Highly Hydrophilic, Two-photon Fluorescent Terpyridine Derivatives

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Highly Hydrophilic, Two-photon Fluorescent Terpyridine Derivatives Containing Quaternary Ammonium for Specific Recognizing Ribosome RNA in Living Cells Wei Du,† Hui Wang,† Yingzhong Zhu,† Xiaohe Tian,*,‡ Mingzhu Zhang,† Qiong Zhang,† Senio Campos De Souza,∥ Aidong Wang,⊥ Hongping Zhou,† Zhongping Zhang,§ Jieying Wu,*,† and Yupeng Tian*,† †

Anhui Province Key Laboratory of Chemistry for Inorganic/Organic Hybrid Functionalized Materials, ‡School of Life Science, and School of Chemistry and Chemical Engineering, Anhui University, Hefei 230601, P. R. China ∥ Department of Chemistry, University College London, London WC1H 0AJ, U.K. ⊥ School of Chemistry and Chemical Engineering, Huangshan College, Huangshan University, Huangshan 245041, P. R. China §

S Supporting Information *

ABSTRACT: A two-photon fluorescent probe (J1) that selectively stains intracellular nucleolar RNA was screened from three water-soluble terpyridine derivatives (J1−J3) with quaternary ammonium groups. The photophysical properties of J1−J3 were systemically investigated both experimentally and theoretically, revealing that J1−J3 possess large Stokes shifts and the two-photon absorption action cross sections range from 38 to 97 GM in the near-infrared region. This indicates that J1 could specifically stain nucleoli by targeting the nucleolar rRNA from the recognition experiments in vitro, the two-photon imaging experiments, and the stimulated emission depletion in vivo. The mechanism of action in which J1 binds to the nucleolar rRNA was researched via both experiments and molecular modeling. The high binding selectivity of J1 to nucleolar RNA over cytosolic RNA made this probe a potential candidate to visualize rRNA probe in the living cells. KEYWORDS: water-soluble, two-photon, rRNA, molecular docking techniques, STED

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complexes15 have been synthesized to overcome these problems and applied for RNA imaging in the living cells. Recently, our group successfully designed ratiometric twophoton fluorescent nucleic acid-selective probes,22 which possessed several advantages over one-photon fluorescent probes, such as minimal photodamage to the livings cells, good tissue penetration, and negligible interference from the background autofluoresence.17−21 However, these reported probes were unable to discriminate nucleolar RNA (rRNA) from cytosolic RNA (mRNA and tRNA). Although fluorescent mRNA probe has recently emerged,23 specific rRNA targeting small molecular system is still rarely reported. Guo et al. reported an indole-based cyanine derivate as a rRNA-selective two-photon fluorescent probe;24 nonetheless, the mechanism of selective rRNA was not presented. Synthesizing novel specific rRNA probes and understanding its binding mode are very valuable to deeply understand the cause of the probes staining with rRNA and to guide the design of new rRNA probes.

INTRODUCTION RNA is one of the first self-replicating molecules leading to the origins of life.1 RNA molecules including transfer RNA (tRNA), messenger RNA (mRNA), and ribosomal RNA (rRNA) in the living cells are well known for a wide variety of functions including transportation and interpretation of genetic information, regulation of gene expression, and some important biological catalytic roles.2−8 In particular, rRNA is the most abundant type of RNA in the cells that can form ribosome with protein in nucleoli.9 Designing and synthesizing specific rRNA fluorescent probes are beneficial for observing the status and activities of rRNA in nucleoli under microscopy as well as in extended super-resolution nanoscopy (e.g., stimulated emission depletion (STED) microscopy, structured illumination microscopy). However, RNA probes for living-cell bioimaging are rare due to their poor nuclear membrane penetration.10,11 SYTO RNA-select is the only commercially available probe for RNA bioimaging in whole living cells at the moment. Nevertheless, it has not been widely used in the research community due to unknown molecular structure and a relatively higher phototoxicity under one-photon laser source.12,13 In the recent years, single-photon fluorescent probes including organic fluorescent compounds14 and metal © 2017 American Chemical Society

Received: June 6, 2017 Accepted: August 1, 2017 Published: August 1, 2017 31424

DOI: 10.1021/acsami.7b08068 ACS Appl. Mater. Interfaces 2017, 9, 31424−31432

Research Article

ACS Applied Materials & Interfaces

Figure 1. (A) Synthetic procedure for target molecules J1−J3. (B) Chemical structures of J1−J3. (C) The spherical plots for J1 and J2 obtained from single-crystal structure determination.

photon microscopy (TPM) and stimulated emission depletion (STED) micrographs indicated that J1 could stain nucleoli in the living cells through interaction with rRNA. The mechanism of action was further testified by molecular docking techniques and 1H NMR. It is suggested that J1 may be a potential candidate for specific rRNA labeling in the living cells.

The quaternary ammonium derivatives were selected for designing the RNA probes because water solubility is the main limitation of organic molecular probe for bioassays.16,25 Quaternary ammonium salts have similar water solubility to that of inorganic salts, which substantially improves the water solubility of organic probes. They are positively charged and interact easily with negatively charged substances in the living organism such as RNA.26,27 The terpyridine as a hydrophobic group is easily immobilized with the groove of RNA by the π−π interactions,28 which can bind DNA or rRNA. The synergistic effect of quaternary ammonium salts and terpyridine groups might be assembled into a new selective rRNA probe. In this work, a novel two-photon fluorescent rRNA probe was screened from a series of terpyridine derivatives: 2,2′:6′,2″ (J1), 3,2′:6′,3″ (J2), and 4,2′:6′,4″ terpyridine (J3) were designed and synthesized into A−π−A′ configurations, which comprised of an aniline with two quaternary ammoniums (acting as acceptors, A), phenyl groups (acting as π-bridges), and terpyridine groups (acting as another acceptors, A′). Recently, the terpyridine (β-, γ-) derivatives with thiophene of our group were reported, but they have no hydrophilia, which would highly limit their application in the living organism.29 The quaternary ammonium groups in the designed molecules can greatly enhance the water solubility, and their positive charges can interact with the negatively charged RNA in the living organism. Moreover, small molecules with A−π−A′ configuration could be designed into two-photon fluorescent probes because quadrupolar configuration molecules can show a two-photon absorption (TPA) cross section.30,31 The three terpyridine derivatives have different electron-withdrawing effects and different steric effects for their neighboring molecules due to the different localization of the nitrogen atoms. J1 can specifically recognize the rRNA through the π−π interactions between the rigid terpyridine groups and the hydrophobic groove. In this article, the two-photon fluorescent properties and the biological application of these three terpyridine derivatives were investigated systematically. Two-

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EXPERIMENTAL SECTION

Measurements and Instruments. All of the chemicals were commercially available and the solvents were purified through conventional methods before using them. DNA employed was calf thymus DNA (ct-DNA) and RNA employed was ribonucleic acid diethylaminoethanol salt Type IX from Sigma-Aldrich for the in vitro experiments. Fourier transform infrared (FT-IR) spectra were obtained on NEXUS 870 (Nicolet) spectrophotometer in the 4000−400 cm−1 region, with samples prepared as KBr pellets. 1H NMR and 13C NMR spectra were recorded on a Bruker Avance 400 and 100 MHz spectrometer (tetramethylsilane as internal standard in NMR) using D2O and DMSO-d6 as solvents. Matrix-assisted laser desorption ionization time-of-flight mass spectrometry was carried out on Bruker Autoflex III Smartbeam and electrospray ionization mass spectrometry (ESI-MS) was confirmed on a Finnigan LCQ spectrometer. UV−vis absorption spectra were performed by the UV-265 spectrophotometer. Fluorescence measurements were recorded on a Hitachi F-7000 fluorescence spectrophotometer. For time-resolved fluorescence measurements, the fluorescence signals were collimated and focused onto the entrance slit of a monochromator with the output plane equipped with a photomultiplier tube (HORIBA HuoroMax-4P). The ζ-potentials of the samples were measured using a Nano Zeta Potential Analyzer (Delsa 440SX Beckman Coulter Limited). Two-Photon Action Cross Section. Two-photon absorption (TPA) cross sections of all of the ligands were obtained by the twophoton excited fluorescence (TPEF) method with femtosecond laser piles and a Ti:sapphire system (680−1080 nm, 80 MHz, 140 fs) as the light source. The two-photon action cross section σΦ values were determined by the following equation

σ Φ = σref Φref 31425

cref nref F c n Fref DOI: 10.1021/acsami.7b08068 ACS Appl. Mater. Interfaces 2017, 9, 31424−31432

Research Article

ACS Applied Materials & Interfaces

Figure 2. (A) UV−vis absorption and single-photon fluorescent spectra for J1−J3 in PBS (pH 7.4). (B) Molecular orbitals for J1−J3 in their ground state (S0) optimized geometries from time-dependent density functional theory calculations. (C) Two-photon absorption cross sections of J1−J3 at different excitation wavelengths from 680 and 1000 nm in PBS.

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Here, the subscripts ref represents the reference molecule (fluorescein). Φ stands for the quantum yield, n is the refractive index, F is the integral area under the corrected emission spectrum, and c is the concentration of the solution (mol/L). The σref value of reference was cited from the literature.32 All of the three compounds were successfully synthesized according to the routes, shown in Figure 1 and Scheme S1, and the 1H NMR and 13 C NMR spectra of B1 and J1 are shown in Figures S1−S12. Preparation of J1. B1 (2 g, 3.72 mmol) together with 60 mL 40% aqueous trimethylamine was sealed in a 50 mL Teflon reactor autoclave and heated to 90 °C for 10 h. After cooling to room temperature, the reserved trimethylamine, tetrahydrofuran, and water were removed under vacuum. The crude product was washed three times using dichloromethane and dried in a vacuum. The product (1 g, 1.52 mmol) was fully dissolved in 20 mL methanol in a 50 mL roundbottom flask and AgNO3 (0.54 g, 3.04 mmol) along with 6 mL acetonitrile was added dropwise in the dark. The mixture was allowed to react for 30 min at room temperature. The reaction was filtered over diatomite, and the removal of solvent in vacuum yielded yellow solids. The preparations of J2 and J3 were similar to that of J1. Yield: 89%. Mp: 197 °C. FT-IR (KBr cm−1): 3430 (s), 3023 (w), 1591 (s), 1523 (m), 1384 (s), 1203(m), 813 (m). 1H NMR (D2O, 400 MHz, ppm) δ: 8.12 (t, J = 20.9 Hz, 2H), 7.89 (d, J = 7.9 Hz, 2H), 7.68 (t, J = 7.6 Hz, 2H), 7.49 (s, 2H), 7.31−7.15 (m, 4H), 6.34 (d, J = 8.3 Hz, 2H), 3.63 (dd, J = 26.1, 18.2 Hz, 4H), 3.46−3.27 (m, 4H), 3.15 (s, 18H). 13C NMR (100 MHz, D2O): δ = 153.89, 153.79, 147.93, 147.56, 145.86, 138.00, 127.57, 125.23, 124.24, 121.95, 116.20, 112.08, 61.05, 53.50, 43.89. Anal. calcd for C31H40N8O6: C, 59.99; H, 6.50; N, 18.05%. Found: C, 60.08; H, 6.34; N, 18.23%. ESI-MS: calcd: M = 620.31, found: [M − 2NO3−]2+/2 = 248.42. J2. Yield: 74%. Mp: 191 °C. FT-IR (KBr cm−1): 3412 (m), 3026 (w), 1600 (s), 1525 (m), 1382 (s), 1205 (m), 786 (m). 1H NMR (D2O, 400 MHz, ppm) δ = 8.06 (s, 2H), 8.02 (d, J = 4.0 Hz, 2H), 7.47 (d, J = 7.2 Hz, 2H), 6.90 (dt, J = 15.9, 7.9 Hz, 2H), 6.71 (d, J = 7.7 Hz, 2H), 6.51 (s, 2H), 5.85 (d, J = 7.5 Hz, 2H), 3.61 (t, J = 4.8 Hz, 4H), 3.32 (t, J = 5.1 Hz, 4H), 3.07 (s, 18H). 13C NMR (100 MHz, D2O): δ = 153.36, 153.26, 147.92, 147.55, 146.26, 134.07, 127.89, 125.21, 124.24, 120.36, 115.62, 112.27, 61.04, 53.56, 43.87. Anal. calcd for C31H40N8O6: C, 59.99; H, 6.50; N, 18.05%. Found: C, 60.12; H, 6.38; N, 18.13%. ESI-MS: calcd: M = 620.31, found: [M − 2NO3−]2+/2 = 248.25. J3. Yield: 68%. Mp: 194 °C. FT-IR (KBr cm−1): 3412 (m), 3026 (w), 1595 (s), 1523 (m), 1352 (s), 1203 (m), 812 (m). 1H NMR (D2O, 400 MHz, ppm) δ: 7.96 (d, J = 5.1 Hz, 4H), 7.13 (d, J = 5.1 Hz, 4H), 6.77 (s, 2H), 6.69 (s, 2H), 5.91 (d, J = 8.0 Hz, 2H), 3.45 (s, 4H), 3.20 (d, J = 7.1 Hz, 4H), 3.13 (s, 18H). 13C NMR (100 MHz, D2O): δ = 152.78, 148.51, 147.87, 146.29, 145.76, 137.47, 127.70, 125.75, 125.10, 121.28, 116, 112, 61, 53.57, 43.80. Anal. calcd for C31H40N8O6: C, 60.05; H, 6.39; N, 18.18%. Found: C, 60.08; H, 6.34; N, 18.23%. ESI-MS: calcd: M = 620.31, found: [M − 2NO3−]2+/2 = 248.25.

RESULTS AND DISCUSSION Synthesis and Characterization. As shown in Scheme S1, the structural difference of the three compounds is the localization of the nitrogen atom within the terpyridine moiety due to the different acetylpyridine in the synthetic process. Teflon reactor autoclave is used in the synthesis of J1−J3, which decreases the loss of trimethylamine in their confined space and shortens the heating reaction time. The synthesis of J1 with a high yield (89%) and that of J2 and J3 with lower yields of 79 and 68% is verified. All of the compounds are obtained at mild reaction temperature to ensure that they are handled efficiently. All of the compounds are characterized by FT-IR, 1H NMR, 13C NMR, mass spectra, and elemental analysis. Crystal Structures. The crystal data collection and refinement parameters of J1 and J2 are listed in Table S1. The selected distances and angles of bonds are given in Table S2. J1 and J2 are crystallized in the triclinic crystal system with the P1̅ space group. As displayed in Figures 1C and S13, the dihedral angle between the benzene ring (P1) and the middle pyridine ring (P2) in J1−J3 is 32.00, 8.62, and 8.42°, respectively, indicating that the planarity of J2 and J3 is better than that of J1. These planarities suggest that J2 and J3 have a delocalized π-electron systems, which favors a two-photon fluorescence response. In addition, Figure S14 shows that the dihedral angles between the two side pyridines of J1−J3 are 6.44, 25.05, and 23.52°, respectively. The terpyridine of J1 is almost planar; in contrast, the terpyridine of J2 and J3 shows a twisted configuration, and the non-coplanar of the side pyridines of J2 and J3 results in a large steric hindrance in the intermolecular interactions. Optical Properties. Figure 2A shows that the UV−vis absorption spectra in phosphate-buffered saline (PBS) of J1−J3 all of the displayed two absorption bands at about 290 and 325 nm, respectively. The higher energy band at about 290 nm is assigned to the πterpyridine−πterpyridine* transitions,33 whereas the lower energy band around 325 nm is assigned to the intramolecular charge transfer (ICT). Figure 2B and Table S3 suggest that the electron density mainly focuses on terpyridine groups at highest occupied molecular orbital and on the aniline groups at lowest unoccupied molecular orbital around 325 nm, which confirms the ICT from the terpyridine to aniline group and quaternary ammonium groups. The electron mobility trends of J1−J3 from the terpyridine part to aniline group and quaternary ammonium groups results from the strong electronwithdrawing ability of the two quaternary ammonium salts. As 31426

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ACS Applied Materials & Interfaces Table 1. Photophysical Properties for J1−J3 in PBS (pH 7.4) J1 J2 J3 a

λmaxa

extinction coefficient (×104 mol−1 L cm−1)

λmaxb

quantum yields

τ (ns)c

Δνd

292 325 283 325 282 331

2.62 2.62 1.72 3.25 1.05 1.70

452

0.19

6.08

8645.34

441

0.29

4.18

8093.49

479

0.15

6.63

9334.65

Peak position of the absorption band. bEmission wavelength of single photon. cFluorescence lifetime (ns). dStokes shift (nm−1).

Figure 3. (A) Absorption spectra against the concentration of J1 from 10 to 100 μM and (B) the plot of absorption against the concentration of J1 from 10 to 100 μM.

fluorescent molecules and have potential application in TPA bioimaging. The water solubility of J1−J3 is also investigated via UV−vis absorption spectral determination. As shown in Figures 3 and S18, J1−J3 are soluble in PBS, and the absorbance is linear to the concentration in the region of 10−100 μM, suggesting that J1−J3 possess excellent water solubility, which is a significant advantage for biological research. Specific RNA Response in Vitro. In solution fluorescence measurements, J1 exhibited RNA “light-switch” effect in both single-photon fluorescent and two-photon fluorescent spectra (Figure 4A−C). In the presence of 50:1 RNA-to-J1 ratio, the single-photon fluorescent intensity of the probe shows an 3.8fold enhancement, association constant K = 1.8 × 104 M−1. In contrast, at the same ratio, the two-photon fluorescent intensity has a 6-fold enhancement. The maximum two-photon action cross section of J1 shows an approximately 4-fold enhancement with bonding to RNA. It indicates that J1 has a turn-on effect with RNA in the two-photon excited fluorescence (TPEF) and the single-photon-excited fluorescence (SPEF), suggesting J1 to be a potential candidate for SPEF and TPEF bioimaging. In contrast to RNA, the “light-switch” effect of J1 with DNA in vitro is weaker (Figure S19), the fluorescence intensity shows only a 3-fold enhancement with the presence of DNA at SPEF, and 4-fold enhancement with the presence of DNA at TPEF spectra. It is further confirmed by the ethidium bromide (EB) displacement assays (Figure S20). The EB is a commercial intercalator for DNA and emits a strong fluorescent signal when associated with DNA. If another DNA intercalator could replace EB from the EB−DNA system, the fluorescence would be quenched due to free EB molecule rapidly quenched in

shown in Figure S16, J1−J3 dissolved in different polar solvents are excited by an identical excitation wavelength (J1: λex = 324; J2: λex = 325; J3: λex = 332). With increasing solvent polarity, one-photon fluorescent spectra of J1−J3 exhibits an obvious red shift. This may be caused by the different polarity of the solvent, which changes the interaction between the solvents and solutes. The fluorescence spectra of J3 exhibit a greater red shift compared that of J1 and J2. This might be a result of the stronger electron-accepting ability of the 4-pyridyl group and the better plane structure of the whole molecule, which consequently increase the ICT of the molecule and, as result, the emission maximum becomes shifted.34 J1−J3 exhibit a large Stokes shift, which can eliminate the influence of excitation spectra. The data of quantum yields (Φ) and fluorescence lifetimes (τ) of J1−J3 in PBS are collected in Table 1. A decreasing quantum yield is observed from J2 (29%) → J1 (19%) → J3 (15%). The two-photon action cross sections (σΦ) of J1−J3 from 680 to 900 nm are shown in Figure 2C. There is no linear absorption in the range of 680−900 nm for J1−J3, implying that there is no energy level corresponding to the singleelectron transition in this spectral range. To further verify the two-photon excited processes, the linear dependence of output fluorescence intensity (Iout) on the square of input laser power (Iin), which is expressed by the log−log plot and shown in Figure S4. The gradient of linear-fitted results is 2.11, 2.16, and 1.89, which suggests a two-photon excitation mechanism. As shown in Table S17, the optimal excitation wavelengths of J1− J3 are at 700 nm and the maximum two-photon action cross section of J1−J3 is 39, 72, and 97 GM, respectively. These results suggest that all of the compounds are two-photon 31427

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Figure 4. (A) SPEF changes of 10 μM J1 with 0.5 mM RNA, inset: fluorimetric titrations curves of 10 μM J1 by 0.5 mM RNA. (B) TPEF changes of 0.5 mM J1 with 25 mM RNA. (C) Two-photon absorption action cross section changes of 0.5 mM J1 with 25 mM RNA. (D) Partial 1H NMR titration spectra for J1 + RNA for in D2O. (E) Models obtained by molecular modeling for the interaction of J1 with RNA fragment.

water system. There is no changed fluorescence intensity, which indicates that J1 has a pretty weak interaction with DNA. As shown in Figure S21, the single-photon fluorescent intensity and two-photon fluorescent intensity of J2 and J3 has no enhancement in spectra, which indicates that neither J2 nor J3 has enough interaction with DNA and RNA. To confirm that J1 is encapsulated by RNA, the 1H NMR titration experiments are carried out in D2O. As shown in Figure 4D, the addition of 2 equiv of RNA generated significant upfield shifts of the terpyridine protons Ha and Hb, which demonstrates the presence of π−π stacking interactions between terpyridine fragments of J1 and RNA.35 Figure S22 shows that 10 μM J1 exhibits a positive ζpotential 9.76 mV and 100 μM RNA exhibits a negative ζpotential −20.84 mV. After they interact with each other, the ζpotential is −5.51 mV, between 9.76 and −20.84 mV. It is indicated that J1 is positively charged and could easily interact with the negatively charged RNA, making it suitable probe for RNA detection in the living organisms. Molecular docking techniques are valuable for understanding the molecular design and the mechanistic research about the nature of RNA interaction (Figures 4E and S23). The docking techniques calculate the interaction of J1 with double-stranded RNA and single-stranded RNA. CDOCK ENERGY of J1 and double-stranded RNA (−37.8431 kJ/mol) fragment is smaller than that of J1 and single-stranded RNA fragment (−19.9767 kJ/mol), which suggests that J1 has a more stable interaction with double-stranded RNA and not with single-stranded RNA. The fact that nucleoli rRNA is found as a double-stranded RNA further explains the high affinity of J1 toward nucleoli rRNA

over cytosolic RNA (e.g., mRNA). In addition, the molecular simulation indicates the presence of π−π interactions between the side pyridine ring of J1 and G21 of RNA for the plane terpyridine of J1. These π−π interactions are not found with J2/J3 and RNA due to the twist in the steric configuration of the side pyridines at J2/J3 (Figure S14). It is further explained that J1 could specifically stain nucleoli RNA in vivo and in vitro. The mechanism of the “light-switch” effect of J1 binding to RNA is proposed as follows. The weak emission of J1 in water could be attributed to the rapid nonradiative decay of fluorophore. When interacting with RNA, the rotation of C− C single bond between pyridine rings is restricted by the steric effect from the π−π interactions, which effectively decrease the nonradiative transition and enhance the fluorescence intensity. We displayed a small molecular system that could concentrate within cell nucleoli, which might offer a tool to study the dynamic alternation of rRNA in the living system. Application for Biological Imaging. Because high cell viability is essential for biological applications, the cytotoxicity of J1−J3 is evaluated by the MTT assay in HepG2 cells (shown in Figure S24). J1−J3 exhibit high cell viability after 24 h of incubation with the concentrations between 10 and 50 μM of all of the compounds. The results demonstrate that all of the compounds show low toxicity in the living cells. To assess the cell entry properties, HepG2 cell is used as a model, whereby 20 μM of J1−J3 are incubated for 30 min followed by confocal microscopy imaging after washing with PBS (×3). As shown in Figure 5, J1 could enter the cell nuclear region within the short-time incubation and display obvious luminescence under one-photon and two-photon laser 31428

DOI: 10.1021/acsami.7b08068 ACS Appl. Mater. Interfaces 2017, 9, 31424−31432

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the fluorescence intensity of Syto 9 decreases to 34% after 150 s of irradiation, whereas the fluorescence intensity of J1 only decreases to 96% after 250 s of irradiation. The finding suggests that photostability of J1 is much higher than that of Syto 9. To understand the cellular uptake mechanism of J1, the HepG2 cells are incubated for 20 min at 4 °C, then incubated with 20 μM J1 for another 20 min at 4 °C, washed, and imaged immediately. In comparison with the imaging of 37 °C, the lowtemperature bioimaging (Figure S26) shows that no luminescence is observed after the cells are incubated with J1 at 4 °C. Cell metabolism and physiological activity would be suppressed under low-temperature conditions; thus, the results suggest that J1 entered cells via an energy-dependent pathway (e.g., endocytosis, active transport), in agreement with our above hypothesis that J1 might interact with nucleoli rRNA via specific intracellular sorting mechanism. In addition, the fluorescence intensity of J1−J3 did not change at pH 6−8 (Figure S27); therefore, bioimaging would not be affected in the living cells. To further confirm whether J1 selectively detected DNA or RNA, a digestion test is carried out using deoxyribonuclease (DNase) and ribonuclease (RNase), as shown in Figure 7C. RNase is a special enzyme that only hydrolyzes RNA and does not disturb DNA and vice versa. After RNase digestion, the two-photon fluorescent signal of J1 in nucleoli is significantly diminished compared with that in the untreated samples. In contrast, following DNase digestion, the two-photon fluorescent signal of J1 in the nucleoli and cytoplasm still remain. These results indicate that J1 could selectively stain RNA in nucleoli. In a parallel experiment, flow cytometry as an alternative tool to access the luminescence change following DNAse/RNAse addition. As shown in Figure 7D, the luminescence does not change after DNase digestion but is decreased after RNase digestion, which again indicates that J1 could selectively stain RNA. To prove that J2 and J3 are not able to interact with RNA, the uptake properties of J1−J3 are evaluated in fixed HepG2 cells. Because plasma membrane and nuclear membrane become permeable after fixation, probes can easily penetrate and come in direct contact with the nucleoli. As shown in Figure S28, J1 clearly stains nucleoli in the fixed cells, but J2 and J3 show similar distribution patterns in the fixed cells, further supporting the higher affinity of J1 interaction with RNA over J2 and J3 compounds. The successful nucleolar rRNA labeling using J1 demonstrates stable fluorescence and high specificity; therefore, we further extend its utility in biosensing to display nucleolar ultrastructure under stimulated emission depletion (STED) nanoscopy in the living cells. The STED micrographs clearly show nucleolus in different regions of cells (1−3) with higher resolution (Figure 8) compared with normal confocal microscopy (Figures 5 and 6). J1 displays superb signal-tonoise ratio and indicates clear fibrillar center (asterisk, fc) and dense fibrillar component (arrow, dfc). It is worth noting that RNA-rich dfc indicated a strong fluorescence, whereas fc, which mainly possesses DNA, shows negative patches. Such results further confirm the high affinity of nucleolar rRNA using J1 and compatibility under STED nanoscopy in the living cells. This utilization is crucial because super-resolution applications are mainly limited in fixed or transfected samples, as well as suffering the risk of alternation the very ultrastructure using concentrated markers during such process.36,37 Thus, J1 might

Figure 5. Confocal fluorescence images of J1−J3 in HepG2 cells. Single-photon imaging (green, λex = 405 nm), two-photon imaging (red, λex = 720 nm), bright field (gray), and merge were shown. Scale bar: 20 μm.

excitation, whereas J2 and J3 show general cytosolic distribution with no selectively on specific organelles. According to the bright field image, the morphology and bright dots observed with J1 correspond to the dark-phase nucleolar regions. Moreover, two-photon confocal fluorescence images of J1 in Hela (human cervical cancer cell), MCF-7 (human breast cancer cell), and MRC-5 (fetal lung fibroblast) have been carried out (Figure S25), and these experiments show that J1 could enter the nuclear region and stain nucleoli for these three kind of cells. To further confirm the subcellular localization of J1, colocalization studies of J1 with Syto 9 (RNA-select commercial dye) and Nuc-red (DNA-select commercial dye) are tested in HepG2 cells using TPM (Figure 6). The results show that J1

Figure 6. Costaining of J1 (λex = 800 nm) with RNA-specific dye Syto 9 and DNA-specific dye Nuc-red. Scale bar: 20 μm.

could readily enter the living HepG2 cells effectively, and predominantly emission is observed similarly as in Syto 9 (Pearson’s Rr = 0.90) in the nuclear region. It also clearly indicates a high selectivity for nucleoli RNA over nuclear DNA (Nuc-red, Rr = 0.22). In addition, the photostability of J1 is examined in comparison to commercially available Syto 9 in the living HepG2 cells via photobleaching experiments. The initial fluorescence intensities from the first scans of costained cells are normalized, and the percentages of the weakened fluorescence signal are calculated. As shown in Figure 7A,B, 31429

DOI: 10.1021/acsami.7b08068 ACS Appl. Mater. Interfaces 2017, 9, 31424−31432

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Figure 7. (A) Confocal fluorescence bioimages of 10 μM J1-pretreated HepG2 cells incubated with Syto 9 for another 20 min. Green channel for Syto 9 (λex = 488 nm, λem = 500 nm) and red channel for J1 (λex = 800 nm, λem = 550 nm). Scale bar: 10 μm. (B) Quantitative analysis of the weakened fluorescence intensities of J1 (red line) and Syto 9 (green line). (C) Deoxyribonuclease (DNase) and ribonuclease (RNase) digest experiments of J1 (λex = 800 nm) using two-photon fluorescent images, scale bar: 20 μm. (C-i) Two-photon fluorescent images of J1 in HepG2 cells. (C-ii) Two-photon fluorescent images of J1 in HepG2 cells after dealing with DNase. (C-iii) Two-photon fluorescent images of J1 in HepG2 cells after dealing with RNase. (D) The histogram of the changes in two-photon fluorescence intensities of J1, J1 + DNase, and J1 + RNase. (E) Flow cytometry of the cellular uptake of J1 in the HepG2 cells. HepG2 cells were incubated with 20 μM of J1 in DMEM for 6 h at 37 °C.

investigated in thick kidney tissue in vitro. The rat kidney slice incubated with 50 μM J1 for 30 min exhibits the nucleoli labeling (white arrow in Figure S29B). The TPEF enhancement is also observed at the depth of ∼100 μm of rat kidney slice after incubation with J1 (Figure S31). It demonstrates that the TPEF of J1 is able to permeate into deep tissue and has the potential applications in monitoring the states and activities of deep organization.

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CONCLUSIONS

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ASSOCIATED CONTENT

In summary, a series of novel terpyridine derivatives with quaternary ammonium groups have been synthesized and characterized. The results indicate that the location of the nitrogen in terpyridine have an effect on their photophysical properties. Large Stokes shifts are verified in all J1−J3, and the two-photon absorption action cross sections range from 38 to 97 GM. In addition, two-photon confocal imaging experiments indicate that J1 could pass through both the plasma and nuclear membrane. Using molecular docking techniques and STED nanoscopy, J1 is shown to efficiently target rRNA in nucleoli through the electrostatic attractions of the quaternary ammonium groups as well as through the π−π interactions of the terpyridine groups. The nucleoli imaging can be observed at the depth of ∼100 μm in 3D multicellular spheroids and rat kidney slice through TPM. With enhanced water solubility, hypotoxicity, and good biocompatibility, J1 would be a potential candidate for selective rRNA fluorescent probe targeting mRNA and tRNA in the living cells.

Figure 8. STED micrographs of J1 staining nucleolus in different numbers with higher resolution. J1 displayed clear fibrillar center (asterisk, fc) and dense fibrillar component (arrow, dfc). Scale bar: 500 nm.

offer a fascinating tool to study nucleolar rRNA in dynamic manner in the living cells under super-resolution nanoscopy. Three-Dimensional Multicellular Spheroid and Tissue Imaging. To evaluate the imaging depth, J1 is applied for 2PEF imaging of 3D multicellular spheroid and rat kidney slices, and the changes in fluorescence intensity with scan depth are then determined in z-scan mode. The 3D multicellular spheroid is used as simulation model of in vitro 3D solid tumor for evaluating the imaging depth using 2PEF (Figure S29A). HepG2 multicellular spheroids incubated with 20 μM J1 are imaged under two-photon confocal microscopy. After treatment with J1, a significant fluorescent enhancement is observed at the depth of ∼100 μm of spheroids for two-photon excitation (Figure S30). In addition, the bioimaging of J1 is

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acsami.7b08068. 31430

DOI: 10.1021/acsami.7b08068 ACS Appl. Mater. Interfaces 2017, 9, 31424−31432

Research Article

ACS Applied Materials & Interfaces

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Detailed characterization and cell-related methods (PDF)

AUTHOR INFORMATION

Corresponding Authors

*E-mail: [email protected] (X.T.). *E-mail: [email protected] (J.W.). *E-mail: [email protected] (Y.T.). ORCID

Wei Du: 0000-0002-7959-6227 Hui Wang: 0000-0002-5253-1204 Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This work was supported by grants from the National Natural Science Foundation of China (51432001, 21602003, 51672002, 51372003, and 21501001), Anhui Provincial Natural Science Foundation of China (1708085MC68), and Anhui University Doctor Startup Fund (J01001962). The Higher Education Revitalization Plan Talent Project (2013). 2017 Returnees innovation and entrepreneurship key support program.

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