Radical Pair-Driven Luminescence of Quantum Dots for Specific

Feb 19, 2016 - Merouani , D. R.; Abdelmalek , F.; Ghezzar , M. R.; Semmoud , A.; Addou , A.; Brisset , J. L. Ind. Eng. Chem. Res. 2013, 52, 1471– 14...
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Radical Pair-Driven Luminescence of Quantum Dots for Specific Detection of Peroxynitrite in Living Cells Wenjuan Zhou, Yuqing Cao, Dandan Sui, and Chao Lu* State Key Laboratory of Chemical Resource Engineering, Beijing University of Chemical Technology, Beijing 100029, China S Supporting Information *

ABSTRACT: There is currently great interest in developing chemiluminescence (CL) probes that can selectively detect peroxynitrite (ONOO−) in living cells. In comparison with other reactive oxygen species (ROS), ONOO− can spontaneously decompose into a series of radicals. Notably, the interaction of quantum dots (QDs) with oxidizing/reducing ROS radicals can generate a strong CL emission by electron-transfer annihilation. Herein, we report a novel CL probe that affords the ability to distinguish ONOO− from other ROS in living cells. ONOO− can activate luminescence of QDs in the absence of excitation source, effectively avoiding background noise and scattering of light from biological matrixes produced by in situ excitation; however, there is no response to other ROS including 1O2, H2O2, •OH, O2•−, and ClO−. The outstanding selectivity of the present CL probe leads us to detect the exogenous release of ONOO− from 3-morpholinosydnonimine (SIN−1) in living cells. These results suggest that this present probe-based CL provides a promising platform for highly selective and sensitive detection of ONOO− in biological systems. ntracellular reactive nitrogen species (peroxynitrite, ONOO−) from the diffusion-controlled reaction between nitric oxide and superoxide radicals can easily react with different biomolecules through direct oxidation or decomposition into highly reactive secondary radicals (e.g., hydroxyl radical and carbonate radicals), leading to potentially serious damage in living cells.1−3 Accordingly, ONOO− has been implicated as a key pathophysiological intermediate in a growing list of diseases, such as cardiovascular, neurodegenerative, and inflammatory disorders.4−6 In addition, ONOO− can act as a cytotoxic effector against invading pathogens, displaying protective activities in vivo.7 Recently, ONOO− also plays key roles in the redox regulation of signaling pathways.8,9 Therefore, accurate quantitation of ONOO− in living cells is vital not only for fundamental studies to understand its role in pathophysiological processes but also for early diagnosis of potential corresponding diseases. However, most of the beneficial properties of ONOO− remain controversial or poorly characterized owing to the lack of reliable methods for monitoring ONOO− in living cells. Chemiluminescence (CL) is the emission of luminescence as a result of chemical reaction without the need of background luminescence. Owing to the absence of autofluorescence interference by photoexcitation, CL represents a promising strategy for detecting biologically important reactive oxygen species (ROS).10−12 However, CL-based detection of ONOO− is limited to the cross-interference from other ROS species.13 Therefore, the utilization of CL for high specificity toward ONOO− over other ROS in living cells still remains a great challenge. Quantum dots (QDs) are highly luminescence materials in a wide emission range from UV to NIR. QD−ROS interactions play a pivotal role in a wide variety of QD CL processes.14−16 In general, QDs are highly susceptible to redox reactivity with oxidizing/reducing radicals to generate oxidized and reduced

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QDs by injecting a hole/electron to QDs, i.e., the injection of an electron from reducing radicals (e.g., O2•−) into the LUMO of QDs to produce a reducing QDs (R•−), and the injection of a hole from oxidizing radicals (e.g., •OH) into the HOMO of QDs to produce an oxidizing QDs (R•+). The generated R•+/R•− QDs can generate a strong CL emission by electron-transfer annihilation.17−19 Such an intriguing interaction between QDs and radicals could promote the development of new strategies for the CL detection of ROS. Notably, multifarious sensing systems using QDs as CL labels are available for ions or small-molecules at present.20,21 However, the responsiveness of QDs to specific ROS remains relatively less studied.22 The reasons are partially due to difficulties in the coexistence of instable oxidizing and reducing ROS. Therefore, it is hoped that new approaches will gain more attention to generate oxidizing and reducing radicals to meet the high demand for develop novel CL approaches suitable for the specific detection of ROS. ONOO− can decompose into oxidizing and reducing radical pair.23,24 In principle, the interaction between QDs and the radical pair from ONOO− could produce the CL emissions by electron-transfer annihilation. In this work, the investigation of the energy levels of the thioglycolic acid (TGA)-capped CdTe QDs using cyclic voltammetry demonstrated that the oxidizing radical •OH from ONOOH is capable of injecting a hole into the valence band (VB) of the CdTe QDs to produce oxidized QDs (QDs•+). Subsequently, it occurs the electron-transfer annihilation between QDs•+ and O2•− from ONOO− to form the excited QDs, which could emit light when they returned to the ground Received: October 11, 2015 Accepted: February 9, 2016

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DOI: 10.1021/acs.analchem.5b03827 Anal. Chem. XXXX, XXX, XXX−XXX

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by Solarbio (Beijing, China). HeLa cells were provided by the American Type Culture Collection (ATCC, Rockville, MD). Dulbecco’s modified Eagle’s medium (DMEM) was purchased from Hyclone (Thermo Fisher scientific, USA) supplemented with 10% fetal bovine serum (FBS, Hyclone) and 1% penicillin/ streptomycin (Hyclone). ONOO− was synthesized in a quenchedflow reactor. Apparatus. The fluorescence spectra were carried out using a Hitachi F-7000 fluorescence spectrophotometer (Tokyo, Japan) with the excitation wavelength of 365 nm. The excitation slit and the emission slit were both maintained at 5.0 nm with a scanning rate of 1200 nm/min. The CL detection was conducted on an Ultra−Weak biophysics chemiluminescence (BPCL) luminescence analyzer (Institute of Biophysics, Chinese Academy of Science, Beijing, China). The CL spectrum of the CL system was measured with high-energy cutoff filters from 400 to 640 nm between the CL cell and the photomultiplier tube (PMT). The UV−vis absorption spectra were obtained on a Shimadzu UV-3600 spectrophotometer (Tokyo, Japan). High resolution transmission electron microscope (HRTEM) image was obtained with a TecnaiG220 TEM (FEI Co., Netherlands) at an accelerating voltage of 200 kV. Cyclic voltammetry (CV) curves of the CdTe QDs were performed on a CHI660E electrochemical workstation (CH Instruments, USA). Electron spin resonance (ESR) spectra were obtained on an ESR spectrometer (JEOL, JES-FA200 spectrometer, Tokyo). The fluorescence images of cells were taken using a DMI 3000B Digital Microscope (Germany Leica Co., Ltd.) with an objective lens (10×, 20×, 40×). For MTT assays, constant temperature microporous fast oscillator (QB-9006, China) and Multimode Plate Readers (PerkinElmer Enspire, USA) were used. Cell Culture and Cytotoxicity Assay. HeLa cells were cultured in DMEM supplemented with 10% heat-inactivated FBS 100 units/mL of penicillin and 100 mg/mL of streptomycin at 5% CO2, 37 °C. Cells were seeded in a 6-well plate at a density of 105 cells/well and incubated in 2.0 mL of DMEM/well for 24 h. Prior to experiments, cells were washed twice with Tris− HCl (pH = 7.4) to remove the remnant growth medium and then incubated in a DMEM medium (2.0 mL) containing various concentrations of CdTe QDs (0.04−5.00 μM) for 6 h. The cell viability was determined by MTT assay according to the manufacturer’s instructions. Briefly, HeLa cells were seeded in a 96-well plate at a density of 104 cells/well and incubated in 200 μL of DMEM/well for 24 h. The culture media were removed, and the cells were incubated with different concentrations of CdTe QDs for 6 h. Then 10 μL of sterile-filtered MTT stock solution and 90 μL fresh medium was added to each well. After 5 h of incubation, the unreacted dye was removed by aspiration, and 110 μL of DMSO was added to each well. The absorbance at 490 nm was measured using a Multimode Plate Readers after all the formazan crystals were dissolved. The cell viability (%) relative to control cells cultured in media without CdTe QDs was calculated from [A]test/[A]control × 100%, where [A]test and [A]control are the absorbance values of the wells (with QDs) and control ones (without QDs), respectively. Fluorescence Imaging of CdTe QD-Loaded Cells. The cellular uptake of the CdTe QDs was determined under a fluorescence microscope. For the fluorescence imaging, the QD-labeled HeLa cells were washed three times with pH 7.4 Tris−HCl and then examined by a DMI 3000B Digital Microscopes with an objective lens (10×, 20×, 40×), excitation at 365 nm.

Scheme 1. Schematic Illustration for CL Emission of the CdTe QDs Triggered by Oxidizing and Reducing Radical Pair from ONOO− Decomposition

states (Scheme 1). However, other ROS including 1O2, H2O2, • OH, O2•−, and ClO− cannot initiate the CL of CdTe QDs. To take advantage of this exciting finding, we also demonstrated the potential use of this approach for monitoring exogenously produced ONOO− from 3-morpholinosydnonimine (SIN−1) in living cells. With high selectivity, signal stability, good sensitivity, and fast response time to ONOO−, the proposed method has great potential in physiological and pathological applications.

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EXPERIMENTAL SECTION Chemicals and Materials. All reagents used were analyticalreagent grade without further purification, and all deionized water from a Millipore water purification system (18.2 MU cm, Milli Q, Millipore, Barnstead, CA, USA) was used throughout the experiments. NaOH, NaClO, FeSO4, KO2, sodium azide (NaN3), and ascorbic acid (AA) were purchased from Beijing Chemical Reagent Company. A mixed working solution of 0.05 M H2O2 and 0.03 M HCl was freshly prepared by volumetric dilution of 36% (v/v) HCl (Beijing Chemical Reagent Company) and commercial 30% (v/v) H2O2 (Beijing Chemical Reagent Company). Stock solution of 0.1 M NaNO2 was prepared by dissolving 0.69 g of NaNO2 (Tianjin Chemical Reagent Company) in 100 mL of deionized water. Working solutions of NaNO2 were freshly prepared by diluting the NaNO2 stock solution with deionized water. ClO− was prepared by diluting commercial NaClO solution in deionized water. 1O2 was generated from the online reaction of NaClO solution (100 μM) with H2O2 (100 μM). •OH was generated from the online reaction of FeSO4 (100 μM) and H2O2 (100 μM) through Fenton reaction. O2•− was prepared by dissolving 1.1 mg of KO2 into 100 mL of dimethyl sulfoxide (DMSO). Working solutions of 3-morpholinosydnonimine (SIN−1, Toronto Research Chemicals Inc.) were prepared by dissolving SIN−1 into NaOH solution (0.01 M) at 37 °C. The commercial CdTe QDs capped with mercapto-carboxylate ligands were purchased from BeiDaJuBang Chem. Co. Ltd. (Beijing, China). The concentration and size of the CdTe QDs were calculated according to the method reported in the literature.25,26 In detail, the particle size (2.8 nm) of the QDs can be determined from the first absorption maximum in virtue of the following empirical formula: D = (9.8127 × 10−7)λ3 − (1.7147 × 10−3)λ2 + (1.0064)λ − 194.84, where the D was the diameter size of the CdTe QDs, and the λ was the wavelength of the first absorption maximum peak of the CdTe QDs. The concentration of the QDs (C) can be calculated upon the absorbance of the QD solution (A) according to Beer’s Law: A = εbC, where ε = 10043 D2.12 and b = 1 cm. 3-(4,5-Dimethylthiazol-2-yl)-2,5diphenyltetrazolium bromide (MTT) and DMSO were supplied B

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Figure 1. (A) Cyclic voltammograms recorded for (a) 50 mM Tris−HCl (pH = 7.4) and (b) 0.74 μM CdTe QDs in 50 mM Tris−HCl (pH = 7.4). The scan rate was 100 mV/s; working electrode: glassy carbon electrode; reference electrode: Ag/AgCl electrode, counter electrode: Pt electrode. (B) The energies for 1Sh and 1Se quantum-confined orbitals of the CdTe QDs and the standard redox potentials of different ROS. (C) Normalized CL intensity of CdTe QDs to various ROS. Inset: spectra overlap of fluorescence (FL) spectrum of the CdTe QDs and CL spectrum of the CdTe QD−ONOO− system. (D) Normalized FL spectra of the CdTe QDs in the presence of different ROS (100 μM). PMT voltage: 400 V; λex = 365 nm; EX slit: 5 nm; EM slit: 5 nm.

CL Measurements. The CL signals were examined with a static CL system. Fresh solutions of ONOOH were prepared by reacting NaNO2 with acidified H2O2, and then it mixed with NaOH (0.2 M) to produce ONOO−. The concentration of the standard ONOO− solution was measured by the absorbance at 302 nm according to Beer’s Law: A = εbC (ε = 1670 M−1 cm−1).27 100 μL of the as-prepared ONOO− solution was injected into 200 μL of CdTe QDs (in 50 mM Tris−HCl, pH = 7.4) by a microliter syringe from the upper injection port. The CL signals were monitored by a PMT (−1000 V) adjacent to the CL quartz cell. Detection Procedure of Exogenous ONOO− in Living Cells. HeLa cells were seeded in a 6-well plate under the conditions as described in the cell culture. After treatment with 0.16 μM CdTe QDs for 4 h, the QD-loaded cells were washed three times with Tris−HCl buffer (50 mM, pH = 7.4) to remove any extracellular QDs. Then the cells were suspended with pancreatin treatment and resuspended in the same Tris−HCl buffer solution. For living cell assays, SIN−1 was utilized to generate ONOO−. SIN−1 solution was incubated at 37 °C for 50 min and then injected into the suspension of the QD-loaded cells. The reaction was performed in the quartz cell, and the CL signals were monitored by a PMT (−1000 V) adjacent to the CL quartz cell.

were depicted in Figure S1A. Moreover, the average size of the CdTe QDs was observed to be about 2.8 nm with good crystal structure from the HRTEM image (Figure S1B). The oxidation and reduction characteristics of the CdTe QDs were determined by cyclic voltammetry measurements. A reversible oxidation wave from 1.113 V with a peak at 1.194 V (marked as A2) was displayed from CdTe QDs (Figure 1A) during anodic potential sweep. In this anodic process, the holes can be injected from an electrode into the CdTe QDs to form the radical cation of CdTe QDs (QDs•+).28 On the other hand, a reversible reduction wave from −0.895 V with a peak at −1.002 V (marked as C1) was observed during the cathodic potential sweep. This cathodic peak was coupled with an anodic peak appearing at −0.624 V (marked as A1), meaning the formation of the stable radical anion of CdTe QDs (QDs•−).29 In addition, the potential difference of 2.11 eV between C1 and A2 was in accordance with the optical band gap (Eg) of 2.10 eV, which can be calculated using the equation Eg = 1240/λg, where λg (nm) was the value of CdTe QDs absorption edges (Figure S1A).30 From the cyclic voltammogram of CdTe QDs, the HOMO energy level of CdTe QDs was calculated to be −5.823 eV, according to the equation EHOME = −((Eonset(ox))+4.71) eV, where the onset oxidation potential (Eonset(ox)) was determined to be 1.113 V.31 Combined with the HOMO−LUMO band gap (Eg) of 2.11 eV, the LUMO energy level of CdTe QDs was calculated to be −3.713 eV.32 Specific CL Response of CdTe QDs to ONOO−. ONOOH could rapidly decompose under physiological conditions. However, ONOOH can be deprotonated into the stable ONOO− in

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RESULTS AND DISCUSSION HOMO−LUMO Band Gap of the CdTe QDs. The absorption and fluorescence properties of the commercial CdTe QDs C

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Figure 2. (A) Effects of scavengers (1.0 mM) for various ROS on the CdTe QD−ONOO− system. PMT voltage: −1000 V. (B) ESR signals of DMPO−•OH and DMPO−O2•− adducts in the ONOO− and the CdTe QD−ONOO− systems. The final concentration of DMPO is 0.06 M. Experimental procedure: 20 μL of 0.3 M DMPO + 32 μL of Tris−HCl/32 μL of 4 μM CdTe (in 0.05 M Tris−HCl buffer solution) + 20 μL of 0.01 M NaNO2 + 20 μL of 1 mM NaOH + 10 μL of 0.02 M HCl−0.05 M H2O2.

alkaline solutions. The homolysis of ONOOH/ONOO− would produce the •OH and O2•− radicals simultaneously.33,34 The proposed energy level diagram of the CdTe QDs and the standard redox potentials of different ROS in the ONOO− system were shown in Figure 1B. The •OH radical can inject holes into the 1Sh quantum-confined orbital of the CdTe QDs with a standard redox potential of 2.3 V.35 In addition, the ONOO− and ONOOH have a comparably potent oxidant with a redox potential of 1.4 and 2.1 V,36 which is capable of injecting the holes into the CdTe QDs. On the other hand, the standard redox potential of the reducing O2•− radical is −0.33 V,35 allowing an electron donation to the hole injected CdTe QDs. It occurs with the electron-transfer annihilation between QDs•+ and O2•− from ONOO− to form the excited QDs,37 which could emit light when they returned to the ground states. With a static injection method depicted in Figure S2, the CL signals of the CdTe QDs were investigated in the presence of various ROS including 1O2, H2O2, •OH, O2•−, ClO−, and ONOO−. As shown in Figures 1C and S3, there were no obvious CL emissions when the CdTe QDs solution was mixed with 1O2, H2O2, •OH, and O2•− (each 100 μM), respectively. Note that the relative weak CL emission can be generated in the simultaneous presence of •OH and •O2− in vitro (Figure S3). However, the lifetime of •OH is short (10−9 s), and •O2− can occur spontaneously in a dismutation reaction in a physiological environment.38,39 Therefore, it is impossible that a large amount of •OH and •O2− can coexist in a living system. In contrast, ONOO− has a longer lifetime (1 s) and can simultaneously decompose to produce •OH and •O2−, producing a strong CL emission. Therefore, a trace amount of •OH and •O2− could not interfere with the detection of ONOO−. In conclusion, the proposed system exhibited a highly selective CL response toward ONOO−. On the other hand, we investigated the fluorescence responses of the CdTe QDs toward such the ROS. As shown in Figure 1D, the fluorescence signals of the CdTe QDs showed an indiscriminate quenching in the presence of different ROS (Figure 1D). These results further demonstrated the superiority of the CdTe QD-induced CL emissions over the fluorescence of the CdTe QDs for the selective detection of ONOO−. Reactive Intermediates in the QD−ONOO− CL System. In order to prove the CL reactive intermediates for the selective response of the CdTe QDs toward ONOO−, the CL spectrum of the ONOO− system in the presence of the CdTe QDs was measured with high-energy cutoff filters from 400 to 640 nm.

The maximum emission wavelength of the proposed CL system was about 540 nm, which was in accordance with the fluorescence (FL) spectrum of the CdTe QDs (inset of Figure 1C), demonstrating that the CL emission was generated from the excited-state QDs. This QD-based CL emission may be attributed to the energy transfer between ONOOH* and QDs34 or electron−hole recombination of QDs.40 In order to identify the CL process of QDs, the reactive intermediates produced in the CdTe QD−ONOO− CL reaction were measured by the scavengers of various ROS.41 As shown in Figure 2A, 1.0 mM NaN3 (a scavenger for 1O2) had little effect on the CL signal of the CdTe QD−ONOO− system. However, the CL emission of the system was almost completely quenched in the presence of 1.0 mM NBT (a scavenger for O2•−), 1.0 mM AA (a scavenger for ROS), and 1.0 mM thiourea (a scavenger for •OH). These results showed that the CL emissions in the system were mainly originated from O2•− and •OH radicals, and the contribution of the ONOOH* energy transformation to the strong CL was less. Moreover, the existence of •OH and O2•− radicals was further confirmed using ESR spectroscopy. As a specific target molecule of •OH and O2•− radicals, DMPO was used to identify the production of •OH and O2•− radicals during the CL reaction of ONOO− with CdTe QDs in 0.05 M Tris−HCl buffer.42 There were significant ESR signals of DMPO−•OH and DMPO−O2•− adducts in both the ONOO− and the CdTe QD−ONOO− systems (Figure 2B), indicating the production of •OH and O2•− radicals in the proposed CL system. In combination with the scavenger tests and the energy level analysis, these results could verify the electron−hole recombination CL mechanism. Analytical Performances of the CdTe QD−ONOO− System. Under the experimental condition of 0.16 μM QDs in 0.05 M Tris−HCl buffer (pH = 7.4), we investigated the relationship between the CL intensity and ONOO− concentration. UV−vis absorption measurements were used to measure the concentration of the ONOO− produced by the online reaction of acidified H2O2 with NaNO2, followed by adding a NaOH solution to stabilize the ONOO−. The UV−vis absorption spectra for the as-prepared ONOO− sample were shown in Figure S4. According to Lambert−Beer and Kubelk−Munk theories, the concentration of ONOO− from 10 mM NaNO2 was calculated to be 460 μM from the absorbance at 302 nm (ε = 1670 M−1 cm−1).27 The CL intensity was found to be linear with the concentration of ONOO− in the range from 0.46 to 46 μM (Figure 3A), and the correlation coefficient was 0.9989. The detection limit for ONOO− (S/N = 3) was 0.1 μM. D

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Figure 3. (A) CL intensity of the CdTe QD−ONOO− system at different concentrations of ONOO−. Inset: the calibration curves for ONOO− in the range of 0.46−46.0 μM. (B) Time-dependent CL intensity of the SIN−1 induced ONOO− system in the presence of CdTe QDs (0.16 μM). SIN−1 (5 mM) was dissolved in 0.01 M NaOH solution and incubated at 37 °C for the generation of ONOO−. PMT voltage: −1000 V.

The relative standard deviation for five repeated measurements for 0.46 μM ONOO− was 2.0%. As a generator of ONOO−, SIN−1 is the most widely used to study the biological effects of ONOO− in physiological environments.2 Herein, the released ONOO− from 5.0 mM SIN−1 was monitored using the proposed CL approach. Figure 3B showed the CL signals over a 2-h incubation period. It can be seen that the concentration of ONOO− generated by SIN−1 increased with incubation time and reached a maximum of 42.1 μM after 50 min of incubation, indicating that SIN−1 had decomposed completely within 50 min. These results corresponded closely with those examined in the other reports.43−45 However, the actual concentration of ONOO− was slightly lower than the expected ideal level (50 μM), which could be attributed to the possible reactivity with ambient CO2.45 Detection of ONOO− in Living Cells. It is well-known that the inherently toxic elements and the thiol ligands would render the CdTe QDs toxic to both cells and living systems.46,47 Therefore, the potential toxicity of the CdTe QDs is a key point for practical applications. In the present work, we used the commercial CdTe QDs with good biocompatibility for cell endocytosis and ONOO− detection. The cytotoxicity of the commercial CdTe QDs was evaluated according to MTT assays on HeLa cancer cells. Figure S5 showed that the cytotoxicity of the CdTe QDs to HeLa cells increased with increasing extracellular QD concentration. Furthermore, various concentrations of CdTe QDs were used to investigate their cellular uptake. The fluorescence microscope images of HeLa cells with CdTe QDs were shown in Figure S6. It can be seen that the cellular uptake was dependent on the concentration of the CdTe QDs with an optimal concentration of 0.16 μM. Next, HeLa cells were first incubated with 0.16 μM of QDs for 4 h (Figure 4A and S7). After the uptake, the cells were washed with Tris−HCl for three times to remove any extracellular QDs. For the living cell assays, the cells were suspended with pancreatin treatment and resuspended in Tris−HCl buffer (50 mM, pH = 7.4). Under the living cell system, we investigated the relationship between the CL intensity and ONOO − concentration in the range of 2.3−460 μM (Figure 4B). Linear regression analysis revealed good linearity between the CL intensity and the log concentration of ONOO− (R2 = 0.998). The relative standard deviation for three repeated measurements of 2.3 μM ONOO− was 2.5%. The effects of the typical interferences including some ions and reductants present in cells were investigated (Table S1). The results demonstrated that

Figure 4. (A) Fluorescence images of HeLa cells after treatment with 0.16 μM CdTe QDs for 4 h. λex = 488 nm; scale bars, 40 μm (top: bright field image; down: fluorescence image); (B) CL signals from the CdTe QDs in HeLa cells by adding various concentrations of ONOO−. Inset: the calibration curves for ONOO− in the range of 2.3−460 μM; (C) CL signals from HeLa cells after addition of ONOO− generated by SIN−1 (0, 0.5, 5.0 mM, respectively). PMT voltage: −1000 V.

these coexistent substances had no influence on the determination of 10 μM ONOO−. Finally, the ONOO− generator SIN−1 was used to support the potential of the proposed method in the biosensing system. The CL measurement was performed by adding a working solution of 0.5 mM SIN−1 into the suspension of QD-loaded HeLa cells. An obvious CL emission generated after injection of SIN−1 (Figure 4C), indicating a fast diffusion of ONOO− into cells. The concentration of ONOO− generated from 0.5 mM SIN−1 was determined to be 4.07 μM. In addition, the CL intensity increased when the SIN−1 concentration was increased to 5.0 mM, and the ONOO− concentration was determined to be 42.8 μM, which corresponded closely with the content of the extracellular examination (42.1 μM). These results demonstrated that the changes in the ONOO− level in living cells could be monitored through the proposed CL method, suggesting the potential of the CdTe QDs as a CL probe for further biochemical research in living systems. E

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CONCLUSIONS To close, a versatile CL method has been developed for highly selective and sensitive detection for ONOO− exogenously produced from SIN−1 in living cells by utilizing the formation of QDs•+ via the interaction between QDs and oxidizing ROS radicals from the decompostion of ONOO−. The generated QDs•+ and reducing radicals can generate a strong CL emission by electron-transfer annihilation without external excitation, allowing the removal of interfering autofluorescence and scattering of light from biological matrixes. This probe features an excellent selectivity for ONOO− over biologically competing ROS, including 1O2, H2O2, •OH, O2•−, and ClO−. This work is the first CL probe available for the detection of ONOO− in living cells. It could be anticipated that although further improvements are certainly required, this proposed strategy would provide a powerful tool to meet the challenges in distinguishing ONOO− from other ROS in biological systems. Current efforts are directed toward tracking endogenous ONOO− production, as well as determining potential signaling events associated with ONOO− in pathophysiological processes.

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.analchem.5b03827. UV−vis absorption and fluorescence spectra of the commercial CdTe QDs; schematic diagram of a static CL setup; CL response of CdTe QDs to various ROS; UV−vis absorption spectra of the ONOO− produced from the reaction of H2O2/HCl with NaNO2 in different concentrations; cells viability values estimated by MTT assays; fluorescence images of HeLa cells after treatment with various concentrations of CdTe QDs for 4 h; fluorescence images of HeLa cells after treatment with 0.16 μM CdTe QDs for 4 h; tolerance limit of various coexistent substances on the determination of 10 μM ONOO− (PDF)

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AUTHOR INFORMATION

Corresponding Author

*Phone/Fax: 86 10 64411957. E-mail: [email protected]. Author Contributions

W.J.Z., Y.Q.C., and D.D.S. contributed equally to this work. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This work was supported by National Basic Research Program of China (973 Program, 2014CB932103), the National Natural Science Foundation of China (21375006 and 21575010), and the Innovation and Promotion Project of Beijing University of Chemical Technology.

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DOI: 10.1021/acs.analchem.5b03827 Anal. Chem. XXXX, XXX, XXX−XXX

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DOI: 10.1021/acs.analchem.5b03827 Anal. Chem. XXXX, XXX, XXX−XXX