Plasma Treatment Converts Phenolic Compounds into Fluorescent

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Plasma Treatment Converts Phenolic Compounds into Fluorescent Organic Nanoparticles for Cell Imaging Yong Liu, Mingzhu Yang, Juanjuan Li, Wei Zhang, and Xingyu Jiang Anal. Chem., Just Accepted Manuscript • DOI: 10.1021/acs.analchem.9b00837 • Publication Date (Web): 22 Apr 2019 Downloaded from http://pubs.acs.org on April 22, 2019

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Analytical Chemistry

Plasma Treatment Converts Phenolic Compounds into Fluorescent Organic Nanoparticles for Cell Imaging Yong Liua,b, Mingzhu Yanga, Juanjuan Lia, Wei Zhanga, and Xingyu Jiang*a,b,c Beijing Engineering Research Center for BioNanotechnology, CAS Key Laboratory for Biomedical Effects of Nanomaterials and Nanosafety, CAS Center for Excellence in Nanoscience, National Center for NanoScience and Technology, No. 11 Zhongguancun Beiyitiao, Beijing 100190, P. R. China a

b

University of Chinese Academy of Sciences, No. 19A Yuquan Road, Beijing 100049, P. R. China

Department of Biomedical Engineering, Southern University of Science and Technology, No. 1088 Xueyuan Road, Shenzhen, Guangdong 518055, P. R. China c

ABSTRACT: Fluorescent organic nanoparticles (FONs) are promising alternatives for biological imaging applications owing to the increasing concerns over the potential toxicity and poor degradability of inorganic particles-based probes. However, synthesis of stable, small-sized FONs in aqueous media remains challenging. Inspired by the self-polymerization chemistry of phenolic compounds, we demonstrate ultrafast synthesis of FONs (phenolic compounds-derived FONs, PhFONs) from a variety of molecular building blocks including dopamine, norepinephrine, pyrogallol, and gallic acid, simply by plasma treatment at the aqueous interface. Specifically, using dopamine as the precursor, poly(dopamine) (PD)-FONs featuring a small size of 3 nm are obtained within 1 min. Compositional and structural characterizations confirm the polymeric architectures in PD-FONs. The PhFONs, with multicolor emissions, excellent biocompatibility, high stability, and sizedependent access into cell nucleus, are suitable for live cell imaging and developing nucleus-targeting imaging platforms.

INTRODUCTION The use of fluorescent materials to image biological samples has triggered broad investigations in chemical/bio sensing1 and monitoring crucial biological events such as cancer metastasis2 and intracellular reagent delivery.3 Fluorescent nanomaterials, both inorganic and organic, are intriguing for these applications owing to their longer retention time and higher stability against photobleaching compared to organic dyes and macromolecules.4 Various formulations, including quantum dots,5 upconversion nanoparticles,6 metallic nanoclusters,7 and organic nanoparticles,8 have been proposed as imaging probes. Among them, fluorescent organic nanoparticles (FONs) based on self-assembled molecular units have received increasing attention owing to their structural variability, low toxicity, and favorable biodegradability. Involved materials such as conjugated polymers,9, 10 semiconducting polymer dots,11, 12 and aggregation-induced-emission nanoparticles,13 have established a burgeoning field with bright prospects in analytical and biological applications. To fabricate FONs, commonly used strategies are reprecipitation,14 miniemulsion,15 and self-assembly.16 For the first two types of methods, organic solvents are inevitably involved, which are hazardous to human body and make the isolation process difficult. The residual organic solvents in FONs may damage cells and animals. These defects are overcome by the self-assembly of watersoluble building blocks such as monodisperse oligomers or

chromophores. However, the assembled FONs lack longterm stability in water. Additionally, most of existing FONs have large sizes (up to hundreds of nanometers), which not only impede their interactions with cells, but also cause stability problems. Therefore, new methodologies to synthesize small-sized FONs in aqueous media is still in a need. Phenolic compounds, which widely exist in mussel adhesive proteins and plant-derived foods/beverages, have exerted profound impact on surface chemistry and drug development.17, 18 Their catachol/pyrogallol-rich skeletons facilitate easy oxidation and polymerization in aqueous environment to form crosslinked assemblies.19 Recently, various micro/nano structures based on these compounds, such as nanoparticles, capsules, and assembled micro/nano composites, have been developed and applied.20, 21 These materials have shown excellent compatibility in biomedical applications, both in vitro and in vivo. Specifically, poly(dopamine) nanoparticles (PDNPs) have emerged as versatile platforms for sensing, drug delivery, and photothermal therapy.22-24 PDNPs are nonfluorescent and widely used as fluorescence quencher.25 A few studies have synthesized PD-based FONs and applied them in sensing and imaging.26-30 To obtain PD-FONs, concentrated H2O2 (up to 6%) is generally required, alone (to break down bulk PD26, 27) or with additives (e.g., nano-sized Fe3O428 and MnO229 to oxidize dopamine into FONs, and polyethyleneimine to

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form conjugated composites30), which causes huge waste of resources and poses severe safety hazards. Besides, the extendibility of using other phenolic compounds to make FONs has not been investigated. Plasma technology provides a green toolbox for robust synthesis, modification, and functionalization of various materials.31 Compared to solution chemistry, it shows irreplaceable advantages such as mild reaction condition, fast processing, and environmental friendliness. Nonthermal plasma in which the produced chemical species do not exist in thermal equilibrium state possesses low temperature but high reactivity.32 Direct contact of plasma with liquid produces various kinds of reactive species in the medium, such as persistent reactive oxygen/nitrogen species (ROS and RNS, e.g., H2O2 and NO3-), short-lived radical/atomic species (e.g., O, •NO, and •OH), and non-radical chemical compounds (e.g., singlet oxygen, 1O2).33 Phenolic compounds can be easily oxidized and subsequently form crosslinked structures, which is generally triggered by oxidizing agents or ultraviolet light.34, 35 Therefore, we anticipate the synthesis of phenolic compounds-derived FONs (PhFONs) by nonthermal air plasma treatment. Under the low temperature, the functional groups from the molecular reactants are well preserved in the FONs, which is beneficial for their chemical/biological applications. Herein, a unique, yet missing interface, the plasma/liquid interface, is presented as a new stimulus for the oxidation and polymerization of various phenolic precursors including two catechol compounds dopamine and norepinephrine, and two pyrogallol compounds pyrogallol and gallic acid. Consequently, FONs based on PD, poly(norepinephrine) (PN), poly(pyrogallol) (PP), and poly(gallic acid) (PG) are obtained. Specifically, PD-FONs appear as ultrasmall nanodots with an average size of 3 nm, are stable against photobleaching and long-term preservation, and show excellent biocompatibility in vitro. These PhFONs exhibit excitation-dependent emission property and size-dependent nucleus-targeting property, and are successfully utilized for live cell imaging. EXPERIMENTAL SECTION Materials. Dopamine hydrochloride and tannic acid were from Sigma-Aldrich. Noradrenaline bitartrate monohydrate and quinine sulfate were from Aladdin. Pyrogallol was from Bide. Gallic acid was from Acros. Hydrochloric acid was from Beijing Chemical Works. Dulbecco’s minimum essential medium (DMEM), fetal bovine serum (FBS), and trypsin were from Invitrogen. Penicillin-streptomycin (PS) was from MP Biomedicals. RedDot 1 far-red nucleus stain was from Biotium. Plasma-triggered synthesis of PhFONs. In a typical synthesis of PD-FONs, dopamine hydrochloride was dissolved in 10 mM Tris-HCl buffer (pH 7.0) to reach a concentration of 10 mM (~2 mg/mL). The solution was dropped onto a Teflon slide (10 μL/drop) to form a droplet array (20-50 droplets/slide), which was then put into the plasma chamber (Plasma etching machine, Chengdu

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Mingheng Co. Ltd) and treated for 1 min at a power of 270 W. The droplets were recollected in a tube and maintained at room temperature for 1 h to complete the reaction. Afterwards, hydrochloric acid was added into the sample (final concentration: 10 mM) to terminate the reaction system. These steps were repeated several times to obtain desired amount of PD-FONs. The freshly prepared sample was used directly for ultraviolet-visible (UV-Vis) absorption spectra and fluorescence excitation/emission spectra measurement. For cell experiments, the sample was subjected to ultrafiltration (cut-off molecular weight Mw=1000) and then lyophilized. To prepare PD-FONs with larger size, 10 mM pH 8.8 Tris-HCl buffer was used. To prepare PhFONs from other phenolic compounds, dopamine hydrochloride was replaced with norepinephrine (noradrenaline bitartrate monohydrate), pyrogallol, gallic acid, and tannic acid. Accordingly, PNFONs, PP-FONs, and PG-FONs were obtained. Characterizations. High-resolution transmission electron microscope (HRTEM) images were obtained on a Tecnai G2 F20 U-TWIN transmission electron microscope. TEM images were obtained on an HT 7700 transmission electron microscope. Images were analyzed by Image J. Xray diffraction (XRD) patterns were acquired at a scan rate of 5o/min in the 2 theta range of 10-80 degree (Cu Ka radiation, Bruker D8 Advance, German). Data were analyzed by High Score Plus. Raman spectra were obtained on a Raman spectrometer (LabRAM HR800). X-ray photoelectron spectroscopy (XPS) analysis was performed on a Thermo Scientific ESCALAB 250XI photoelectron spectrometer. Data were analyzed by XPS Peak 41 and CasaXPS. The binding energies were calibrated with C 1s (284.8 eV). Fourier transforming infrared (FT-IR) spectra were recorded on a FT-IR spectrometer (PerkinElmer, Spectrum one). KBr crystals were used as the matrix for sample preparation. Ultraviolet-visible (UV-Vis) absorption spectra were recorded on a UV-Vis spectrophotometer (UV-2450, Shimadzu, Japan). Florescence excitation/emission spectra were recorded on a spectrofluorophotometer (RF-5301PC, Shimadzu, Japan). The band pass was set as 5 nm for both excitation and emission measurements. The quantum yield of PD-FONs was measured using quinine sulfate in 0.1 M H2SO4 (literature quantum yield 0.54 at 360 nm) as a reference. Time-resolved fluorescence spectra were recorded on a steady state/transient state fluorescence spectrometer (NanoLOG-TCSPC). The fluorescence lifetime (τ) of PDFONs was measured under the excitation of 340 nm. The decay trace was fitted using biexponential functions Y(t) based on non-linear least squares analysis:

( )

Y(t) = α1exp -

t

τ1

( )

+ α2exp -

t

τ2

Where α1 and α2 are the fractional contributions of timeresolved decay lifetime of τ1 and τ2. τ was calculated based on the following equation:


τ=

α1τ21 + α2τ22 α1τ1 + α2τ2

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Analytical Chemistry Photostability test was performed on a spectrofluorophotometer. PD-FONs were continuously illuminated with 400 nm light source for 2 h. Fluorescence emission intensities at different time points were recorded and compared. Long-term storage stability test was performed at room temperature for 3 months. UV-Vis absorption spectra and fluorescence emission spectra before/after preservation were recorded and compared. Confocal laser scanning microscope (CLSM) images were obtained on a Zeiss LSM 710 confocal microscope system. Images were processed by ZEN 2.3. Cytotoxicity of PD-FONs. NIH 3T3 fibroblasts were cultured in complete DMEM containing 10% FBS and 1% PS. The passage of cells was conducted using 0.25% trypsin containing 0.02% ethylenediamine tetraacetic acid every 23 days. Unless otherwise specified, PhFONs were dissolved in DMEM containing 10% FBS and 1% PS in cell experiments. For cell viability assay, NIH 3T3 fibroblasts were seeded in 96-well cell culture plates (Costar, Corning, NY, USA) at a density of 104 cells/well and cultured overnight. The cell culture medium was then replaced with PD-FONs solution (10 to 120 μg/mL). After incubation for 24 h, cells were rinsed with PBS and incubated with Cell Counting Kit-8 (CCK-8) reagents for 2 h. Cell viability was calculated based on the optical degree (OD) at 450 nm. For hemolytic property assay, human red cells were collected, diluted to 4%, and incubated with PD-FONs (10 to 150 μg/mL) in 96-well plates for 3 h. Samples were then centrifuged at 10050 rpm for 3 min and measured on a microplate reader at 540 nm. Cell imaging using PhFONs. NIH 3T3 fibroblasts were seeded in 30 mm confocal dishes (Costar, Corning, NY, USA) at a density of 1.5×105 cells/dish and cultured for 24 h. The cell culture medium was then replaced with different PhFONs solutions (100 μg/mL). After incubation for 4 h, cells were rinsed 3 times with PBS to remove excess PhFONs, and then observed under the confocal microscope using 405 nm laser. For multicolor cell imaging, PD-FONs were used and cells were excited by 405 nm and 488 nm lasers. For colocalization, cells were stained with RedDot 1 (1:200 dilutions in DMEM) for 10 min after the uptake of PD-FONs and then observed using 405/488 nm (for PD-FONs) and 543 nm (for RedDot 1) lasers. For real time tracking of the uptake of PD-FONs, cells were incubated with PD-FONs in PBS, and then observed using 405 nm laser at different time points (0, 15, 30, 60, 90, 120 min). RESULTS AND DISCUSSION Plasma-induced fluorescence in phenolic compounds. We examine the effect of plasma treatment on dopamine monomer. We introduce the dopamine precursor onto a Teflon slide as individual droplets and then expose the droplet array to plasma atmosphere. These droplets with small sizes (10 μL/drop) resemble microliter scale reactors that ensure sufficient interfacial interactions and fast reaction kinetics. The plasma is generated by electrical breakdown of low-pressure air in a plasma

etching machine (so-called glow discharge plasma, a type of nonthermal plasma). It should be noted that the temperature of dopamine droplets can drop down to 0 oC in the low-pressure plasma since ice grains are observed sometimes. We dissolve dopamine chloride (10 mM) in neutral medium (pH 7.0 Tris-HCl buffer, 10 mM) to inhibit its spontaneous self-polymerization under alkaline conditions. As expected, the control sample (dopamine solution in the air) shows negligible color change within 2 hours. However, the droplets immediately change into pale yellow after plasma treatment for 1 min (Figure 1a), suggesting the oxidation of dopamine. Strikingly, the product shows intense green fluorescence upon irradiation by violet light. The oxidized dopamine exhibits a broad absorption band from 300 nm to 600 nm with no distinctive peaks (Figure 1b), which differs from bulk PD aggregates having a characteristic peak at 420 nm. Its emission peaks shift accordingly with different excitation wavelengths (Figure 1c). The strongest emission is located at 430 nm under 340 nm excitation (Figure S1). Such a variation is correlated with the wide absorption spectrum, indicating the inherent chemical heterogeneity of the product. We also evaluate the response of other phenolic compounds to plasma treatment. Excitation-dependent fluorescence emissions are observed in norepinephrine, pyrogallol, and gallic acid, but not in tannic acid (Table S1 and Figure S2) This is probably due to the huge, sable skeleton of tannic acid.

Figure 1. Plasma-triggered synthesis of PD-FONs. (a) Digital photographs of dopamine and plasma-treated dopamine under daylight (up) and violet light illumination (bottom). C: control sample. P: plasma-treated sample. (b) Ultraviolet-visible (UV-Vis) absorption and fluorescence emission spectra (excitation at 400 nm) of plasma-treated dopamine. (c) Normalized fluorescence emission spectra of plasma-treated dopamine under different excitation wavelengths. Formation of PhFONs. Transmission electron microscope (TEM) images confirm the presence of welldefined PhFONs in the plasma-treated samples. Interestingly, FONs synthesized from different molecular precursors show different size and morphology. The two catechol-derived FONs, PD-FONs and PN-FONs, appear as quasi-spherical nanoparticles (Figure 2a-2b), while the two pyrogallol-derived FONs, PP-FONs and PG-FONs, possess nonuniform, worm-like morphologies (Figure 2c-2d). These PhFONs all have broad size distributions. The average sizes of PD-FONs, PN-FONs, PP-FONs, and PGFONs are 3.36±0.21 nm, 7.80±1.32 nm, 17.62±3.20 nm, and



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12.82±3.28 nm, respectively. The irregular shape and large size of PP-FONs and PG-FONs are probably due to their pyrogallol structures that are easier to be oxidized and form crosslinked oligomers compared to catechol compounds.

Figure 2. TEM images of PhFONs. (a) PD-FONs (HRTEM images). (b) PN-FONs. (c) PP-FONs. (d) PG-FONs. The insets show the size distribution calculated from TEM images. Scale bar: 20 nm. The yield of PhFONs is correlated with the droplet size used for reaction, the smaller size, the higher yield. For PDFONs, the absorbance and fluorescence intensity of the plasma-treated sample progressively increase as the droplet size decreases from 50 μL to 5 μL (Figure S3). This indicates that the plasma has an effective penetration depth at the liquid interface. Droplets larger than 50 μL do not favor the synthesis of PhFONs since large ice grains appear. To compromise favorable yield and scaled synthesis, 10 μL is applied to prepare PhFONs. Morphological, compositional, structural, and optical analysis of PD-FONs. We select PD-FONs for further characterizations. As shown by high-resolution TEM (HRTEM) images, most PD-FONs display amorphous morphologies (Figure 3a), which is also supported by the X-ray diffraction (XRD) pattern and Raman spectrum. In the XRD pattern, the broad peak band centered at 25o (0.34 nm) is attributed to disordered carbon atoms (Figure S4a).36 Since PD-FONs possess low content of lattice structures, no obvious D or G bands are detected in the Raman spectrum (Figure S4b). Nevertheless, for a number of PD-FONs, well-resolved lattice fringes can be observed under TEM (Figure 3a-3b, Figure S5). The interplanar spacing of 0.21 nm is close to the (100) facet of graphite.37 This partial crystallinity is possibly due to plasma etching of the polymer matrix at the plasma/liquid interface.38

Figure 3. Morphology, composition, and structure of PDFONs. (a) HRTEM images of PD-FONs. The red circles mark those with observable lattice fringes. Scale bar: 10 nm. (b) Selected examples of crystalized PD-FONs in (a). (c) Element content (atomic ratio) in dopamine and PD-FONs analyzed by XPS. DA: dopamine. (d-e) Expanded XPS spectra of PD-FONs including C 1s (d) and N 1s (e). B. E.: binding energy. (f) Fourier transforming infrared (FT-IR) spectra (2000~500 cm-1) of dopamine and PD-FONs. X-ray photoelectron spectroscopy (XPS) reveals that the plasma does not introduce any exogenous element to the product (Figure S6a). However, the element content in PDFONs is quite different from that in dopamine, featured by a dramatic increase of O content accompanied by a decrease of both C and N contents (Figure 3c). This is probably due to oxygen doping during plasma-induced oxidation. In the expanded image of the C 1s peak, the signals at 284.7 eV and 286.2 eV are from of C-C, C=C, CO, and C-N functional groups (Figure 3d and Figure S6b). The peak at 288.4 eV is assigned to quinone groups (C=O), which is consistent with the signal at 531.0 eV (O=C) in O 1s region (Figure S6c).39 This indicates the oxidation of phenol groups to quinone groups, which is featured as the initiation of dopamine polymerization.40 The N 1s region is fitted with two peaks at 400.0 eV and 401.9 eV, which can be assigned to cyclized R2NH (possibly from 5,6dihydroxyindole (DHI))41 and unreacted RNH2 groups respectively (Figure 3e). These results suggest that the formation of PD-FONs follows the “eumelanin” model that envisages a polymeric skeleton based on the oxidation and cyclization of dopamine into DHI units (Figure S7). Results from Fourier transforming infrared (FT-IR) spectroscopy further support the observations in XPS. The


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Analytical Chemistry broad band in the range of 3500~3000 cm-1 includes the stretching vibrations of N-H (3500~3300 cm-1), O-H (3500~3200 cm-1), and Ar-H (3100~3000 cm-1) groups (Figure S8). The peaks at 1600 cm-1 and 1502 cm-1 confirm the benzene ring-based skeletons of PD-FONs. Importantly, four peaks appear in the FT-IR spectrum of PD-FONs yet are missing in that of dopamine (Figure 3f). The peak at 1635 cm-1 is assigned to C=O stretching vibrations of quinone groups, which agrees well with the XPS data.42 The peak at 1553 cm-1 and 665 cm-1 originate from C=C stretching vibrations and C-N-C in plane bendings in the indole structures as earlier proposed.43 The strong peak at 1050 cm-1 indicates the presence of C-O-C groups in PD-FONs. Therefore, we hypothesize that plasma-induced O insertions mainly appear as C-O-C linkers between adjacent rings in PD assemblies (Figure S9). Collectively, these characterizations validate the oxidation of dopamine and the polymeric structures in PDFONs. Time-resolved photoluminescence measurements reveal that PD-FONs possess biexponential decay property with an average lifetime of 5.03 ns (Figure 4a).44 The quantum yield (QY) of PD-FONs is 0.8% using quinine sulfate as the reference (Figure S10), which is comparable to reported values (1.0 %).28 The low QY is probably due to nonradiative dissipation from the eumelanin-like structures of PD.45 PD-FONs are stable against photobleaching with no decrease in fluorescence intensity during continuous light illumination for 2 h (Figure 4b). Besides, they show identical absorption spectra and approximately 10% increase in fluorescence intensity after preservation for 3 months under ambient conditions (Figure 4c-4d).

Cytotoxicity of PD-FONs. We test the in vitro biocompatibility of PhFONs applying PD-FONs as the example. In the 24 h CCK-8 assay, PD-FONs are nontoxic to 3T3 fibroblasts at concentrations up to 120 μg/mL, with cell viability higher than 90% and negligible damages to cell morphology (Figure 5a-5b). Besides, they show no damage to human erythrocytes at concentrations up to 150 μg/mL (Figure 5c), thus are suitable for blood-contacting applications.

Figure 5. Cytotoxicity of PD-FONs. (a) Cell viability (CCK8) assay using NIH 3T3 cells. (b) Bright field images showing the morphology of normal cells and cells treated with 100 μg/mL PD-FONs for 24 h. Scale bar: 50 μm. (c) Hemolytic property of PD-FONs tested by incubation with human red cells. Cell imaging applications of PhFONs. To underscore the analytical applications, we study the uptake and imaging behaviors of PhFONs in 3T3 fibroblasts using confocal laser scanning microscope (CLSM). Under 405 nm excitation, the cells display intense fluorescence after the incubation with PhFONs, indicating the successful uptake of these nanoparticles. The fluorescence appears yellow for PP-FONs and green for the other three FONs (Figure S11), which is consistent with their emission spectra. Interestingly, for PD-FONs (3.36±0.21 nm) and PN-FONs (7.80±1.32 nm), the fluorescence is located in both the cytoplasm and the nucleus, while for PP-FONs (17.62±3.20 nm) and PG-FONs (12.82±3.28 nm), it is observed only in the cytoplasm. Nanoparticles smaller than 9 nm can enter cell nucleus through passive nucleus diffusion.46 Thus, we attribute this difference to the size effect of these FONs. To further confirm this hypothesis, we synthesize PD-FONs with even larger size of 32.04±3.08 nm in pH 8.8 Tris-HCl buffer (Figure 6a) and investigate their intracellular distribution. As expected, the access into cell nucleus is limited for the larger PD-FONs (Figure 6b), which differs from their smaller counterpart (Figure 6c-6d). Such sizedependent nucleus-targeting property of PhFONs allows many functionalities to be tailored such as cell nucleus imaging, nucleus drug delivery, and theranostics.

Figure 4. Optical property of PD-FONs. (a) Time-resolved fluorescence spectrum of PD-FONs at 340 nm excitation. (b) Fluorescence intensity of PD-FONs under continuous illumination (excitation at 400 nm) for 2 h. (c-d) UV-Vis absorption (c) and fluorescence emission spectra (excitation at 400 nm, inset: fluorescence intensity) (d) of freshly-prepared PD-FONs and those preserved for 3 months.



Figure 6. Size-dependent nucleus access of PD-FONs. (a) TEM images of PD-FONs synthesized in pH 8.8 Tris-HCl buffer. Scale bar: 100 nm. (b) CLSM images of cells treated with the larger PD-FONs. Scale bar: 50 μm. (c) HRTEM images of PD-FONs synthesized in pH 7.0 Tris-HCl buffer. Scale bar: 20 nm. (d) CLSM images of cells treated with the smaller PD-FONs. Scale bar: 50 μm. The excitation-dependent emission spectra of PhFONs make them suitable for multicolor bioimaging. For demonstration, PD-FONs show green and yellow colors when excited by 405 nm and 488 nm lasers respectively (Figure S12). Accordingly, the cells exhibit green and yellow fluorescence after the uptake of PD-FONs (Figure 7a-7b). The colocalization of PD-FONs and RedDot 1 (a fluorescent dye staining the cell nucleus as red) further confirms their access into the cell nucleus (Figure 7c-7d), which is indicated by the overlap of these two signals in the nucleus region (Figure 7e-7f). We track the cellular uptake process of PD-FONs by real time CLSM imaging (Figure 8 and Figure S13). PD-FONs distribute around the cells upon their immediate addition into the culture medium (Figure 8a). Discernable fluorescence signals appear in the cytoplasm 15 min after treatment, indicating the penetration of PD-FONs through the cell membrane (Figure 8b). PD-FONs start entering the cell nucleus 1 h post-treatment, and reside there thereafter (Figure 8d-8f). As time prolongs to 2 h, the fluorescence becomes increasingly stronger inside the cells but weaker outside, which is due to the continuous uptake and accumulation of PD-FONs. After incubation for 2 h, the background signal (free PD-FONs in the solution) becomes nearly negligible, suggesting that most of PD-FONs have been harvested by the cells. These results uncover the time-dependent, nonspecific internalization of PD-FONs into subcellular structures.

Figure 7. PD-FONs for multicolor imaging of live cells. (ab) Confocal laser scanning microscope (CLSM) images of NIH 3T3 fibroblasts loaded with PD-FONs under 405 nm (a) and 488 nm (b) excitation. (c-d) Colocalization of RedDot 1 and PD-FONs under 405/543 nm (c) and 488/543 nm (d) excitation. Scale bar: 50 μm. (e-f) Fluorescence intensity of different channels along the paths (indicated as white arrows) in (c) and (d).

Figure 8. Real time tracking of the uptake and distribution of PD-FONs in live cells. CLSM images show NIH 3T3 fibroblasts at different time points after PD-FONs treatment. (a) 0 min. (b) 15 min. (c) 30 min. (d) 60 min. (e) 90 min. (f) 120 min. Cells are excited by 405 nm laser. Scale bar: 50 μm.


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CONCLUSION In summary, a new class of FONs based on polymerized phenolic compounds, PhFONs, are synthesized by plasma treatment at the aqueous interface. As a representative, PD-FONs show excellent stability and biocompatibility. Multicolor imaging of live cells is achieved using these particles as fluorescence probes. Additionally, their sizedependent access into cell nucleus clearly suggests a potential to develop nucleus-targeting staining agents. Owing to their reactive surface groups, these nanoparticles can be easily modified with other structures such as bioactive molecules or inorganic nanocomposites to realize autonomous functionalities. Future efforts will be devoted to improving the throughput of this synthesis method and exploring other applications using these FONs.

ASSOCIATED CONTENT Supporting Information Fluorescence spectra of PD-FONs (Figure S1). Synthesis of FONs from other phenolic compounds (Table S1 and Figure S2). Effect of droplet size on the synthesis of PD-FONs (Figure S3). XRD pattern and Raman spectrum of PD-FONs (Figure S4). HRTEM images of PD-FONs (Figure S5). XPS spectra of PD-FONs and energy assignments (Figure S6). Proposed mechanism of PD-FONs formation (Figure S7). Complete FTIR spectrum of PD-FONs (Figure S8). Proposed mechanism of plasma-induced oxygen doping in PD-FONs (Figure S9). Quantum yield measurement of PD-FONs (Figure S10). Cell imaging using different PhFONs (Figure S11). CLSM images of PD-FONs solution (Figure S12). Real time CLSM images of cells treated with PD-FONs (Figure S13).

AUTHOR INFORMATION Corresponding Author * [email protected] (Xingyu Jiang)

ORCID Xingyu Jiang: 0000-0002-5008-4703

Notes The authors declare no competing financial interests.

ACKNOWLEDGMENT We thank the National Key R&D Program of China (2017YFA0205901), the National Natural Science Foundation of China (21535001, 81730051, 21761142006), and the Chinese Academy of Sciences (QYZDJ-SSW-SLH039, 121D11KYSB20170026) for financial support.

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