaging in plants has unique challenges and obstacles when compared to other model systems.1,2 Plant staining is often complicated by strong endogenous autofluorescence of plant tissues along with the impermeability of the plant cell wall to large macromolecules and protein-based labels.3 These factors impose significant challenges for plant bioimaging leading to the use of a relative select class of fluorophores with emission spectra outside the autofluorecence region including blue (DAPI),4 yellow/orange (rhodamines)5 and longer-wavelength dyes (Alexa Fluor® 647).6 In addition, researchers have observed that the emission intensity of many of these commercial dyes is much lower when compared to mammalian cell staining. This is due to the highly unfavorable bioimaging conditions within


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plant cells which induces emission quenching in conventional fluorophores in a phenomenon known as aggregation-caused quenching (ACQ).7 These highly undesired outcomes have also limited the multiplex imaging capabilities of conventional fluorophores with green fluorescent protein (GFP) technologies. This is particularly tragic given the important role GFP has played in understanding cellular protein biogenesis, transport and localization in plant biology.8−16 Despite the many tangible advantages of GFP technology, there are still limitations which need to be considered. One important limitation is the resolution limit which can be obtained considering that GFP itself is an inherently large protein.17,18 Therefore, to obtain the precise sub-cellular location of an unknown GFP-tagged proteins, one must examine the colocalization with a target specific exogenous fluorophore.19,20 However, the enhanced ACQ effects in plants drastically limits the synergistic relationship between conventional exogenous fluorophores and fluorescent proteins. One largely unexplored remedy to this problem is the replacement of conventional fluorophores with aggregation-induced emission luminogens (AIEgens) for multiplex fluorescent imaging in plant biology. AIEgens do not suffer from the ACQ effect and are able to stay highly emissive even in unfavorable bioimaging conditions like what is found inside the plant cell.21 The AIE phenomenon results from the restriction of intramolecular rotations of molecules with rotating units such as phenyl rings.22 The rotor-containing fluorophores undergo low-frequency rotations in dilute solutions which induce non-radiative decay pathways in their excited states leading to weak emission. However, in the aggregated state these rotations are blocked by intermolecular steric interactions which opens the radiative pathway.23 As previous literature has shown, AIEgens have performed better in many aspects relative to commercial fluorophores and have unique turn on responses which can be used to directly visualize biological analytes24 and


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processes.25−26 In particular, AIEgens also exhibit higher photostability than many conventional probes which would be of key impotence in plant bioimaging where high lase power is often required. To date, a large library of AIEgens exist in the literature but virtually no reports exist of their application in plant biology.27−28 RESULTS AND DISCUSSION Recognizing the great potential of AIEgens in plant biology, we first set out to systematically investigate the uptake and distribution of AIEgens using cheap and readily available onion epidermal cells as a proof-of-concept as shown in Figure 1. We selected diphenylimidazole-In (DPI-In) as our AIEgen of choice due to its positively charged structure, red emission and favorable AIE properties according to our previous reports.29 The onion epidermal cells were incubated with pure DPI-In under isotonic conditions for 10 min. As shown in Figure 1(5A), DPI-In brightly stains the cell wall of the onion epidermal cells. The plant cell wall is primarily made up of three classes of polysaccharides including cellulose, hemicellulose and pectins.30 Lignin is a hydrophobic biopolymer that fills the spaces between these three polysaccharide components. Unlike the highly hydrophilic polysaccharide components of the cell wall, lignin is hydrophobic and thus impermeable to water. Lignin plays an important role in water retention and makes it possible for the plants vascular tissue to conduct water efficiently.31 The fact that DPI-In stains the cell wall under these conditions is expected since DPI-In itself is relatively hydrophobic and positively charged leading it to have a high retention affinity within the hydrophobic lignin domains of the cell wall. This is not unique to DPI-In and is exhibited by many other similarly structured polysaccharide staining commercial dyes such as propidium iodide (PI), safranin, Calcofluor white, congo red, ruthenium red and aniline blue.32 Since multiplex imaging with fluorescently tagged membrane proteins is of particular importance in


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plant cell biology, we devised a simple strategy to both increase the solubility and deliver efficacy of DPI-In to the plant cell membrane by fabricating saponin AIE nanoparticles (saponin@DPI-In NPs). Saponins are a class of naturally occurring bioactive and biocompatible amphiphilic glycosides with unique membrane permeabilization properties.33,34 Previous reports have demonstrated the unique ability of saponins to permeabilize the plasmalemma and cell wall of plant cells to macromolecules.35,36 Saponin@DPI-In NPs were prepared by adding 200 µL of DPI-In (5 µg/mL in THF) drop-wise to 1 mL of PBS in a scintillation vial. The mixture was vigorously stirred at room temperature for 10 min with a gentle stream of N2 to help slowly evaporate the THF and form small dye aggregates. Then 20µL of saponin (1mg/mL) was subsequently added and stirred at room temperature for another 5 min. The saponin@DPI-In NPs fabrication method yielded uniform self-assembled nanoparticles around 50 nm in size as confirmed by DLS and TEM (Figure S1). The nanoparticle self-assembly process is possible due to the hydrophobic characteristics of the DPI-In nanoaggregates and the amphiphilic properties of saponin which interact to form stable core-shell complexes. This fabrication method also allowed for other AIEgens to be easily substituted for DPI-In if one desired a different emission wavelength or cellular target. As shown in Figure 1(5B), saponin@DPI-In NPs were incubated with onion epidermal cells under isotonic conditions and were again found to stain the cell wall area. However, it could not be conclusively determined if saponin@DPI-In NPs permeated past the cell wall to the cell membrane from simple visual observation of the confocal laser scanning microscope (CLSM) images. To solve this technical problem, a plasmolysis experiment was performed by exposing the onion epidermal cells to a 1M sucrose solution followed by rehydration to isotonic conditions with saponin@DPI-In NP incubation. This method effectively overcomes the immense turgor pressure of the intracellular space and allows a large quantity of


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dye to forcibly enter the cell. As shown in Figure 1(6C), the intracellular space shows bright fluorescence. This is partially due to the AIE effect where even at high concentrations bight fluorescence can be maintained without quenching. Further work is being conducted to test whether we can use this unique AIE effect for an in situ turgor pressure sensor as we believe the AIE emission intensity would be ratiometrically proportional to the internal osmotic pressure.37 Upon close examination, it can also be observed that the cell wall is still slightly fluorescent due to some hydrophobic retention of DPI-In. From these results we can conclude that pure DPI-In stains hydrophobic lignin domains in the cell wall. The permeabilization across the cell wall can be enhanced via saponin encapsulation which effectively increases the hydrophilicity of DPI-In allowing it to diffuse past the lignin domains and localize near the plasma membrane. Due to immense internal turgor pressure, DPI-In will not naturally diffuse inside the plant cell. However, high concentrations of saponin@DPI-In NPs can be directed to the cytoplasm if the plant cell undergoes plasmolysis followed by rehydration. Although onion cells are readily available, they are not commonly used as model plant organisms because of their long lifecycle. Therefore, we turned our attention to developing methods and optimized staining protocol for Arabidopsis thaliana, which is the most popular model organism for plant biology.38,39 The fluorescent staining procedure involved incubating an Arabidopsis tissue sample in a saponin solution (1 mg/mL) for 10 min. This was followed by a second incubation period with the fluorescent saponin@DPI-In NPs for 10 min. The Arabidopsis tissue was then mounted on a glass slide for CLSM investigation. This staining procedure worked well for a variety of Arabidopsis tissue types and morphologies as illustrated in Figure 2. This included staining Arabidopsis leaf cross-sections, stem cross-sections and various segments along the root system. As shown in Figure 2A−D, the Arabidopsis stem cross-section staining


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showed highly inhomogeneous staining characteristics. A closer investigation into the literature allowed us to identify that the brightly stained region was the primary xylem as illustrated in Figure 2A. The primary xylem contains an abundance of hydrophobic lignin which attracts, retains and further aggregates DPI-In molecules. The AIE properties of DPI-In allow the dye to stay highly emissive even in the aggregated state and further enables for the direct visualization of lignin in vivo. Xylem is an important vascular tissue in plants responsible for transporting water from the roots to the leaves.40 Lignin is an important component in this process providing a water-impermeable layer within the xylem vascular bundles. Previous reports have shown that the composition and vascular architecture of xylem changes as the plant develops from a seedling to a mature plant. Exactly how early stage protoxylem is converted to mature metaxylem is still uncertain and of key interest in plant biology.41 Given the high brightness and specificity to primary xylem lignin, saponin@DPI-In NPs may be utilized to monitor the lignin composition and morphological changes that occur during plant development.42 This sensitivity maybe further utilized for lignin quantification in other applications such as assessing the effects of chemical, physical and biological treatments for paper production.43 Furthermore, Arabidopsis thaliana root tissue was stained using saponin@DPI-In as shown in Figure 3. Interestingly, DPI-In shows some unique tissue selectivity which can be attributed to both its AIE properties and lignin specificity. Figure 3G shows a diagram of the Arabidopsis root architecture and cell landscape.44 There are three primary development zones within the Arabidopsis root. These include the meristematic zone which is characterized by a high rate of cell division, high cell density and an underdeveloped cell wall. Next region is the elongation zone which is characterized by auxin regulated cell wall formation and stiffening. The last region is the differentiation zone where the root hairs, secondary cell wall and lignin


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biosynthesis machinery are formed.30 As shown in Figure 3A−C, saponin@DPI-In strongly stains the meristematic zone relative to the elongation and differentiated zones. The higher florescence intensity within the meristematic zone can be explained by considering that there is a much higher cell density relative to other zones. The meristematic zone is the only area where plant stem cells are located and undergoing the highest rate of mitosis cell division.45 Additionally, these cells have less developed cell walls so the amount of exposed hydrophobic membrane lipids is greatest within the meristematic zone which leads to its increased emission. Figure 3D−F shows that further up the main taproot into the differentiated zone, saponin@DPIIn stains root hairs for the same reasons. Additionally, the lignin containing xylem in the center of the root in Figure 3E is also prominently visible. Fluorophore photostability is an important parameter to quantify in addition to tissue selectivity. This is particularly important for plant bioimaging which often requires high laser power over extended periods of time to monitor growth dynamics or protein movement.46 Therefore, highly photostable fluorophores are highly coveted and necessary for accurate colocalization experiments with fluorescent proteins. Propidium iodide (PI) is the closest commercial dye to DPI-In in terms of both its application and structure. PI has found extensive applications in plant bioimaging although it suffers from the ACQ effect. Both PI and DPI-In are positively charged and have similar emission wavelengths. Our previous reports have shown that fluorophores with AIE properties exhibit higher photostability when compared to many commercial dyes. This is because AIEgen aggregates are protected and only their outer shell photobleached while their inside core stays highly emissive. As shown in Figure 4A, the photostability of DPI-In was tested against PI with continuous irradiation time at 514 nm in Arabidopsis thaliana seedling root tissues. Remarkably, the signal intensity of saponin@DPI-In


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NPs changed only slightly whereas PI fell by almost 50%. This can be visualized in the CLSM images of Figure 4B−Ε which shows the photobleaching effects on both dyes. The superior photostability of saponin@DPI-In NPs relative to PI along with its membrane specificity compelled us to investigate its application as a colocalization probe for unknown GFP-tagged proteins in plant cells.47 Our preliminary experimental results in onion epidermal cells led us to hypothesize that saponin effectively increased the water solubility of DPI-In allowing it to diffuse past the cell wall and localize at the plasma membrane. To test whether saponin@DPI-In NPs could be used as a counter stain for subcellular membrane protein localization, we performed a colocalization analysis of DPI-In and PI with yellow fluorescent protein (YFP) tagged PIP2A aquaporin in Arabidopsis thaliana mutant root tissue as shown in Figure 5.48 Aquaporins are membrane proteins used to facilitate water transport in cells.49 We chose to examine YFP tagged aquaporins because they are integral membrane proteins and assured to be localized within the membrane.50 As shown in Figure 5A−C, the emission intensity of saponin@DPI-In is visibly brighter than that of PI (F-H). More importantly, saponin@DPI-In colocalization with YFP tagged PIP2A aquaporin is significantly higher both in terms of colocalization rate (Figure 5E) and Pearson’s correlation coefficient (Figure 5J) when compared to PI. Staining with pure DPI-In also had better colocalization results compared to PI but not as significant. This is because without saponin encapsulation, more of the DPI-In would be captured in the outer cell wall due to hydrophobic interactions. The high colocalization with fluorescently tagged membrane proteins like aquaporin coupled with its inherently high photostability and unique tissue specific turn-on emission properties makes saponin@DPI-In a unique probe in the plant bioimaging space. The high membrane specificity and compatibility


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with fluorescent proteins brings up the exciting prospect of using AIEgens to identify the subcellular distribution of unknown fluorescently tagged proteins.

CONCLUSION Just as the application of GFP technology in plants took several years longer to optimize than in mammalian cells, so too has the application of AIE technology lagged behind in plants. This study hopes to rescind this trend and offer new insights and methods for applying AIEgens to plant bioimaging using the plant model system Arabidopsis thaliana. This work has outlined a fabrication procedure for red emissive saponin DPI-In AIE nanoparticles which were able to traversed the cell wall and localize at the plasma membrane. We demonstrated that saponin DPIIn had unique tissue specific turn-on emission properties due to its hydrophobic affinity. This tissue selectivity was used to distinguish meristematic zones in root tissue and primary xylem in stem tissue. Furthermore, DPI-In had higher photostability compared to commercial plant bioimaging dyes and exhibited higher colocalization to YFP labeled PIP2A aquaporin membrane proteins when compared to PI. We suspect that this report will energize the AIE community to investigate more unique and original research into plant bioimaging.

EXPERIMENTAL SECTION Materials. Saponin (Quillaja) was purchased from Alfa Aesar, Inc (USA). DPI-In was purchased from AIEgen Biotech, Inc (Hong Kong) or synthesized according to our previous report.29 All of the standard chemicals were purchased from Sigma–Aldrich including the agar


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and growth media components. Tetrahydrofuran (THF) distilled from sodium benzophenone ketyl under nitrogen. Milli-Q water was purified via the Milli-Q Plus System (Millipore Corporation). Arabidopsis Material and Growth Conditions. Arabidopsis (wild type Col-0 and HisYFP-PIP2;1::pip2;1/2) seeds were surface-sterilized and stored in double-distilled water at 4° C for 4 days. These seeds were then mixed with 0.1% w/v agar solution, sown on agar growth media and dispensed on 9-cm diameter glass plates. The growth medium was made of 9 mM KNO3, 0.4 mM Ca5OH (PO4)3, 2 mM MgSO4, 1.3 mM H3PO4, 50 µM Fe-EDTA, 70 µM H3BO3, 14 µM MnCl2, 0.5 µM CuSO4, 1 µM ZnSO4, 0.2 µM Na2MoO4, 10 µM NaCl, 0.01 µM CoCl, 10 g/L sucrose, 1 mg/L thiamine HCL, 0.1 mg/L pyridoxine, 0.1 mg/L nicotinic acid, 100mg/L myo-inositol, and 0.8% bacteriological agar, pH 5.7. In each glass plates, 18–20 seeds were sown on the surface of the agar medium, 1.5–2 cm apart from each other. The plates were placed in a plant growth chambers with a 16-h-light/8-h-dark regime with a constant temperature at 22 ± 2° C. 7-day-old Arabidopsis seedlings were harvested for root tissue. For leaf and stem tissue, the Arabidopsis seedlings were transplanted to soil and grown further for another 3-weeks until maturity. Characterization. The absorption spectra were recorded using a Shimadzu UV-2600 spectrometer. The emission spectra were recorded using a Perkin–Elmer LS 55 spectrofluorometer. A Leica DMI 6000 fully motorized inverted microscope was used for confocal laser scanning microscopy. Synthesis of DPI-In. 4-Benzyl-1-methylpyridium iodide was first prepared through methylation of 4-benzylpyridine. A solution of DPI-CHO (0.3 mmol, 79 mg) and 4-benzyl-1-


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methylpyridium iodide (0.3 mmol, 93.4 mg) in dry EtOH was refluxed for 10–12 h. After cooled down, the reaction mixture was concentrated under reduced pressure and purified by silica gel chromatography with dichloromethane/MeOH (98:2−95:5) as eluent to afford the product (126 mg, 0.23 mmol) as a red solid. Then the counter ion of the product was changed from I− to PF6− after stirring it with KPF6 in acetone at room temperature for 4 h. Yield: 73%. m.p. = 198.6– 199.7 °C. 1H NMR (400 MHz, DMSO- d6): δ (TMS, ppm) 8.10−8.06 (d, 1H), 7.96−7.95 (m, 1H), 7.90−7.88 (m, 1H), 7.77−7.74 (d, 1H), 7.65−7.59 (m, 5H), 7.49−7.45 (m, 4H), 7.32−7.28 (m, 3H), 4.13 (s, 3H), 3.78 (s, 3H), 1.82 (s, 6H). 13 C NMR (100 MHz, DMSO- d 6 ): δ (TMS, ppm) 180.33, 143.27, 142.63, 141.82, 141.75, 135.85, 134.97, 133.02, 130.25, 129.58, 129.18, 129.09, 128.86, 128.80, 128.19, 127.47, 126.65, 122.72, 114.91, 111.75, 51.81, 34.14, 31.52, 25.33. HRMS (MALDI-TOF), m/z calcd. for (C29 H28 N3 )+: 418.2278; found 418.2289 (M+). Fabrication of Saponin AIE NPs. Saponin@DPI-In NPs were prepared by adding 200 µL of DPI-In (5 µg/mL in THF) drop-wise to 1 mL of PBS in a scintillation vial. The mixture was vigorously stirred at room temperature for 10 min with a gentle stream of N2 to help slowly evaporate the THF and form small dye aggregates. Then 20µL of saponin (1mg/mL) was subsequently added and stirred at room temperature for another 5 min. Characterization immediately followed the synthesis to limit any effects of aggregation. CLSM Fluorescent Imaging and Photostability. For root tissue, 7-day-old Arabidopsis seedlings were harvested from the agar culture. The root tissue was directly incubated with either saponin (1mg/ml) followed by the saponin@DPI-In NPs or pure DPI-In (200 µL of a 5 µg/mL solution in THF added to the root tissue suspended in 1 mL of PBS) for 10 minute intervals at room temperature. For leaf and stem tissue, a vibratome was used to make 100 µm sections and the tissue was incubated with saponin@DPI-In NPs or pure DPI-In in a similar manner. PI was


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used to stain the plant tissue according to the supplier’s specifications (2 µl of a 1 mg/mL solution). Before imaging on the confocal microscope, the cells were washed with PBS three times. The sample was placed on a concave microscope slide and imaged using a fluorescent microscope (Leica DMI 6000) at 30% laser power. Excitation wavelength was 488 nm for DPIIn, 514 nm for YFP nm and 561 for PI. For the DPI-In samples, a 600−625 nm emission filter was used. For the YFP samples, a 520−540 nm emission filter was used. For the PI samples, a 600−630 nm emission filter was used. Pearson’s correlation coefficient and colocalization rate were calculated using graphpad prism software. Photostability comparison between saponin@DPI-In NPs and PI was obtained using a Leica DMI 6000 fluorescent microscope at 30% laser power with continuous irradiation time. Signal intensity is defined by I/I0, where I0 is the initial fluorescence intensity and I is the fluorescent intensity after each subsequent scan. Colocalization statistics including colocalization rate and Pearson’s correlation coefficient were calculated using the statistical toolbox in GraphPad Prism 5 software using 95% confidence intervals.

ASSOCIATED CONTENT Supporting Information. Supporting figures show DLS characterization of saponin@DPIIn NPs (Figure S1).

AUTHOR INFORMATION Corresponding Author


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*Corresponding author: B. Z. Tang (email: [email protected]) Author Contributions A. N. performed all the bioimaging experiments, analyzed the data and wrote the paper. W.K. and N. L provided all the Arabidopsis thaliana plant samples. Z. S. synthesized and characterized DPI-In. R.T.K.K. and A. N. helped prepare the paper. R. T. K. K, N. L. and B.Z.T. helped supervise and support the research.

Funding Sources No financial conflict of interest.

Acknowledgement This work was partially supported by the National Basic Research Program of China (973 Program; 2013CB834701 and 2013CB834702), the University Grants Committee of Hong Kong (AoE/P-03/08), the Research Grants Council of Hong Kong (16301614, 16305015, 16308016, N_HKUST604/14 and A-HKUST605/16), and the Innovation and Technology Commission (ITC-CNERC14SC01 and RE:ITCPD/17-9). B.Z.T. is also grateful for the support from the Guangdong Innovative Research Team Program of China (201101C0105067115) and the Science and Technology Plan of Shenzhen (JCYJ201602229205601482). A. N. recognizes support from the Hong Kong PhD Fellowship Program.

ABBREVIATIONS


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AIE, Aggregation-Induced Emission; AIEgen, Aggregation-Induced Emission luminogen; DPIIn, diphenylimidazole-In; saponin@DPI-In NPs, saponin DPI-In nanoparticle; CLSM, confocal laser scanning microscope.

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FIGURES


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Figure 1. Fluorescent staining procedure for Arabidopsis thaliana and onion epidermal cells. (1) Plant tissue extraction, (2) fabrication of saponin@DPI-In NPs (3) incubation with either DPI-In or saponin@DPI-In NPs for 10 min, (4) CLSM sample preparation and bioimaging. (5) Fluorescent staining of onion epidermal cells in different tonic environments using CLSM. (A) Under the control isotonic conditions without saponin treatment, DPI-In fluorescence is electrostatically associated with the cell wall. (B) Under isotonic conditions with saponin


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treatment, DPI-In fluorescence is associated with the plasmalemma and inner cell wall due to electrostatic interactions. (6) Under hypertonic conditions (onion epidermal cells exposed to 1 M sucrose followed by rehydration) with saponin treatment, DPI-In fluorescence is associated with the retracted plasmalemma and cytoplasm due to cell plasmolysis as shown in (C).

Dye

concentration and staining time was 0.5 µg/mL for 10 min. All scale bars are 100 µm.


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Figure 2. CLSM images of various Arabidopsis thaliana plant tissues including leaf, stem and root stained with 0.5 µg/mL saponin@DPI-In for 10 min. The primary xylem of the stem crosssection is brightly stained due to the high hydrophobic lignin concentration. (A) Diagram of an Arabidopsis stem cross-section, (B) bright field, (C) fluorescence and (D) merged bright field and fluorescence. All scale bars are 200 µm.


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A

B Elongation Zone

C

G

Merged

Meristematic Zone D

Differentiation Zone

E

F Merged

Figure 3. Arabidopsis thaliana root tissue stained with 0.5 µg/mL saponin@DPI-In for 10 min. (A−C) meristematic zone, scale bars are 200 µm and (D−F) differentiation zone, scale bars are 50 µm. (G) Arabidopsis root architecture diagram.


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110

A

Signal Intensity (%)

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B Saponin @DPI-In 0 min

100

C Saponin @DPI-In 15 min

90 80 70

D

E

PI 0 min

PI 15 min

Saponin@DPI-In PI

60 50 0

5

10

15

Time (min)

Figure 4. (A) Photostability comparison between saponin@DPI-In NPs (red) and PI (blue) in Arabidopsis thaliana seedling root tissues with continuous irradiation time at 488 nm and 561 nm, respectively at 30% laser power. Signal intensity is defined by I/I0, where I0 is the initial fluorescence intensity and I is the fluorescent intensity after each subsequent scan. CLSM images at different time intervals for saponin@DPI-In NPs at (B) 0 min, (C) 15 min and PI at (D) 0 min, (E) 15 min, respectively. Scale bars are 50 µm for all images.


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C

YFP-PIP2A

Merged

Colocalization rate (%)

Saponin @DPI-In

B

Sap@DPI-In pixel intensity

150

A

G

100

50

0

100

*

H

80 60 40 20 0

0

20

40

60

PI

80

DPI-In

Saponin@DPI-In

YFP-PIP2A pixel intensity 150

PI

F

E

YFP-PIP2A

Merged

I

Pearson's correlation

D

PI pixel intensity

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


100

50

0

1.0

*

J

0.8 0.6 0.4 0.2 0.0

0

20

40

60

80

PI

DPI-In

YFP-PIP2A pixel intensity

Figure 5. CLSM images of the colocalization analysis between saponin@DPI-In NPs (A−C) and PI (D−F) with YFP-PIP2 aquaporin in Arabidopsis thaliana root tissue. All scale bars are 50 µm. (G and I) showing the colocalization scatter plot for saponin@DPI-In NPs and PI, respectively. Statistical comparison showing (H) Colocalization rate and (J) Pearson’s correlation coefficient statistics. Asterisks (*) indicate a significant difference with a 95% confidence interval.

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Uptake, Distribution, and Bioimaging Applications of Aggregation

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