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Mitochondrial Peroxynitrite Mediates Anthracycline Induced Cardiotoxicity as Visualized by a Two-Photon Near-Infrared Fluorescent Probe Xilei Xie, Fuyan Tang, Guangzhao Liu, Yong Li, Xingxing Su, Xiaoyun Jiao, Xu Wang, and Bo Tang Anal. Chem., Just Accepted Manuscript • DOI: 10.1021/acs.analchem.8b03207 • Publication Date (Web): 10 Sep 2018 Downloaded from http://pubs.acs.org on September 10, 2018

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Analytical Chemistry

Mitochondrial Peroxynitrite Mediates Anthracycline Induced Cardiotoxicity as Visualized by a Two-Photon Near-Infrared Fluorescent Probe Xilei Xie,†,* Fuyan Tang,† Guangzhao Liu, Yong Li, Xingxing Su, Xiaoyun Jiao, Xu Wang,* and Bo Tang* College of Chemistry, Chemical Engineering and Materials Science, Key Laboratory of Molecular and Nano Probes, Ministry of Education, Collaborative Innovation Center of Functionalized Probes for Chemical Imaging in Universities of Shandong, Institute of Molecular and Nano Science, Shandong Normal University, Jinan 250014, P. R. China. E-mail: [email protected]; [email protected]; [email protected] ABSTRACT: Anthracyclines rank among the most efficacious anticancer medications. However, their clinical utility and oncologic efficacy are severely compromised by the cardiotoxicity risk facing the early diagnosis difficulty and the puzzled molecular mechanism. Herein, a two-photon excitable and near-infrared emissive fluorescent probe, TPNIR-FP, was fabricated and endowed with extraordinary specificity, sensitivity, and rapid response toward peroxynitrite (ONOO−), as well as mitochondriatargeting ability. With the aid of TPNIR-FP, we demonstrate that mitochondrial ONOO− is upregulated in the early stage and contributes to the onset and progression of anthracycline cardiotoxicity in cardiomyocyte and mouse models, therefore represents an early biomarker to predict the subclinical cardiotoxicity induced by drug challenge. Furthermore, TPNIR-FP is proved a robust imaging tool to provide critical insights into drug induced cardiotoxicity and other ONOO− related pathophysiological processes.

Anthracyclines (doxorubicin, epirubicin, daunorubicin, etc.) rank among the first-line anticancer drug categories, but their clinical application is extensively hampered by the cardiotoxicity risk. Their adverse effects range from subclinical myocardial dysfunction to serious cardiomyopathy and heart failure which may finally result in cardiac transplantation and even death. What’s more, these complications may manifest even decades after anthracycline administration.1−4 Current recommendation for cardiotoxicity monitoring is mainly based on the assessment of the left ventricular ejection function by echocardiography or multigated acquisition scan.5−7 However, this approach is not ideal because the impairment can be detected only after the dysfunction has already emerged. Therefore, identification of early biomarkers to predict the subclinical cardiotoxicity before the damage becomes uncontrollable and irreversible, is of significant benefit. On the other hand, the molecular pathogenesis of anthracycline cardiotoxicity still remains fairly controversial. The wide accepted hypothesis is the overgeneration of reactive oxygen species (ROS) by interfering with the electron transport chain in mitochondria.8,9 Besides, nitric oxide synthases are upregulated and nitric oxide (NO) is demonstrated a crucial molecule involved in the cardiotoxic pathophysiology.10,11 Interestingly, peroxynitrite (ONOO−) originates from the reaction of superoxide radical and NO, thus can simultaneously represent both the oxidative and nitrative stress levels. Moreover, as one of the most powerful endogenous toxins, ONOO− dysregulates the signal transduction by oxidizing and nitrating the biomolecules, induces mitochondria apoptosis and subsequent cell death, and

finally results in organ dysfunction.12 Therefore, mitochondrial ONOO− is supposed to play a pivotal role in the pathogenesis of anthracycline cardiotoxicity.13 However, there is no intuitive and visible evidence to support this point of view. Herein, we propose that ONOO− is overgenerated in mitochondria and initiates the cardiomyocyte apoptosis and myocardial dysfunction, and can serve as a potential biomarker to early predict the anthracycline cardiotoxicity. To prove this issue, the major challenge is how to accurately monitor the subtle concentration variations of mitochondrial ONOO− in cardiomyocytes and cardiac tissues due to its extremely short half-life (~10 ms), nanomolar homeostatic concentration,12 and other elusive nature in physiological environments. Fluorescent probe based assay provides an efficient methodology to real-time visualize molecular events in live cell and animal models by benefiting from its high resolution and noninvasive characteristic.14−30 In particular, two-photon fluorescent probes with near-infrared (NIR) emission are preferred for live tissue imaging due to their increased penetration depth and eliminated background fluorescence from some intrinsic biomolecules.31−33 Up to now a number of fluorescent probes for ONOO− have been reported,34−45 only few of them are endowed with satisfactory analytical parameters, capability for mitochondria targeting and live tissue imaging. To this end, we judiciously created a two-photon excitable and NIR emissive fluorescent probe (TPNIR-FP) for ONOO− detection and visualization. TPNIR-FP displayed not only satisfactory specificity, sensitivity, and rapid response toward ONOO−, but also mitochondria-targetable characteristic. These favorable properties enabled the direct visualization of

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mitochondrial ONOO− upregulation during anthracycline induced cardiotoxicity in both live cardiomyocytes and cardiac tissues. In particular, in combination with other bioanalytical methods, the direct correlation between elevated ONOO− levels and anthracycline cardiotoxicity has been well established.

EXPERIMENTAL SECTION Synthesis of TPNIR-FP. The probe was synthesized via a one-step procedure from TPNIR-NH2 outlined in Scheme S1. TPNIR-NH2 was prepared according to the previous literature.46 2-(4-Nitrophenyl)-2-oxoacetic acid (78 mg, 0.4 mmol) was dissolved in anhydrous dichloromethane (4 mL). After addition of oxalyl chloride (106 µL, 1.2 mmol) and DMF (2 drops), the mixture was refluxed at 40 °C for 1 h. The solvent was evaporated, and the residue was added into a mixture of TPNIR-NH2 (16 mg, 0.05 mmol) and triethylamine (28 µL, 0.2 mmol) in anhydrous dichloromethane (5 mL). The reaction mixture was stirred at room temperature for 1 h. The solvent was evaporated, and the residue was purified by column chromatography on silica gel with dichloromethane and methanol (200/1, V/V) to afford a solid (18 mg, 70% yield). 1H NMR (400 MHz, DMSO-d6): δ 11.42 (s, 1H), 8.69 (s, 1H), 8.40 (d, J = 8.9 Hz, 2H), 8.34 (d, J = 8.9 Hz, 2H), 8.25 (d, J = 9.4 Hz, 1H), 7.95 (s, 2H), 7.93 (s, 1H), 7.47 (dd, J = 9.4, 2.2 Hz, 1H), 7.30 (d, J = 1.9 Hz, 1H), 3.70 (q, J = 7.0 Hz, 4H), 3.08 (br s, 4H), 1.26 (t, J = 7.0 Hz, 6H). 13C NMR (100 MHz, DMSO-d6): δ 186.62, 161.81, 161.69, 158.23, 155.47, 150.52, 148.46, 143.31, 142.96, 137.45, 131.96, 131.79, 131.02, 126.98, 123.82, 122.31, 120.97, 119.58, 119.18, 118.69, 118.26, 95.65, 45.53, 26.54, 24.45. HRMS (ESI): calculated for C29H26N3O5+ (M+) 496.1867, found 496.1865. Cell Culture and Imaging. H9c2 cells were cultured in high glucose DMEM supplemented with 10% fetal bovine serum, 1% penicillin, and 1% streptomycin at 37 °C in a 5% CO2 ⁄95% air incubator MCO-5AC (SANYO, Tokyo, Japan). One day before imaging, the cells were detached and were replanted on glass-bottomed dishes. For monitoring of ONOO− in H9c2 cells, the cells were pretreated with 3morpholinosydonimine hydrochloride (SIN-1, 1.0 mM) for 30 min before being washed three times with PBS, then incubated with TPNIR-FP (5 µM) for 30 min before rinsed with PBS. For the scavenging assay, cells were treated with uric acid (100 µM) for 2 h in advance. Prior to the colocalization experiments, H9c2 cells were pretreated with SIN-1 (1.0 mM) for 30 min before being washed three times with PBS, then costained with the corresponding organelle marker (50 nM) and TPNIR-FP (5 µM) for 30 min and then rinsed for three times with PBS. For monitoring of ONOO− during anthracycline cardiotoxicity, H9c2 cells were pretreated with the corresponding concentration (0, 10, 20, and 20 µM) of doxorubicin (Dox) or epirubicin (Epi) for 6 h before being washed three times with PBS, then incubated with TPNIR-FP (5 µM) for 30 min before rinsed with PBS. For the scavenging assay, cells were treated with uric acid (100 µM) for 2 h in advance. For cell apoptosis staining, H9c2 cells were pretreated with the corresponding concentration (0, 10, 20, and 20 µM) of Dox or Epi for 6 h before being washed three times with PBS, then incubated with fluorescein-annexin V (5 µL, beyotime) and propidium iodide (10 µL, beyotime) for 30 min before rinsed with PBS. For the scavenging assay, cells were treated with uric acid (100 µM) for 2 h in advance. Cell imaging was performed on Leica TCS SP8 confocal laser

scanning microscope (one-photon) and Zeiss LSM 880 confocal microscope (two-photon). Mouse Models. All animal experiments were in strict accordance with the National Institutes of Health (NIH) Guide for the Care and Use of Laboratory Animals. Kunming mice were randomly divided into the control group, the lower dose group, the higher dose group, and the scavenging group. The mice were intraperitoneally injected with the corresponding dosage of drugs (0, 25, 50, and 50 mg/kg of Dox or Epi) and then injected with TPNIR-FP through the tail vein. For the scavenging group, the mice were pretreated with uric acid (100 mg/kg) intraperitoneally before drug administration and probe loading. All the mice were then anesthetized, and dissected to isolate the hearts, which were then fixed with 4% paraformaldehyde for 12 h. Then the samples were dehydrated with ethanol and embedded in paraffin before 6 µm sectioning of the left ventricles. Two-photon fluorescence images of ONOO− were acquired using a Zeiss LSM 880 confocal laser scanning microscope. As to immunohistochemistry staining assays, the sections were washed with PBS buffers for three times and blocked with normal goat serum for 1 h. And then, an anti-nitrotyrosine primary antibody was utilized to label nitrotyrosine overnight, followed by colored with rhodamine (TRITC)-labeled secondary antibody for 1 h. After that, the sections were imaged by excited at 543 nm with a Zeiss LSM 880 confocal laser scanning microscope and the emission was collected between 580−640 nm. HE staining was performed using hematoxylin and eosin under standard protocols.

RESULTS AND DISCUSSION Design, Synthesis, and Fluorescence Response of TPNIR-FP. The newly designed fluorescent probe (TPNIRFP) was fabricated by incorporating the α-ketoamide functionality developed by our lab44 onto an amino group containing fluorophore, TPNIR-NH2 (Scheme 1). In this design, TPNIR-NH2 was selected as the optimal fluorophore due to its large rigid conjugated system and lipophilic cation moiety which enables two-photon imaging, NIR emission, and mitochondria-targeting.46 We envisioned that the probe TPNIR-FP would accumulate into the mitochondria, and the α-ketoamide bridge would be rapidly cleaved by mitochondrial ONOO− to release the fluorophore TPNIR-NH2, which subsequently resulted in the fluorescence restoration (Scheme 1). TPNIR-FP was readily prepared via a one-step procedure outlined in Scheme S1, and characterized by HRMS, 1H NMR, and 13C NMR. Scheme 1. Design and Reaction Pathway of TPNIR-FP for ONOO− Imaging

We first compared the optical properties of TPNIR-FP and TPNIR-NH2 (Figure S1). They exhibited maximum absorption at 550 nm and 570 nm, respectively. When excited at 570 nm, TPNIR-NH2 featured a strong fluorescence emission at 630 nm, and its quantum yield (Φf) was determined to be 0.101. By contrast, TPNIR-FP was essentially nonfluorescent around 630 nm (Φf = 0.012) mainly due to the donor-excited photoinduced electron transfer (d-PeT) process from the


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Analytical Chemistry excited fluorophore to the electron-deficient nitro-substituted benzene moiety.47 The fluorescence response of TPNIR-FP in the presence of ONOO− was examined. After incubation with different concentrations of ONOO− (0−5 equiv), a dose-dependent fluorescence increment was observed (Figure 1a), and the intensity at 630 nm exhibited excellent linear correlation (R2 = 0.995) with the added ONOO− concentration (Figure 1b). The detection limit was determined to be 34 nM based on the formula of 3σ/k, which enabled real-time visualization of the traces of endogenous ONOO− in living systems.

Figure 1. (a) Fluorescence spectra of TPNIR-FP with ONOO− titration (0−5 equiv). (b) Linear correlation between fluorescence intensity of TPNIR-FP at 630 nm and ONOO− concentration. (c) The time course of fluorescence intensity of TPNIR-FP at 630 nm after adding 10 µM of ONOO−. (d) Fluorescence responses of TPNIR-FP toward 100 µM analytes unless otherwise denoted: (1) 10 µM ONOO−, (2) blank, (3) H2O2, (4) t-BuOOH, (5) NO, (6) 1 O2, (7) O2•−, (8) ClO−, (9) •OH, (10) 5 mM GSH, (11) Cys, (12) Hcy, (13) H2S, (14) NO3−, (15) NO2−, (16) SO42−, (17) SO32−, (18) CO32−, (19) PO43−, (20) Na+, (21) K+, (22) Ca2+, (23) Zn2+, (24) Cu2+, (25) Fe2+, (26) Fe3+, and (27) vitamin C. The final concentration of TPNIR-FP was 2 µM. All experiments were done in phosphate buffer (50 mM, pH 7.4) with 1% DMSO at 37 °C. λex/λem = 570/630 nm.

The above fluorescence variation was attributed to the ONOO− mediated cleavage of α-ketoamide bridge and concomitant release of TPNIR-NH2 fluorophore, which finally resulted in the restored fluorescence emission. To further confirm this reaction pathway, the reaction mixture of TPNIRFP and ONOO− was subjected to HPLC and HRMS analyses. As a result, the HPLC chromatogram of the reaction mixture displayed that a new peak with the same retention time as TPNIR-NH2 appeared (Figure S2). Furthermore, the mass spectrum of the reaction mixture showed that two new peaks emerged at m/z 319.1814 and 166.0166 (Figure S3), which agreed well with the released fluorophore TPNIR-NH2 and the byproduct p-nitrobenzoic acid, respectively. These results clearly validated the ONOO− triggered decomposition pathway of TPNIR-FP described in Scheme 1. The time-dependent fluorescence variation of TPNIR-FP in the presence of ONOO− was recorded. After addition of 10 µM of ONOO−, the fluorescence intensity of TPNIR-FP (2 µM) at 630 nm rapidly increased and finally reached a plateau within 10 seconds (Figure 1c), indicating that TPNIR-FP holds

rapid response and is capable of efficiently capturing ONOO− despite its elusive properties. Additionally, the effect of pH value was evaluated. TPNIR-FP held higher fluorescence OFF/ON ratio toward ONOO− within an extremely narrow pH range of 7.0−8.0 (Figure S4), which overlapped well with the mitochondrial pH in live cells. ONOO− (pKa = 6.8)12 presents higher nucleophilicity under neutral and alkaline conditions. On the other hand, the fluorescence of TPNIR-NH2 was gradually quenched when the pH value was higher than 8.0 (Figure S5).48,49 Therefore, TPNIR-FP exhibits the weakly alkaline pH-dependent response, which is well corresponding to its potential mitochondria-targeting ability discussed in the later section. Subsequently, the specificity of TPNIR-FP was examined by screening its fluorescence response to a panel of biologically relevant species (Figure 1d and Figure S6). As expected, TPNIR-FP achieved significant fluorescence enhancement only in the presence of ONOO−. Negligible fluorescence variations were induced by other potential interfering substances, including reactive oxygen and nitrogen species (H2O2, t-BuOOH, NO, 1O2, O2•−, ClO−, and •OH), reactive sulfur species (GSH, Cys, Hcy, and H2S), anions (NO3−, NO2−, SO42−, SO32−, CO32−, and PO43−), cations (Na+, K+, Ca2+, Zn2+, Cu2+, Fe2+, and Fe3+), and vitamin C. H2O2 possesses similar peroxide moiety and reactivity with that of ONOO−, thus often cross-reacts with ONOO− probes. Therefore, we particularly investigated the fluorescence response of TPNIR-FP toward high amounts of H2O2 (Figure S7). Encouragingly, no signal increment was observed in presence of 500 µM of H2O2, and only 1.5-fold fluorescence increase was detected when the concentration of H2O2 was as high as 1 mM. In contrast, 10 µM of ONOO− treatment resulted in ca. 8-fold signal increment. Collectively, TPNIRFP demonstrates sufficient specificity for ONOO− visualization in complex biological surroundings. Moreover, TPNIR-NH2 was incubated with a variety of reactive oxygen and sulfur species to evaluate its stability. As a result, the fluorescence remained constant in most cases, although some fluorescence decrement (no more than 10%) was observed (Figure S8). Therefore, TPNIR-NH2 was relatively stable under physiological conditions during the fluorescence imaging process. Visualizing ONOO− in Mitochondria. The cytotoxicity of TPNIR-FP was assessed in H9c2 cardiomyocytes by the standard MTT assay (Figure S9). The IC50 value of TPNIR-FP was calculated to be 141 µM, indicating its minimal cytotoxicity. Taken together, the aforementioned characteristics in terms of high sensitivity, sufficient specificity, rapid response, and negligible cytotoxicity, render TPNIR-FP particularly favorable for tracking ONOO− fluctuation in live cells. As shown in Figure 2, the probeloaded cells displayed faint fluorescence upon excitation at 800 nm using a two-photon laser. By contrast, the cells emitted striking red fluorescence after pretreated with 3morpholinosydonimine hydrochloride (SIN-1), a wellestablished ONOO− generator. Meanwhile, the red fluorescence was evidently suppressed by a widely used ONOO− scavenger uric acid. These observations demonstrate that TPNIR-FP is competent to visualize ONOO− levels in live cells.



Figure 2. Two-photon fluorescence images of ONOO− in H9c2 cardiomyocytes. Cells were treated with (a) PBS, (b) 1.0 mM SIN-1, or (c) 1.0 mM SIN-1 and then 100 µM uric acid, followed by incubation with 5 µM TPNIR-FP. (d) Relative fluorescence intensities of (a)−(c). Emissions were collected from 600 to 640 nm and excited at 800 nm. Scale bar: 10 µm.

It is known that lipophilic cations like TPNIR-FP tend to accumulate into the mitochondria, thus colocalization experiment was performed to confirm the subcellular distribution of TPNIR-FP (Figure 3). The red fluorescence of TPNIR-FP almost completely overlapped with the green signal from Mito-Tracker Green, a commercial mitochondrial marker. The intensity profiles of linear regions of interest (ROIs) across a selected cell of the two channels altered in close synchrony and the Pearson’s colocalization coefficient was determined as high as 0.93. To further confirm the above result, additional colocalization experiments were performed by costaining cells with TPNIR-FP and Lyso-Tracker Green (lysosome marker), ER-Tracker Green (endoplasmic reticulum marker), or Golgi-Tracker Green (Golgi apparatus marker), respectively. Nearly no colocalization was observed in all the three cases (Figure S10). Collectively, TPNIR-FP is capable of accumulating and imaging in mitochondria where ONOO− dominantly originates and functions.

Figure 4. Two-photon fluorescence images of ONOO− upregulation in H9c2 cardiomyocytes induced by Dox and Epi. Top line: cells were treated with (a) PBS, (b) Dox (10 µM, 6 h), (c) Dox (20 µM, 6 h), or (d) uric acid (100 µM, 2 h) plus Dox (20 µM, 6 h), followed by incubation with TPNIR-FP (5 µM, 30 min); (e) Relative fluorescence intensities of (a)−(d). Bottom line: cells were treated with (f) PBS, (g) Epi (10 µM, 6 h), (h) Epi (20 µM, 6 h), or (i) uric acid (100 µM, 2 h) plus Epi (20 µM, 6 h), followed by incubation with TPNIR-FP (5 µM, 30 min); (j) Relative fluorescence intensities of (f)−(i). Emissions were collected from 600 to 640 nm and excited at 800 nm. Scale bar: 20 µm.

Figure 3. H9c2 cells were costained with (a) Mito-Tracker Green (50 nM) and (b) TPNIR-FP (5 µM). (c) Colocalized image. (d) Intensity profiles of ROIs across a selected cell of the two dyes. Cells were pretreated with 1.0 mM SIN-1 for 30 min before labeled with dyes. The excitation wavelengths were (a) 488 nm and (b) 561 nm, respectively, and the emissions were collected at (a) 500−540 nm and (b) 600−640 nm. Scale bar: 20 µm.

Visualizing ONOO− Mediated Cardiomyocyte Apoptosis During Anthracycline Cardiotoxicity. Subsequently, TPNIR-FP was applied to monitor the mitochondrial ONOO− fluctuation during anthracycline induced cardiotoxicity using the cultured H9c2 cardiomyocyte model. Doxorubicin (Dox) and epirubicin (Epi), two representative anthracyclines, were selected to induce the myocardial cytotoxicity. Compared to the cells in control group, the drug treated cells exhibited dose-dependent fluorescence increase, and the increased fluorescence was effectively suppressed by the ONOO− scavenger uric acid (Figure 4). The solid data evidently demonstrated that cardiomyocytes experienced the mitochondrial ONOO− burst after administration of Dox or Epi, and the upregulated ONOO− concentrations were positively dependent on the drug dosages.

Figure 5. Confocal images of fluorescein-annexin V and PI stained H9c2 cell apoptosis induced by (a) Dox and (b) Epi. H9c2 cardiomyocytes were treated with the same regimen as used in Figure 4. First column: cells were pretreated with PBS as control. Second column: cells were pretreated with Dox or Epi (10 µM, 6 h). Third column: cells were pretreated with Dox or Epi (20 µM, 6 h). Fourth column: cells were pretreated with uric acid (100 µM, 2 h), and then Dox or Epi (20 µM, 6 h). Images of fluorescein (green) were obtained by collecting the emissions at 510−540 nm upon excitation at 488 nm. Images of PI (red) were obtained by collecting the emissions at 610−640 nm upon excitation at 543 nm. Scale bar: 20 µm.


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Analytical Chemistry Generally, ONOO− impairs the mitochondrial energy metabolism, activates the apoptotic program, and finally induces mitochondria and cell apoptosis. Thereby, we anticipated that mitochondrial ONOO− would be a proapoptotic factor to initiate the cardiomyocyte apoptosis after anthracycline administration. Therefore, fluoresceinannexin V/propidium iodide (PI) double staining was performed to image the H9c2 cardiomyocytes, which were treated with the same regimen as used in Figure 4. The fluorescein labeled annexin V binds externalized phosphatidylserine on the outer membrane in the early stage of apoptosis, while PI can only enter cells and stain nucleus in late apoptosis. When treated with lower dosage (10 µM) of Dox or Epi, green fluorescence signal from fluoresceinannexin V was observed but red fluorescence of PI was absent (Figure 5, 2nd column), confirming the early stage of cardiomyocyte apoptosis. After treatment with higher dosage (20 µM) of Dox or Epi, the cells showed enhanced green fluorescence and the red signal appeared (Figure 5, 3rd column), indicating that apoptotic level increased and partial cardiomyocytes entered late apoptosis. We noticed that with the increasing dosage of drug treatment, mitochondrial ONOO− was gradually upregulated (Figure 4), and concomitantly cell apoptosis level increased (Figure 5). To unveil the correlation between elevated ONOO− level and cardiomyocyte apoptosis, the cells were pretreated with uric acid to selectively remove the overgenerated ONOO− (Figure 4d and 4i). As a result, both the green and red fluorescence signals were apparently suppressed (Figure 5, 4th column), suggesting that removal of ONOO− by a scavenger efficiently attenuated the cardiomyocyte apoptosis. Therefore, we conclude that ONOO− plays a crucial role in initiating and promoting the cardiomyocyte apoptosis, and represents a potential biomarker for early diagnosis of drug induced cardiotoxicity. Visualizing ONOO− Burst and Related Cardiotoxicity in Mouse Models. Subsequently, ONOO− fluctuation and related cardiotoxicity were visualized in the mouse model to further validate the aforementioned conclusions. Kunming mice were randomly divided into the control group, the lower dose group, the higher dose group, and the scavenging group. The mice were intraperitoneally injected with the corresponding dosage of drugs (0, 25, 50, and 50 mg/kg of Dox or Epi) and then injected with TPNIR-FP through the tail vein. For the scavenging group, the mice were pretreated with uric acid (100 mg/kg) intraperitoneally before drug administration and probe loading. All the mice were then anesthetized, and heart left ventricles were isolated and cut into slices for two-photon fluorescence imaging and other bioanalysis. Compared with the control group, the drug administration groups presented striking red fluorescence in a dose-dependent manner, and the fluorescence was obviously blocked by uric acid treatment (Figure 6a). These fluorescence changes in myocardial tissues were highly consistent with those in cultured H9c2 cardiomyocytes, further validated the endogenous ONOO− upregulation during Dox and Epi induced cardiotoxicity. The nitration of tyrosine residues indicates not only that ONOO− has been generated nearby, but also the irreversible impairment on the involved proteins.50 Therefore, nitrotyrosine level was detected by immunohistochemical staining. Compared with the control and scavenging groups, drug administration induced widespread and unambiguous occurrence of nitrotyrosine in myocardial tissues (Figure 6b),

suggesting the potential cardiac damage caused by ONOO−. Moreover, haematoxylin and eosin (HE) staining was performed to identify the morphological changes after drug administration (Figure 6c). The control group showed a regular myofibrillar alignment, while the drug administration group exhibited serious disorganization of myofibrillar arrays and cytoplasmic vacuolization. Uric acid pretreatment removed the elevated ONOO− and markedly counteracted the pathological changes, indicating that ONOO− definitely contributed to the drug induced cardiac damage. Taken together, these outcomes validate the overgeneration and pivotal contribution of ONOO− during anthracycline induced cardiotoxicity in animal models, in support of the conclusion achieved in cultured H9c2 cardiomyocyte models.

Figure 6. Visualization of ONOO− mediated cardiotoxicity in mouse models induced by anthracycline administration. First column: mice were pretreated with PBS as control. Second column: mice were injected with Dox or Epi (25 mg/kg). Third column: mice were injected with Dox or Epi (50 mg/kg). Fourth column: mice were injected with uric acid (100 mg/kg), and then Dox or Epi (50 mg/kg). After preadministration for 1 h, all the mice were injected with TPNIR-FP (50 µM, 100 µL) through the tail vein. After 0.5 h, the mice were anesthetized, and heart left ventricles were isolated and cut into slices for (a) two-photon fluorescence imaging of ONOO−, (b) immunohistochemical assay of nitrotyrosine, and (c) HE staining. Two-photon fluorescence images of ONOO− were obtained by collecting the emissions at 600−640 nm upon excitation at 800 nm. Scale bar: 50 µm.

CONCLUSION In summary, we have proposed a unique fluorescent probe TPNIR-FP for ONOO− detection and visualization. TPNIR-FP



achieves an exceptionally rapid, selective, and sensitive fluorescence turn-on response toward ONOO−, and is characterized by two-photon imaging, NIR emission, and mitochondrial localization, which enables tracking of the subtle concentration variations of mitochondrial ONOO− in cardiomyocytes and cardiac tissues. With the aid of TPNIRFP, we have demonstrated that mitochondrial ONOO− burst emerges in the early stage after anthracycline administration and plays a crucial role in initiating and promoting the cardiomyocyte apoptosis, thus represents an early biomarker to predict the subclinical cardiotoxicity induced by drug challenge. These findings provide novel critical insights into diagnosis and management of drug induced cardiotoxicity. Meanwhile, TPNIR-FP is proved a robust imaging tool to explore ONOO− biofunction in a broad range of pathophysiological processes.

ASSOCIATED CONTENT Supporting Information The Supporting Information is available free of charge on the ACS Publications website. Experimental details, supplementary data, and characterization of compounds (PDF)

AUTHOR INFORMATION Corresponding Author *E-mail: [email protected]. Fax: +86-531-86180017. *E-mail: [email protected]. *E-mail: [email protected].

Author Contributions †

These authors contributed equally to this work.

Notes The authors declare no competing financial interest.

ACKNOWLEDGMENT This work was supported by the National Natural Science Foundation of China (21535004, 91753111, 21505088, 21775093, and 21390411) and the Key Research and Development Program of Shandong Province (2018YFJH0502).

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