A fluorescent/photoacoustic probe for imaging of hydroxyl radical and high-throughput screening of natural products to attenuate acute kidney injury

doi:10.21203/rs.3.rs-2923115/v1

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A fluorescent/photoacoustic probe for imaging of hydroxyl radical and high-throughput screening of natural products to attenuate acute kidney injury

https://doi.org/10.21203/rs.3.rs-2923115/v1

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The hydroxyl radical (•OH) has been shown to play a crucial role in the occurrence and progression of acute kidney injury (AKI). Therefore, the development of a robust •OH detection tool holds great promise for the early diagnosis of AKI and high-throughput screening (HTS) of inhibitors to attenuate AKI. In this work, we report the design and synthesis of an activatable fluorescent/photoacoustic (PA) probe (CDIA) for sensitive and selective imaging of •OH in AKI. CDIA has near-infrared (NIR) fluorescence/PA channels and fast activation kinetics, enabling the detection of the onset of •OH in an AKI model. The positive detection time of 12 hours using this probe is superior to the 48-hour detection time for typical clinical assays, such as blood urea nitrogen (BUN) and serum creatinine (sCr) detection. Furthermore, a method has been established using CDIA for HTS of natural •OH inhibitors from herbal medicines. Puerarin has been screened out by activating the Sirt1/Nrf2/Keap1 signaling pathway to protect renal cells in AKI. Overall, this work provides a versatile and dual-mode tool for illuminating the •OH-related pathological process in AKI and for screening additional compounds to prevent and treat AKI.

Biological sciences/Chemical biology/Small molecules

Health sciences/Nephrology/Kidney diseases/Acute kidney injury

Acute kidney injury

Hydroxyl radical

Molecular probe

High-throughput screening

Natural products

Acute kidney injury (AKI) is a serious complication that affects hospitalized patients at a high risk of developing progressive chronic kidney disease or end-stage renal disease. It is responsible for over 1.5 million deaths globally each year,^[1] making the timely diagnosis and treatment of AKI critical for preventing and managing kidney diseases. In vitro diagnostic methods have been employed to monitor renal function and prevent AKI. However, the current diagnostic approach for kidney disease in clinical settings relies on static analysis of blood urea nitrogen (BUN) and serum creatinine (sCr),^[2] which only increase after a 50% decrease in glomerular filtration rate (GFR).^[3] Non-invasive imaging strategies, such as single-photon emission computed tomography (SPECT) and magnetic resonance imaging (MRI), are limited by in vitro diagnostic methods based on static analysis, which are difficult for dynamic monitoring of renal functional impairment.^[4] Molecular optical imaging offers a noninvasive way to occurrence and development of different kinds of diseases in organisms in real time with high spatiotemporal resolution,^[5–7] making it a promising tool for in situ detection of molecular processes associated with renal disease. Many fluorescent probes have been reported for AKI imaging, but the shallow penetration depth of fluorescence limits their further application for in vivo imaging.^[8] In contrast, photoacoustic (PA) imaging, a hybrid imaging modality that detects sound rather than photons, has minimal attenuation due to negligible sound scattering in living organisms. This property offers a deeper penetration depth (7–10 cm) and higher spatial resolution for biological imaging.^[9] Activatable PA molecular probes have already been explored for deep-tissue imaging,^{[10, 11]} and the deeper imaging in the kidney avoids biological background signal fluctuations, enabling earlier and more precise detection of renal dysfunction.

Therapeutic options for preventing AKI involve the use of small molecular agents like N-acetyl cysteine (NAC), which help reduce oxidative damage.^[12] In recent studies, nanoreagents with reactive oxygen species (ROS)-modulating capabilities have been explored as potential therapeutic strategies for ROS-associated AKI.^{[13, 14]} DNA origami nanomaterials, for example, have demonstrated a strong ability to scavenge ROS, preserve kidney structures, and improve AKI condition.^[15] However, the sensitivity of nanomaterials to the biological environment, including subtle changes in shape, morphology, or physical properties, as well as the lack of practical guidelines, pose significant challenges in this field of research.^[16] Complementary and alternative medicine could potentially play a unique role in treating AKI. In China, traditional Chinese medicine (TCM) has been used for thousands of years to prevent diseases. Natural products, such as epigallocatechin-3-gallate,^[17] saikosaponin-D^[18] and resveratrol^[19], have been employed to treat drug-induced nephrotoxicity, which supports the exploration of effective natural products from medicinal plants for AKI treatment. Nevertheless, developing natural products into effective therapies for AKI presents challenges due to the complicated ingredients and multiple-target nature of natural products, and testing their efficacy and safety in disease models can be a time-consuming and costly process. Therefore, direct visualization of AKI in vivo and rapid screening of efficient natural products for AKI treatment are highly desirable but challenging goals.

AKI is a complex condition triggered by ischemic and inflammatory components that ultimately result in the loss of renal epithelial cells and tubular cell dysfunction. Among various pathological factors associated with AKI, oxidative stress (OS) has been identified as the most crucial.^[20] OS is caused by ROS such as superoxide anion, hydrogen peroxide, and hydroxyl radical (•OH). While ROS play a crucial role in maintaining physiological functions, their hyper-regulated or unregulated accumulation can cause oxidative damage to biomolecules, perturb cell membrane, macromolecular and organelle functions. Severe pathophysiological conditions can enhance ROS production, leading to vascular dysfunction, inflammation, and tubular cytotoxicity, commonly observed in the pathogenesis of AKI.^[21] Animal studies have shown that the occurrence of AKI increases oxidative damage in the body while decreasing tissue antioxidant status.^{[22, 23]} Although the exact mechanism of ROS generation in AKI remains unknown, ROS has been identified as an early hallmark of AKI,^{[24, 25]} especially •OH with the highest reduction potential (2.31 V) compared with other ROS.^[26] Therefore, it can be chosen as a target for intervention of AKI.

Herein, a •OH activatable fluorescent/photoacoustic probe (termed CDIA) has been designed and synthesized for imaging of AKI in murine model. Previous research has shown that the reaction of •OH with a bulky phenol like diiodophenol favors an electron transfer reaction, resulting in the formation of a phenoxyl radical that leads to the decomposition of aromatic compounds.^[27] Additionally, taking cues from the efficient reaction between •OH and aspirin,^[28] a turn-on molecular probe can be rationally designed and synthesized. Thus, the probe CDIA is designed to have two components: 1) the signaling moiety NIR hemi-cyanine dye^[29] and 2) a target-responsive substrate iodosalicylic acid, which is recognized and cleaved by •OH (Fig. 1). The probe is initially non-fluorescent and non-photoacoustic in an aqueous solution, because the hydroxyl group is in a "caged" state with diminished electron-donating ability. Upon exposure to •OH, the iodosalicylic acid component of the molecule undergoes a specific reaction to release the free dye (CyOH) and transition into an "uncaged" state. As a result of the enhanced electron-donating ability from the oxygen atom of the free dye, the product exhibits turned-on NIR fluorescence and PA signals (Fig. 1). Thus, the probe can be activated by •OH to produce its NIR fluorescence and PA signal. With its exceptional sensitivity and selectivity to •OH, CDIA has proven effective in visualizing the endogenous •OH production in living cells and mice subjected to various stimuli, without being affected by other ROS. Moreover, CDIA can be applied for the high-throughput screening (HTS) of natural •OH scavengers from herbal medicines in live cells, which could reveal further mechanisms involved in the chemical regulation of natural products to attenuate AKI.

Synthesis and Characterization of CDIA

The construction of CDIA involved three main steps: i) Synthesis of the symmetric cyanine dye 1 (CyCl), ii) synthesis of the hemi-cyanine precursor 2 (CyOH), and iii) synthesis of the probe CDIA by conjugating CyOH and 3,5-diiodosalicyclic acid. The hemi-cyanine precursor CyOH was via a retro-knoevenagel reaction using the symmetric cyanine dye CyCl and resorcinol,^[30] followed by its reaction with a 3,5-diiodosalicyclic acid to afford the hemi-cyanine CDIA. According to the integration ratio of proton in the ¹H NMR spectrum (Supplementary Fig. 1–4) and mass spectrum (Supplementary Fig. 5), the dual-mode probe CDIA had been successfully conjugated with a molecular weight of 674.55.

Spectral Response of CDIA to •OH In Vitro

In order to confirm the response of the probe to •OH, optical spectra analyses were utilized to track the changes of the probe both qualitatively and quantitatively. Initially, CDIA was non-fluorescent and had an NIR absorption peak at 650 nm (Fig. 2a). Upon exposure to •OH at 37 ℃, the probe exhibited a significant decrease in the absorption peak at 650 nm and a new peak at 690 nm emerged (Fig. 2b), corresponding to the free dye CyOH. Additionally, CDIA demonstrated an increase in fluorescence intensity at 720 nm (Fig. 2c and Supplementary Fig. 6a) and a rise in fluorescent quantum yield to 0.27 following its reaction with •OH (Table S1). This was attributed to the effective reaction between •OH and iodosalicylic acid subunit, leading to the formation of a CyOH moiety with a strong electron-donating phenolate group of the fluorophore and thus strong fluorescence. Good linearity between the fluorescence intensities and the concentrations of •OH was observed with a limit of detection (LOD) of 5.87 nM (Fig. 2d), indicating its ultrasensitivity towards •OH compared with other previously reported works (Table S2). The probe exhibited remarkable sensitivity and fast response to Fenton reagent, completing the oxidation within 3 min (Fig. 2e). In addition, CDIA showed negligible NIR fluorescence responses to other interfering analytes, including other ROS and metal ions (Fig. 2f, Supplementary Fig. 7,8), indicating its high specificity of NIR activation channel towards •OH. The probe functioned well across the physiological pH range of 6.0–9.0 (Fig. 2g and Supplementary Fig. 9) and demonstrated excellent stability (Fig. 2h,i), suggesting its potential for detecting •OH in living organisms. The PA properties of CDIA were also studied, and the PA signal of the probe at 690 nm significantly increased in the presence of •OH, consistent with the absorption changes upon treatment with •OH (Supplementary Fig. 10). All the results confirmed that CDIA had excellent photoactivation properties to detect •OH in vitro. Furthermore, high-performance liquid chromatography (HPLC) and mass spectrum further demonstrated that the probe (HPLC retention time, CDIA: T_R = 6.949 min) was totally hydrolyzed and converted into free hemi-cyanine dye CyOH (T_R = 10.596 min) after incubation with •OH (Supplementary Fig. 11).

We proceeded to test whether the probe was capable of enabling ratiometric imaging of •OH in vitro using RGB image analysis with a smartphone.^[31] The fluorescence color of CDIA changed noticeably from blue to green upon reacting with various concentrations of •OH (Fig. 2j). True-color photos captured with a smartphone were separated into red (R), green (G), and blue (B) channels and digitized through an image-processing algorithm. The ratio of green and blue channel intensities (G/B ratio) could then be used to measure the concentration of •OH. Therefore, it could readily detect concentrations of •OH as low as 2 µM with the naked eye from the G/B ratio, indicating that CDIA has a wide range of applications for identifying •OH.

Numerous chemical compounds, including biologically significant ones such as DMSO,^[32] thiourea,^[33] glutathione (GSH),^[34] NAC,^[35] 2,2,6,6-tetramethyl-1-piperidinyloxy (TEMPO)^[36] and thyroid hormones,^[37] like triiodothyronine (T₃) and thyroxine (T₄), that have the capability to scavenge •OH. Unfortunately, a reliable and sturdy technique for measuring their reactivity is not yet accessible, which obstructs the understanding of many essential mechanisms in both biological and chemical systems.^[38] The use of CDIA probe makes it possible to calibrate and quantify the scavenging efficiency of known scavengers, thereby facilitating deeper exploration of cellular antioxidant capability. By detecting the response of CDIA to Fenton reagents in the presence of different scavenger concentrations, we were able to determine their scavenging efficiency. Supplementary Fig. 12 showed that ranking of scavenging effectiveness was as follows: T₄ > T₃ > TEMPO > GSH ≈ thiourea > NAC ≫ DMSO. The reason for the excellent effectiveness of T₄ and T₃ was their ability to directly react with •OH, which was similar to CDIA. Therefore, we utilized the extremely sensitive CDIA probe to validate and compare the antioxidant properties of several typical •OH scavengers in vitro.

NIR Fluorescent Response of CDIA to Dynamic Changes of •OH in Living Cells

After confirming that the probe has good cytocompatibility (Supplementary Fig. 13), CDIA was applied to detect •OH in cultured cells. HK-2 cells (an immortalized proximal tubule epithelial cell line from normal adult human kidney) were subjected to nephrotoxic drug cisplatin (cis-diammineplatinum dichloride, CDDP).^[39] The half maximal inhibitory concentration (IC₅₀) was defined as the concentration which was necessary to induce the apoptosis percentage to 50% (Supplementary Fig. 14). HK-2 cells were treated with CDDP at the corresponding IC₅₀ for 24 h, respectively. In control groups, HK-2 cells were preloaded with 10 mM thiourea (an acknowledged scavenger of •OH) for 2 h before treated CDDP. All groups were then incubated with CDIA (10 µM) for NIR fluorescence imaging. After incubation for 30 min, the NIR fluorescence of CDDP-treated HK-2 cells were remarkably enhanced by 22.97-fold (Fig. 3a–c), confirming the feasibility of CDIA for drug-induced cell injury imaging. Co-staining experiments demonstrated that CDIA, following •OH activation, was predominantly localized in the mitochondria region (Fig. 3d and Supplementary Fig. 15–17). Conversely, a significant reduction in fluorescence intensity was observed in HK-2 cells pre-treated with thiourea, further verifying that the fluorescence amplification originated from CDIA after activation by overexpressed •OH in HK-2 cells. Meantime, monitoring of intracellular •OH over time using CDIA showed no evidence of photoactivating or photobleaching (Supplementary Fig. 18,19, and Video S1).

In order to detect endogenous •OH in a broad range of contexts, we investigated the usage of CDIA in various model cells. Specifically, we evaluated its performance in RAW264.7 mouse macrophages and HeLa cells by treating them with PMA, a protein kinase C (PKC) activator that induces endogenous •OH.^[38] The cells were co-incubated with PMA (at concentrations of 200 or 500 ng mL^− 1) for 4 h, followed by incubation with 10 µM CDIA at 37°C for 30 min. The results showed a significant increase in NIR fluorescence signal compared to the control group (Supplementary Fig. 20,21). Additionally, we tested the effectiveness of CDIA in detecting OS in HepG2 cells caused by acetaminophen (APAP), which was a component of many cold medications that could cause liver injury and OS when taken in excess.^{[40, 41]} The results showed a dose-dependent increase in NIR fluorescence signal with different concentrations of APAP (Supplementary Fig. 22). These findings suggested that CDIA was a highly effective tool for intracellular •OH imaging.

The PA signals and spectra of CDIA were examined after it was incubated with cells. Supplementary Fig. 23 showed that the PA signal at 690 nm of HK-2 cell pellets treated with CDDP was higher than that of HK-2 cell pellets treated with both CDDP along with thiourea. The PA signals of RAW264.7, HeLa, and HepG2 cell pellets were consistent with the results obtained from confocal imaging (Supplementary Fig. 24–26). These findings not only demonstrated that CDIA could be selectively activated by •OH produced by cells, but also indicated its potential for use in PA imaging in living cells.

Low molecular weight chitosan (LMWC),^[42] a copolymer derived from chitin and composed of D-glucosamine and N-acetylglucosamine, was selected as the nanocarrier for renal targeting owing to its biocompatibility and biodegradability.^[43] LMWC has the ability to bind to the megalin receptor,^{[44, 45]} which is predominantly present on renal tubular epithelial cells. CDIA was encapsulated in LMWC to create the nanoprobe CDIA@LMWC NP. Theoretically, CDIA@LMWC NP remained stable under physiological conditions and could be selectively internalized into the lysosome through megalin receptor-mediated endocytosis, after which CDIA was released due to the acidic pH environment. Utilizing an ionic cross-linking technique (Supplementary Fig. 27),^[46] the CDIA@LMWC nanoparticles were synthesized, and exhibited a uniformly dispersed, spherical morphology measuring 50 nm on average, as observed through transmission electron microscopy (TEM) (Fig. 2k). The average hydrodynamic diameter of 58.3 ± 1.9 nm, as observed by dynamic light scattering (DLS) analysis, was attributed to the presence of hydration layers (Supplementary Fig. 28a). The targeting efficacy of the LMWC-based nanocarrier was evaluated using an FITC-labeled LMWC nanoparticle, which demonstrated good linearity in fluorescence intensity and successful conjugation of FITC to LMWC NP. The zeta potentials of LMWC NP, FITC-LMWC NP, and CDIA@LMWC NP were + 37.4, + 36.8, and + 32.9 mV, respectively, indicating that these colloidal particles could exhibit favorable stability and transportability during glomerular filtration. To assess renal cell targeting, an FITC-labeled LMWC nanoparticle (FITC-LMWC NP) was synthesized. The successful conjugation of FITC to LMWC NP was validated by a notable fluorescence peak observed at 518 nm in the fluorescence spectra (Supplementary Fig. 29) and a strong correlation between the concentration of the sample and the fluorescence intensity at 518 nm within the range of 0 to 10 µg/mL. The amount of FITC that had been conjugated to the FITC-LMWC NP was measured to be 63.84 ± 0.22 µg/mg using a standard curve method (Supplementary Fig. 30). The zeta potentials of LMWC NP, FITC-LMWC NP, and CDIA@LMWC NP were measured to be + 37.4, + 36.8, and + 32.9 mV, respectively (Supplementary Fig. 28 and Table S3), indicating good stability and transportation in glomerular filtration (Supplementary Fig. 31). These above findings suggested that LMWC could serve as a suitable nanocarrier for renal targeting.^[47]

To determine the encapsulation efficiency (EE) and loading content (LC) of the nanoprobe, the ultraviolet-visible-near infrared (UV-Vis-NIR) standard curves of CDIA were used (Fig. 2a). The EE and LC of CDIA were then measured to be 83.62 ± 0.17% and 10.57 ± 0.04%, respectively (Table S4). In vitro release behaviors of the nanoprobe CDIA@LMWC NP was investigated using the dialysis method at different pH levels.^[48] The nanoprobe remained stable over a 24-hour incubation period at pH 7.4 and 6.8 (Fig. 2l), while its release rate was significantly accelerated in an acidic environment of pH 5.0 (Fig. 2m). After incubation for 4 h, the accumulative release rate of CDIA reached 72.96 ± 0.53%, indicating effective delivery and release of the nanoprobe into the acidic lysosomes of renal cells. It is worth noting that the introduction of LMWC did not have a noticeable effect on the reaction of CDIA and •OH (Supplementary Fig. 32).

With the good biocompatibility (Supplementary Fig. 33), we further optimized the incubation time and concentration of FITC-LMWC NP by using confocal laser scanning microscopy. The results demonstrated that fluorescence signals of FITC increased gradually with improving incubation time, and almost no extracellular fluorescence was observed (Supplementary Fig. 34). Thus, a concentration of 20 µg/mL of FITC-LMWC NP and CDIA@LMWC NP with an incubation time of 2 h were chosen for subsequent experiments. In order to investigate the cell specificity of cellular uptake, HK-2 cells, AML-12 cells, H9c2 cells, HepG2 cells, HUVEC cells, and MCF-7 cells were incubated with FITC-LMWC NP. The selective accumulation of FITC-LMWC NP in HK-2 cells was shown by the increased green fluorescence observed through confocal fluorescence imaging. In contrast, minimal fluorescence was observed in the other cell lines (Fig. 3e and Supplementary Fig. 35). To investigate the subcellular distribution of the nanocarrier in cells, FITC-LMWC NP was incubated with HK-2 cells and co-stained with Lyso Tracker Red, a lysosome probe, and Hoechst 33342, a nucleus dye. By confirming the localization of FITC-LMWC NP in the lysosomal compartment (Fig. 3f and Supplementary Fig. 36) and ruling out mitochondrial regions (Supplementary Fig. 37), the yellow fluorescence resulting from the overlap of green FITC and red LysoTracker signals provided evidence for the targeting capability of nanocarrier to HK-2 cells. Further investigation was carried out to understand the mechanism underlying the increased cellular uptake of FITC-LMWC NP in HK-2 cells. Studies using ethylene diamine tetraacetic acid (EDTA), free LMWC, or glucosamine as megalin receptor inhibitors significantly reduced the uptake of FITC-LMWC NP (Fig. 3g and Supplementary Fig. 38),^[49] demonstrating that the LMWC nanocarrier was proposed to have been taken up by HK-2 cells via an endocytosis pathway that was mediated by the megalin receptor.

The production of •OH induced by nephrotoxic drugs in HK-2 cells was quantified using the commercial ROS probe dichlorodihydrofluorescein diacetate (H₂DCFDA). HK-2 cells were treated with CDDP, aristolochic acid I (AAI), and citrinin (CTN) at the respective IC₅₀ concentrations for 24 h (Supplementary Fig. 14).^{[50, 51]} The fluorescence intensity in HK-2 cells experienced a gradual increase with increasing concentration of the nephrotoxic drugs (Supplementary Fig. 39). Confocal fluorescence imaging and flow cytometric assays were then performed to evaluate the feasibility of using CDIA@LMWC NPs for imaging nephrotoxic drug-induced injury in HK-2 cells. The cells were incubated with CDDP for 24 h, and the fluorescent changes of 20 µg/mL CDIA@FITC-LMWC NP-loaded HK-2 cells were monitored in real time using confocal fluorescence imaging (Video S2). Initially, bright green fluorescence was observed in the lysosomes (Fig. 3h), indicating good lysosomal location and cell viability. Subsequently, an increment in the red fluorescence of CDIA within HK-2 cells was observed in the confocal fluorescence images, which signified an escalation in •OH production throughout CDDP treatment. By collecting the fluorescence signals in the chondriosome (Fig. 3i), the fluorescence changes could be observed (Fig. 3j), demonstrating the excellent application of CDIA@FITC-LMWC NP for discriminative imaging of CDDP-induced HK-2 cell injury. Meanwhile, CDIA@LMWC NP was also utilized for monitoring •OH generation during the process of nephrotoxic drugs AAI and CTN stimulation. The NIR fluorescence of AAI- and CTN-treated HK-2 cells were remarkably enhanced by 22.63- and 24.57-fold, respectively (Supplementary Fig. 40,41). However, a remarkable decrease in fluorescence intensity was observed in HK-2 cells pretreated with thiourea, which confirming the feasibility of CDIA@LMWC NP for drug-induced cell injury imaging.

Likewise, PA images and spectra of CDIA@LMWC NP after incubation with HK-2 cells were also studied. On the premise that LMWC would not interfere with the reaction between the probe CDIA and •OH (Supplementary Fig. 42), PA signal of HK-2 cell pellets after treatment with AAI or CTN at 690 nm was higher than the PA signal of HK-2 cell pellets treated with AAI or CTN along with thiourea (Supplementary Fig. 43,44). These above results displayed that the dual-mode nanoprobe CDIA@LMWC NP could be employed to visualize the dynamic changes of •OH on nephrotoxic drugs-induced renal injury.

High-Throughput Screening of Natural Products

With the impressive results obtained using CDIA@LMWC NP for live-cell imaging, we proceeded to investigate its potential as a screening tool for identifying antioxidants that could reduce •OH formation. To this end, we established a straightforward and effective HTS platform using CDIA@LMWC NP. Specifically, HK-2 cells were pre-treated with various natural products with potential antioxidative activity at 50 µM for 12 h before administering 75 µM of CDDP for 24 h. Subsequently, images and quantitative analysis of endogenous •OH levels were conducted after the cells treated with 20 µg/mL CDIA@LMWC NP (Fig. 4a).

By utilizing this approach, the rapid and quantitative detection of endogenous •OH generation was ensured (Fig. 4b). Our findings demonstrated that many natural products selected were able to significantly reduce fluorescence signals, indicating a down-regulation of •OH generation. Of particular note, puerarin (Pue), a key active ingredient found in the herbal medicine Kudzu root,^[52] was shown to be highly effective in reducing fluorescence signals compared to the untreated control. This phenomenon suggested that this flavonoid compound could have potential as an agent for controlling the accumulation of nephrotoxic drug-generated •OH in live cells. The results demonstrated that CDIA@LMWC NP could serve as a valuable tool for screening natural compounds that regulate endogenous •OH variations in a simple yet powerful way. Most kits for the activity assay and screening of biological molecules work on the basis of optical density (OD) values at specific wavelengths for the designed chromogenic reagents. Here, to achieve more portability and efficiency, we used CDIA@LMWC NP with high sensitivity to detect •OH content and establish a HTS system to rapidly discover natural products to attenuate AKI (Fig. 4c). In this work, 50 natural compounds with a variety of chemical skeletons (e.g. coumarins, sesquiterpenes, monoterpenoids, diterpenoids, triterpenoids, flavones, steroids, alkaloids, polyphenols, and quinones) from a natural compound library have been evaluated for their antioxidant effects on AKI (Table S5). Compared with the control group, weak absorption intensity ratio at 690/650 nm (A₆₉₀/A₆₅₀) of CDIA was observed for the wells corresponding to compounds J9 (chlorogenic acid), K10 (quercetin, Que) and K11 (puerarin, Pue) (Fig. 4d), indicating potential •OH scavenging effect. In comparison with the control group, the relative A₆₉₀/A₆₅₀ of each well were interpreted intuitively as shown in Fig. 4e. Because the •OH scavenging effect of chlorogenic acid and Que had been clearly demonstrated by some previous works,^{[53, 54]} compound K11 (Pue) was selected as a potential therapeutic agent to attenuate AKI.

In Vivo Duplex Imaging of AKI

CDIA had almost identical fluorescence in the PBS and urine (Fig. 5a and Supplementary Fig. 45), indicating the excellent stability of CDIA in mice. Moreover, the absence of acute or chronic toxicity of CDIA and CDIA@LMWC NP was confirmed in mice (Supplementary Fig. 46), along with negligible hemolysis toxicity to red blood cells (Supplementary Fig. 47), indicating the potential of CDIA@LMWC NP for in vivo applications, specifically for monitoring renal diseases. The capacity of CDIA@LMWC NP for duplex imaging of AKI was assessed in a mouse model utilizing CDDP, a known nephrotoxic cancer treatment drug, as the model drug.^[55] A total of five groups, including a control group receiving saline and four treatment groups receiving the same total CDDP dosage but with different dosing frequencies, were established by random assignment of mice in this experiment. (Fig. 5b).^[56] CDIA@LMWC NP was injected intravenously (i.v.) after the CDDP dose, and in vivo imaging of the mice was conducted within the NIR window at various time intervals. Simultaneously, a transurethral catheter was used to collect urine samples from the mice, which were subsequently imaged while being exposed to 650 nm irradiation. Mice administrated with a high initial dose of CDDP exhibited signs of kidney damage after 12 h (Fig. 5c,d, Group I and Group II). There was a gradual increase in the NIR fluorescence signals and PA intensities in the kidneys over time, which suggested renal damage progressed from mild to severe levels (Fig. 5e,f and Supplementary Fig. 48). Group IV that received a low initial dose of CDDP treatment experienced delayed onset of kidney dysfunction, while repeated doses of CDDP also resulted in kidney damage. The urine of the mice displayed comparable fluorescence signal variations to those observed in the in vivo NIR fluorescence (Fig. 5g,k). Whereas the NIR fluorescence and PA amplitude in the kidney signaled the status of kidney injury that had built up over time, the urine signal indicated the the degree of transient renal damage. The pathological analysis of kidney slices further validated the above findings (Supplementary Fig. 49). Notably, the use of a dual-mode imaging approach for monitoring kidney dysfunction allowed for the identification of CDDP-induced kidney injury 36 h prior to clinical diagnosis based on BUN and sCr levels (Supplementary Fig. 50), providing additional confidence for AKI diagnosis. The LMWC-targeted kidney monitoring method facilitated the timely detection of renal disease and furnished valuable information on the evolution of kidney injury during treatment, which could potentially contribute to treatment optimization.

In order to further explore the effect of remission on renal injury, the control group of mice received treatment with saline or nephroprotective antioxidant Que or NAC (10 mg kg^–1 day^–1, intraperitoneal (i.p.) injection) 3 d prior to CDDP administration. At different post-treatment timepoints, whole-body longitudinal NIR fluorescence imaging (Fig. 5i) and renal site PA imaging (Fig. 5j) were conducted simultaneously. The NIR fluorescence and PA intensities increased gradually over time and peaked at 2 h after injection. However, these signals were significantly reduced in mice that received pre-treatment with Que or NAC (Fig. 5l,m). Similar results were observed in the analysis of the mice urine (Fig. 5h,n). To confirm that CDIA@LMWC NP was activated in situ in the kidneys, ex vivo fluorescence images of the kidneys and other organs outside the kidneys were taken. Fluorescence signals were detected solely in the excised kidneys of mice 3 h after the injection of CDIA@LMWC NPs, consistent with the in vivo NIR imaging data (Supplementary Fig. 51). This signal was 8.25 times greater than that observed in the control mice. The NIR fluorescence and PA signals indicated that CDIA@LMWC NP accumulated efficiently in the kidney region and reacted with •OH, which was overexpressed in AKI mice.

Molecular Mechanism Responsible for the Protective Effects of Pue on AKI

Initial research indicates that Pue has hepatoprotective properties in rats with CCl₄-induced liver damage by reducing OS and inflammation, leading to improved intrahepatic metabolism.^[57] Considering Pue was screened to be a potential antioxidant agent in vitro, we further tested the protective effects of Pue in vivo and explored the molecular mechanism for the antioxidant effects of Pue on AKI. Mice were treated with saline, 18 mg/kg CDDP, or Pue (5 mg/kg or 10 mg/kg, i.v. injection) 3 days prior to CDDP administration (Fig. 6a). PA images of mice kidney revealed that PA amplitude increased after CDDP administration while efficiently reduced if the mice were pre-treated with Pue (Fig. 6b,c). Pathological analysis on the kidney slices showed similar results (Fig. 6d). In order to gain a deeper understanding of how Pue achieved the restoration of redox homeostasis in the progression of AKI, we examined alterations in crucial signaling molecules within the ROS regulatory network. Sirtuin 1 (Sirt1), a NAD⁺-dependent histone deacetylase, is a critical component in maintaining cellular redox balance and resistance to OS.^[58] The transcription factor NF-E2 p45-related factor 2 (Nrf2), a transcription factor that plays a crucial role in cellular antioxidant pathways, is under the control of Sirt1 and can be reduced by excessive ROS levels.^[59–61] To evaluate the levels of Sirt1, Nrf2, and associated proteins such as haem oxygenase-1 (HO-1), DJ-1, and Kelch-like ECH-associated protein 1 (Keap1) in vitro, both qRT-PCR and western blot analyses were utilized.^[62–64] A qRT-PCR analysis was performed to determine the status of the Sirt1/Nrf2 and DJ-1/Nrf2/Keap1 axis. The results showed that the relative expressions of Sirt1, Nrf2, DJ-1, and HO-1 were significantly increased, while Keap1 was decreased at the mRNA level in HK-2 cells that were pre-treated with Pue (Fig. 6e–i). These findings were further supported by the western blot analysis, which revealed that Pue could elevate the expression of DJ-1, the protein that was typically suppressed under OS.^[65] Subsequently, there was an upregulation of Sirt 1, Nrf2, and HO-1, as well as a downregulation of Keap1. No significant changes in the expression of these proteins were observed in HK-2 cells treated with Pue alone (Fig. 6j–n). After translocation to the nucleus, Nrf2 bound to antioxidant response elements, resulting in the upregulation of the antioxidant gene (HO-1) and a decrease in •OH content. Moreover, DJ-1 prevented the association of Nrf2 with its inhibitor protein, Keap1, and subsequent ubiquitination of Nrf2. In cases of low DJ-1 expression, Nrf2 was ubiquitinated to a greater extent, which was associated with a decrease of Nrf2 protein. Furthermore, the treatment of CDDP inhibited the activity of superoxide dismutase (SOD) and increased the level of MDA in HK-2 cells. However, pre-treatment of Pue could reversed the ROS-related imbalance induced by CDDP (Fig. 6o,p). The results suggest that Pue may exert a protective effect on cells by modulating the expression of key proteins involved in antioxidant defense pathways. In addition, Pue was found to have an IC₅₀ value of 80.52 ± 1.22 µM (Fig. 6s) for •OH scavenging. Like Que (Fig. 6r), Pue could directly react with •OH to achieve the effect of antioxidation. In brief, the data showed that Pue effectively decomposed the predominant cellular ROS, •OH, induced by CDDP into harmless substances like H₂O and O₂. Additionally, the remaining Pue activated the Sirt1/Nrf2/Keap1 signaling pathway. These results highlighted the significance of the reaction between Pue and •OH, as well as the crucial role of Pue in activating the Sirt1/Nrf2/Keap1 signaling pathway to quench ROS (Fig. 6q). Ultimately, this protective mechanism prevented renal cell apoptosis and underscored the potential therapeutic value of Pue in protecting kidney.

To sum up, a probe has been created that can detect AKI through the activation of NIR fluorescence and PA signal in response to •OH. The dual-mode imaging probe comprises a NIR hemi-cyanine dye (CyOH) coupled with a •OH-responsive substrate iodosalicylic acid. With high sensitivity and specificity towards •OH, CDIA possesses a fast activation kinetics and high biocompatibility. After introduction of LMWC for enhancing renal targeting, CDIA@LMWC NP accumulates in the kidney without any active intervention and triggers its fluorescence and PA signals to indicate the expression level of OS related to AKI. The positive time of 12 h using this probe is superior to that of 48 h for typically clinical assays including BUN and sCr detection. Moreover, CDIA@LMWC NP displays excellent ability for HTS of natural •OH scavengers from herbal medicines, uncovering additional mechanisms in the chemical regulation of natural products to attenuate AKI. Additional investigations have revealed that Pue, a natural antioxidant medication selected through screening, shields renal cells in cases of AKI by disintegrating CDDP-induced •OH. Furthermore, it activates the Sirt1/Nrf2/Keap1 signaling pathway to regulate ROS-related genes, which restores redox homeostasis in the renal cells. Given its effortless preparation and reactive activity that is dependent on the situation, we predict that the duplex imaging probe CDIA could be a potential option for detecting clinical AKI and other ROS-related ailments in patients, thus illuminating the ROS-related pathological process and improving the overall diagnostic efficacy.

Materials and Reagents

All the oligonucleotides used in this study were custom-synthesized by Sangon Biological Engineering Technology & Services Co., Ltd. (Shanghai, China). Low molecular weight chitosan (LMWC > 90% deacetylation) with number-average molecular weight of 5000 was purchased from Sigma-Aldrich (St. Louis, MO, USA). Sodium tripolyphosphate (TPP, purity > 98%) was purchased from Macklin Biochemical Co., Ltd (Shanghai, China). Sulfo-Cyanine7 carboxylic acid (Cy7) was obtained from Xi’an ruixi Biological Technology Co., Ltd (Xi’an, China). 3-(4,5-dimethylthiazol)-2,5-diphenyltetrazolium-bromide (MTT), dimethyl sulphoxide (DMSO), Dulbecco's Modified Eagle Medium/Nutrient Mixture F-12 (DMEM/F12), Dulbecco's Modified Eagle Medium (DMEM), fetal bovine serum (FBS) and phosphate buffer solutions (PBS, 137 mM NaCl, 2.7 mM KCl, 4.3 mM Na₂HPO4, 1.4 mM KH₂PO4, pH 7.4) were purchased from Gibco (Gaithersburg, USA). Fluorescein isothiocyanate (FITC, purity > 95%), lipopolysaccharide (LPS), vitamin C (VC), citrinin (CTN), aristolochic acid Ⅰ (AAⅠ), cis-diammineplatinum dichloride (CDDP), quercetin (Que), puerarin (PUE) and other reagents were purchased from Sigma-Aldrich Reagent, Ltd. (Shanghai, China). Trizol RNA isolation reagent, LysoTracker® Green, MitoTracker® Green, and Hoechst 33342 were purchased from Invitrogen Company (Carlsbad, CA, USA). A stock solution (1 mM) of CDIA was prepared by dissolving an appropriate amount of CDIA in chromatogram class DMSO. The HiScript II Q RT SuperMix for qPCR, HiScript II One Step qRT-PCR SYBR Green Kit, 180 kDa Prestained Protein Marker, One-Step PAGE Gel Fast Preparation Kit (10%) and enhanced chemiluminescence (ECL) detection kit were purchased from Vazyme Biotech Co., Ltd. (Nanjing, China). Enhanced BCA Protein Assay Kit, Total Superoxide Dismutase Assay Kit with WST-8 and Lipid Peroxidation MDA Assay Kit were purchased from Beyotime Biotech Co., Ltd. (Shanghai, China). Serum creatinine (sCr) and blood urea nitrogen (BUN) assay kits were obtained from Nanjing Jiancheng Biotech Co., Ltd (Nanjing, China). All the organic solvents were of analytical grade or the best grade available. All aqueous solutions were prepared in ultra-pure water obtained from a Milli-Q water purification system (18 M Ω cm).

Apparatus

The compound was characterized by ¹H NMR and mass spectra. ¹H NMR spectrum was measured on a Bruker DRX600 spectrometer and referenced to the residual proton signals of the solvent. Absorption spectra were recorded on an UV-2550 UV-Vis-NIR spectrophotometer (Shimadzu Company, Japan). Fluorescence spectra were measured on an F-7000 spectrofluorometer (Hitachi, Japan). HPLC spectrum was measured with a 1260 Infinity II spectrometer (Agilent Technologies, USA). Mass spectrum was measured with a InfinityLab LC/MSD Series spectrometer (Agilent Technologies, USA). All pH measurements were carried out with a Sartorius PB-10 (Sartorius scientific instruments, Beijing, China) containing a composite glass electrode. The morphology of the nanoprobe was characterized at a HITACHI HT7700 transmission electron microscope (TEM) operated at 120 kV. The particle size and size distribution were obtained by dynamic light scattering (DLS) at 25°C by means of a 90 Plus/BI-MAS equipment (Brookhaven, USA). Zeta potential measurement was performed at 25°C on a Malvern Zetasizer-Nano Z instrument. MTT assay was performed using a microplate reader (Infinite M200 Pro, Tecan). Confocal fluorescence imaging of cells was performed on a confocal laser scanning microscope (LSM800, Zeiss, Germany) and processed using the ZEN imaging software. Flow cytometric assay was performed using MACSQuant Analyzer 10 (Miltenyi Biotec, Germany). PCR reactions were performed on the Quant Studio 6 Flex Real-Time PCR System (Applied Biosystems, Carlsbad, California, USA). In vivo fluorescence imaging experiments were performed on a live animal imaging system (Tanon ABL X6, China). The hematoxylin and eosin (H&E) staining images were acquired on a digital pathology slice scanner using NanoZoomer 2.0 RS (Hamamatsu, China). The photoacoustic (PA) signal of diseases areas were measured by Visualsonic Vevo 2100 LAZER system.

Measurement of Fluorescence Quantum Yields

The fluorescence quantum yields were measured using Cy7 in DMSO (Φ_F = 0.3) as the standard and calculated with the following equation,

Φ _{F (x)} = Φ_{F (std)} (\(\frac{{{A}}_{{s}{t}{d}}}{{{A}}_{{x}}})\) (\(\frac{{{I}}_{{x}}}{{{I}}_{{s}{t}{d}}})\) (\(\frac{{{\eta }}_{{x}}}{{{\eta }}_{{s}{t}{d}}})\)²

where subscript x designates CDIA, subscript std designates Cy7, A stands for the absorbance at excitation wavelength, I stands for the integrated fluorescence intensity, and η stands for the refractive index of the solvent in the measurement.

Preparation of Analyte Solutions

•OH: Hydroxyl radical was generated by Fenton reaction. To generate •OH, various amounts of Fenton reagent (Fe²⁺/H₂O₂ = 1/6, molar ratio) were added into DMF/0.1 mM H₂SO₄ = 1/1 (v/v).^{[40] 1}O₂: Singlet oxygen was generated in situ by addition of the H₂O₂ stock into a solution containing 10 eq of HClO. ONOO⁻: A mixture of sodium nitrite (0.6 M) and hydrogen peroxide (0.7 M) was acidified with hydrochloric acid (0.6 M), and sodium hydroxide (1.5 M) was added within 1–2 s to make the solution alkaline. The excess hydrogen peroxide was removed by passing the solution through a short column of manganese dioxide. The peroxynitrite concentration was estimated by using an extinction coefficient of 1670 M^− 1•cm^− 1 at 302 nm. C_ONOO− = Abs₃₀₂ nm /1.67 (mM).^[66]NO: Nitric oxide was used from a stock solution prepared by sodium nitroprusside. ClO⁻: hypochlorite was delivered from commercial aqueous solution. TBHP: Tert-Butyl hydroperoxide was delivered from commercial aqueous solution. H₂O₂: Hydrogen peroxide was delivered from commercial aqueous solution. O₂^•−: Superoxide solution was prepared by adding KO₂ to dry dimethylsulfoxide and stirring vigorously for 10 min. T₄: Freshly prepared thyroxine solution (5 mM) was added directly. T₃: Freshly prepared triiodothyronine solution (5 mM) was added directly. TEMPO: Freshly prepared stock solution of 2,2,6,6-tetramethyl-1-piperidinyloxy (100 mM) was added directly. Thiourea: Freshly prepared thiourea (100 mM) was added directly. GSH: Glutathione (100 mM) was added directly. NAC: N-Acetylcysteine (100 mM) was added directly.

Synthesis of CDIA@LMWC NP

0.25 mL of dimethyl sulfoxide (DMSO) containing 5 mg of CDIA was added to 4.25 mL of LMWC solution (5 mg of LMWC) to obtain a mixed solution. 2 mL 1.25 mg/mL TPP solution was added dropwise into the above mixture solution while stirring at 1000 rpm. The conjugation reaction was maintained for 1 h, and then the product was then purified by continuous dialysis (molar weight cutoff, MWCO 10000) against deionized water for 48 h. The final product was freeze-dried and kept in desiccators for use.

Synthesis of FITC-LMWC

5 mg of FITC was dissolved in 500 µL DMSO and added to 50 mg LMWC immersed in 5 mL deionized water. After 16 h of incubation at room temperature in the dark, the mixture was dialyzed (MWCO 1000) for 48 h at room temperature in the dark, followed by lyophilization.

Synthesis of FITC-LMWC NP

FITC-LMWC NP was synthesized by using an ionic crosslinking method.^[67] In brief, 5 mg FITC-LMWC was dissolved in 5 mL 1% acetic acid solution, 2 mL 1.25 mg/mL TPP solution was added dropwise into FITC-LMWC solution while stirring at 1000 rpm. The conjugation reaction was maintained for 30 min, and the product was then purified by successive dialysis (MWCO 10000) against deionized water for 48 h. Then the product solution was adjusted to 7.4. The final product was freeze-dried and kept in desiccators for use. The amount of FITC conjugated on FITC-LMWC NP was determined by absorption at 490 nm according to a previous method.^[68]

Fluorescence Measurement of FITC-LMWC NP

FITC-LMWC NP was diluted with deionized water to obtain final concentrations: 0, 1, 2, 3, 4, 5, 6, 7, 8, 9, 10 µg/mL. The solution was excited at 490 nm and the emission wavelengths were collected from 500 to 700 nm. The fluorescence spectra was measured on Cary Eclipse Fluorescence Spectrophotometer and both excitation and emission slits were set to 5.0 nm. The standard curve of fluorescence intensity was recorded at λ_ex/λ_em = 490/518 nm using a spectrofluorometer.

Characterization of Nanoparticles

TEM measurement was performed with a JEOL JEM-200CX TEM at an accelerating voltage of 200 kV. The samples were prepared by dropping 10 µL of 10 µM CDIA onto copper grids, and dried for 3 min. Then the residual solution was blotted off using filter paper. Dynamic light scattering and zeta potential measurements of LMWC NP, FITC-LMWC NP and CDIA@LMWC NP were performed at 25°C on Mastersizer 2000 particle size analyzer and Malvern Zeta sizer-Nano Z instrument. The samples were dispersed in double distilled water and the pH value of the solution was adjusted to 7.4 before the analysis.

Determination of EE and LC

To determine the encapsulation efficiency (EE) of CDIA@LMWC NP, a dialysis method has been used.^[69] 10 mL freshly prepared CDIA@LMWC NP containing 7.14 mg CDIA was placed in a dialysis bag (MWCO 5000), immersed in 90 mL phosphate buffer solution and shaken at 500 rpm using a magnetic stirrer. 5 mL dialysis sample solution was collected and the concentration of probe was measured by the UV-Vis-NIR absorption method. The EE of CDIA was calculated as: EE (%) = (W₁ − W₂)/W₁×100, where W₁ and W₂ were the weights of added CDIA and unloaded CDIA, respectively. The loading content (LC) of CDIA was calculated as: LC (%) = (W₁ − W₂)/W_t×100, where W₁ and W₂ were the weights of added CDIA and unloaded CDIA, respectively, and W_t was the weight of LMWC NP. The determinations were performed at least five times for each calculation.

In Vitro Release of CDIA

The in vitro release profile of CDIA release from CDIA@LMWC NP was carried out at pH 7.4, 6.8, 5.0, and 4.0 using a dialysis method with free CDIA solution at the same concentration as a control. 1 mL of 1 mg/mL CDIA@LMWC NP was placed in a dialysis bag (MWCO 5000), immersed in 50 mL PBS and incubated at 37°C in a shaking platform at 500 rpm. At predetermined intervals, 1 mL dissolution sample was collected and replaced with the same volume of release medium. The concentration of CDIA released from CDIA@LMWC NP was measured by the UV absorption method as mentioned.

General Procedure for UV-Vis-NIR Absorption Spectra Measurement

All the UV-Vis-NIR absorption spectra measurements were performed in 10 mM phosphate buffer (pH 7.4). To build a standard curve of CDIA content, different final concentrations (0, 1, 2.5, 5, 7.5, 10, 15, 20 µM) of CDIA was added into PBS buffer. The absorbance at 650 nm was measured using a spectrophotometer. To build a standard curve of FITC content, different final concentrations (0, 3, 6, 9, 12, 15 µg/mL) of FITC was added into PBS buffer. The absorbance at 490 nm was measured using a spectrophotometer. For •OH detection, 10 µM CDIA was added into different concentration of freshly prepared •OH (0–50 µM, prepared by Fenton reaction). After rapidly mixing the solution and incubating at 37°C for 30 min in a thermostat, a 3 mL portion of the reaction solution was transferred to a 10 × 10 mm quartz cell for in vitro detection.

General Procedure for Fluorescence Spectra Measurement

CDIA was dissolved in DMSO to make a 5 mM stock solution, which was diluted to the required concentration of testing solution for measurement. Fluorescence spectra was recorded with λ_ex/_em = 650/720 nm and both excitation and emission slit widths were set to 5.0 nm. Meanwhile, a blank solution without •OH (control) was prepared and measured under the same condition for comparison. For •OH response detection, 50 µL 200 µM CDIA was dissolved in 450 µL PBS, and the solution was incubated with 500 µL Fenton reagent at various concentrations (0 − 200 µM) at 37°C for 30 min. The fluorescence spectra of CDIA were recorded using a spectrofluorometer with the excitation wavelength at 650 nm and the emission wavelengths from 675 to 875 nm.

For CDIA selectivity detection, different kinds of analyte solutions were incubated with 10 µM CDIA solution at 37°C for 30 min, respectively, and then recorded using a spectrofluorometer. All the experiments were repeated at least five times.

For fluorescence kinetics measurement, 10 µM CDIA was added into Fenton reagent at various concentrations (0, 10, 30, 50, 100 µM) at 37°C, respectively. The fluorescence intensity was examined with increasing time (0, 0.5, 1, 2.5, 5, 7.5, 10, 12.5, 15, 17.5, 20, 22.5, 25, 27.5, 30 minutes). Then the fluorescence was excited at 650 nm and measured at 720 nm. All the experiments were repeated at least five times.

Cell Culture

AML-12 cells, H9c2 cells, HUVEC cells, MCF-7 cells and HK-2 cells were obtained from KeyGEN Biotech Co., Ltd (Nanjing, China). AML-12, H9c2, HUVEC and MCF-7 cells were routinely grown in Dulbecco's modified Eagle's medium (DMEM) containing 10% FBS, 100 µg/mL streptomycin, and 100 U/mL penicillin. HK-2 cells were cultured in Dulbecco's modified Eagle's medium/Ham's nutrient mixture F12 (DMEM/F12) supplemented with 10% FBS, 100 µg/mL streptomycin, and 100 U/mL penicillin. The above cells were grown in a 100% humidified atmosphere containing 5% CO₂ at 37°C. The medium was replenished every day and the cells were subcultured after reaching confluence.

Cytotoxicity Assay

To investigate the cytotoxicity of CDIA, standard MTT tests were performed in AML-12, H9c2, HUVEC, MCF-7 and HK-2 cells. HepG2 cells growing in log phase were seeded into 96-well plates at a seeding density of 1 × 10⁴ cells per well in 200 µL complete medium and allowed to attach for 24 h. After rinsing with PBS, HepG2 cells were incubated with 200 µL culture media containing serial concentrations of CDIA (0, 1, 2, 5, 10, 15, 20 µg/mL) for 24 h. Then 20 µL of MTT (5 mg/mL in PBS) was added to each well and followed by incubated for 4 h under the respective conditions. After 4 h, the supernatants containing unreacted MTT were discarded carefully, and 150 µL DMSO was added to each well to dissolve the produced blue formazan. The absorbance was recorded at 570 nm using a microplate reader after 10 min of shaking. Cell viability was determined by the following formula: Cell viability (%) = 100 × (OD₁-OD₂)/(OD₃-OD₂), where OD₁, OD₂, and OD₃ were OD value of treatment group, OD value of blank group, and OD value of control group, respectively. All the experiments were repeated at least five times. The similar cytotoxicity assay was applied to nanoparticles (LMWC NP, FITC-LMWC NP and CDIA@LMWC NP) of concentrations from 0 to 2 mg/mL to HK-2 AML-12, H9c2, MCF-7 and HK-2 cells, respectively.

To evaluate the cytotoxicity of CDDP, AAI, and CTN to HK-2 cells, HK-2 cells were dispersed in 96-well microtiter plates at 1 × 10⁴ cells per well in a total volume of 200 µL. The plates were maintained in a humidified atmosphere with 5% CO₂ at 37°C for 24 h. After the original medium has been discarded, all the drugs with 0.1% DMSO at final concentrations of 1 ~ 1000 µM was diluted in fresh medium, then the medium was added to the HK-2 cells respectively, and incubated for 24 h. The cell viability test methods were the same as above.

Confocal Fluorescence Imaging of •OH in Model Cells

Several different cell models were established: (i) PMA (phorbol myristate acetate) was applied to induce endogenous •OH. RAW264.7 mouse macrophages and HeLa cells were exposed to PMA (200 or 500 ng/mL) diluted with serum-free medium for 4 h at 37°C, respectively. (ii) Acetaminophen (APAP) is a component of many cold medications, which may cause liver injury and oxidative stress in liver with overdose. HepG2 cells were exposed to APAP (5 or 10 mM) diluted with serum-free medium for 24 h at 37°C. (iii) HK-2 cells were exposed to 75 µM CDDP diluted with serum-free medium for 24 h at 37°C. (iv) HK-2 cells were exposed to 44 µM AAI diluted with serum-free medium for 24 h at 37°C. (v) HK-2 cells were exposed to 30 µM CTN diluted with serum-free medium for 24 h at 37°C. After incubation, the above cells were stained with CDIA (10 µM for 30 min) or CDIA@LMWC NP (20 µg/mL for 2 h), and then incubated with 5 µg/mL Hoechst 33342 for 25 min, rinsed three times with PBS (pH 7.4) to perform fluorescence imaging with a CLSM at stationary parameters including the laser intensity, exposure time, and objective lens. Hoechst 33342 was excited at 405 nm with a violet laser diode and the emission was collected from 420 to 480 nm. CDIA or CDIA@LMWC NP was excited with a 640 nm helium − neon laser and emission was collected from 650 to 750 nm. All images were digitized and analyzed by a ZEN imaging software.

Confocal Fluorescence Imaging for Determination of Optimal Incubation Conditions

HK-2 cells were seed into 35-mm confocal dishes (Glass Bottom Dish) at a density of 1 × 10⁴ per dish and incubated in complete medium at 37°C. After 24 h, the medium was replaced with fresh serum-free culture medium containing FITC-LMWC NP (0.5 µg/mL, 1 µg/mL, 2 µg/mL, 5 µg/mL, 10 µg/mL, 20 µg/mL) and incubated at 37°C for 10 min, 40 min, 100 min and 130 min, respectively. Cell imaging was then carried out after washing cells three times with PBS buffer (pH 7.4) to remove any FITC-LMWC NP on the surface of cells. The fluorescence of cells was visualized using LSM800 with a Plan-Apochromat 63×/1.40 Oil DIC M27 lens (Zeiss) at stationary parameters including the laser intensity and exposure time. The fluorescence signal of cells incubated with FITC-LMWC NP was excited at 488 nm with an argon ion laser and the emission was collected from 500 to 650 nm.

Cellular Uptake Assay

The cellular uptake assay of FITC-LMWC NP was observed using confocal fluorescence imaging, HK-2 AML-12, H9c2, HUVEC, MCF-7 and HK-2 cells were seeded into 35-mm confocal dishes (Glass Bottom Dish) at a density of 1 × 10⁴ per dish and incubated in complete medium for 24 h at 37°C. Then the above cells were exposed to 20 µg/mL FITC-LMWC NP diluted with serum-free medium for 100 min at 37°C, respectively. After incubation, the cells were stained with 1.0 µM Hoechst 33342 for 25 min, rinsed three times with PBS (pH 7.4) to perform fluorescence imaging with a LSM800 at stationary parameters including the laser intensity, exposure time, and objective lens. Hoechst 33342 was excited with a violet 405 nm laser diode and the emission was collected from 420 to 500 nm. FITC was excited at 488 nm with an argon ion laser and the emission was collected from 500 to 650 nm. All images were digitized and analyzed by a ZEN imaging software.

For competitive inhibition assay, HK-2 cells were seeded into 35-mm confocal dishes (Glass Bottom Dish) at a density of 1 × 10⁴ per dish and incubated with 10 µg/mL FITC-LMWC NP diluted into serum-free medium in the absence or presence of 1 µM EDTA, 50 µg/mL LMWC, or 50 µg/mL glucosamine for 2 h at 37°C. Once uptake was terminated, the cells were rinsed three times with PBS (pH 7.4) to remove extracellular FITC-LMWC NP. The fluorescence of cells was visualized using LSM800 and fluorescence signal of cells incubated with FITC-LMWC NP was excited at 488 nm with an argon ion laser and the emission was collected from 500 to 650 nm. All images were analyzed by a ImageJ software.

Flow Cytometric Assay

For flow cytometric assay, cells were seeded at a density of 2 × 10⁵ cells per well in 6-well plates and incubated with different treatments at 37°C for different times to produce normoxic or hypoxic cells. After a certain time, the medium was replaced with cell culture medium containing CDIA (10 µM) or CDIA@LMWC NP (20 µg/mL). After incubation, the different treatment groups of cells were washed three times with PBS solution, trypsinized, harvested, rinsed with PBS and resuspended in 500 µL PBS medium, and subjected to flow cytometric assay using MACSQuant Analyzer 10. The data were analyzed with FCS Express V3.

Cellular ROS Detection

For detection of •OH production, HK-2 cells were dispersed in 96-well microtiter plates at 1 × 10⁴ cells per well in a total volume of 200 µL. The plates were maintained in a humidified atmosphere with 5% CO₂ at 37°C for 24 h. After rinsing with PBS three times, HK-2 cells were incubated with 200 µL of culture media containing serial concentrations (0, 5, 10, 20, 40, 60, 80, 100 µM) of CDDP, AAI and CTN for 24 h at 37°C, respectively. After incubation, the HK-2 cells were treated with 10 µM commercial ROS probe dichlorodihydrofluorescein diacetate (H₂DCFDA) for 30 min, and then the fluorescence intensity was measured using a spectrofluorometer at λ_ex/λ_em = 492/524 nm.

RNA Extraction and qRT-PCR

To study the protective effect of Pue on the HK-2 cells, 75 µM CDDP and 25 µM Pue or 50 µM Pue were administered, respectively. HK-2 cells were pretreated with 25 µM Pue or 50 µM Pue for 12 h before 75 µM CDDP administration. The total RNA was extracted from the HK-2 cells of each group to detect Sirt11, HO-1, Nrf2, Keap,1, and DJ-1 mRNA levels.

Total RNA was isolated from HK-2 cells using Trizol reagent, and cDNA was prepared using a HiScript II Q RT SuperMix for qPCR. Quantitative RT-PCR was carried out using HiScript II One Step qRT-PCR SYBR Green Kit with 20 µL reaction mixtures. Primer names and sequences for qRT-PCR are listed in Supplementary Table S6. PCR reactions were performed on the Quant Studio 6 Flex Real-Time PCR System with the following program: step 1, 95°C for 3 min to activate the Taq polymerase; step 2, 95°C for 3 s to denaturize DNA; step 3, 60°C for 31 s for annealing/extension (39 cycles for steps 2 and 3). The relative mRNA levels were quantified by the 2^−ΔΔCt method and all data were normalized to GAPDH (the internal control).

Western Blot Analysis

Immunoblotting analysis of proteins in HK-2 cells was performed by homogenizing frozen cells in the lysis buffer containing 50 mM Tris-HCl, 150 mM sodium chloride, 2 mM ethylene diamine tetraacetic acid (EDTA), 2 mM ethylene glycol tetraacetic acid (EGTA), 1% Triton-X 100, and a protease inhibitor (pH 7.4). The supernatants of the mixture were obtained by centrifugation at 4°C (30 min, 16,000 × g), and their total protein concentrations were further determined using the Enhanced BCA Protein Assay Kit. Protein extracts were separated on sodium dodecyl sulfatepolyacrylamide gel electrophoresis (SDS-PAGE, 8–12% gels) and blotted onto PVDF membranes. After blocking with 5% fatfree milk, the membranes were incubated at 4°C overnight with the following primary antibodies: anti-Nrf2 (1:1000, A21176), anti-Keap1 (1:1000, A1820), anti-HO-1 (1:1000, A19062), anti-DJ-1 (1:1000, A19097), anti-Sirt1 (1:1000, A19667), and anti-GAPDH (1:1000, A19056) from ABclonal Biotech Co., Ltd. (Wuhan, China). The bound antibodies were detected using horseradish peroxidase (HRP)-conjugated IgG (ABclonal) and visualized with enhanced chemiluminescence (ECL) detection reagents. GAPDH was used as a loading control.

MDA and SOD Assays

To assess the OS level in HK-2 cells, the levels of SOD and MDA were tested with commercially available kits (Beyotime Biotechnology, China). Briefly, HK-2 cells were homogenized in ice-cold 0.1 M phosphate buffer (pH 7.4), then the homogenates were filtered and centrifuged using a refrigerated centrifuge at 4°C (20 min, 16,000 × g). The obtained supernatants were used to determine the SOD enzyme activity and the lipid peroxidation level by measuring MDA content. The SOD enzyme activity was expressed as a unit of activity per milligram of protein and the MDA content was expressed as micromole per gram of protein.

Animal Model

ICR-Swiss male mice (20–25 g), 5–6 weeks of age, were purchased from the Laboratory Animal Center and Institute of Comparative Medicine at Yangzhou University. The animals were bred in stainless steel metabolic cages with free access to food and double distilled water under standard conditions of humidity (50 ± 10%), temperature (25 ± 2°C) and light (12 h light/12 h dark cycle). All animal study protocols were designed in according to guidelines set by the National Institute of Health Guide for the Care and Use of Laboratory Animals, and approved by the Animal Ethical Experimentation Committee of China Pharmaceutical University. The mice were randomly assigned to 4 experimental groups (group I: mice were treat once with CDDP at a dosage of 18 mg/kg; group II: mice were treated twice with CDDP at a dosage of 9 mg/kg; group III: mice were treated thrice with CDDP at a dosage of 6 mg/kg; group IV: mice were treated with CDDP at a dosage of 3 mg/kg for six times.). The control groups were treated with PBS (0.2 ml) or NAC (10 mg kg^− 1 day^− 1 during the study, i.p. injection) or quercetin (Que, 10 mg kg^− 1 day^− 1 during the study, i.p. injection ) 3 days prior to CDDP administration.^[70] NIR fluorescence and PA imaging were performed for 2 h after i.v. injection of 10 mg/kg CDIA@LMWC NP.

Hemolysis Assay

Hemolytic acitvity of CDIA was carried out according to the previous protocol.^[71] Fresh mouse blood was diluted by physiological saline and centrifuged to isolate red blood cells (RBCs) from serum. Then RBCs were incubated with physiological saline (negative control), triton X-100 (10 g L^− 1, positive control), or CDIA at concentrations of 1, 2.5, 5, 10, 20, and 40 µM at 37°C for 2 h. After incubation, RBCs were centrifuged at 3000 r.p.m. for 10 min and the supernatant of each sample was collected and transferred to a 96-well plate. The absorbance of hemoglobin at 540 nm was measured by using a microplate reader. Because CDIA had intrinsic absorption at 540 nm, the absorption of sample group (A_sample) was deducted by the absoption of CDIA at 540 nm at the same concentration. The hemolysis ratio of RBCs was calculated as follows: Hemolysis (%) = (A_sample – A_negative)/ (A_positive – A_negative) ×100%, where A_sample, A_negative, A_positive were denoted as the absortion of samples (after deduction by the intrinsic absorption of CDIA), negative and positive control, respectively. The same operation was used for the hemolysis test of CDIA@LMWC NP.

In Vivo NIR Fluorescence Imaging in Living Mice

NIR fluorescence imaging were conducted for 2 h after i.v. injection of CDIA@LMWC NP. Fluorescence images of CDIA@LMWC NP were acquired using a live animal imaging system (Tanon ABL X6, China) with excitation at 650 ± 10 nm and emission at 720 ± 10 nm and an acquisition time of 0.1 s. NIR fluorescence intensities of kidneys in living mice were analyzed by the region of interest (ROI) analysis. Mice were euthanized after imaging at different timepoints post-treatment of cisplatin. Major organs were collected and placed into 4% paraformaldehyde for histological examination.

PA Imaging

PA imaging was performed using a LAZR Tight TM imaging enclosure coupled with a Vevo LAZR PA imaging system (Fujifilm VisualSonics) equipped with a MS-250 linear array transducer (21 MHz, 70% 6 dB two-way bandwidth, 256 elements) to detect ultrasound (US) signals and a tunable Nd: YAG laser system (OPOTEK, 680 − 970 nm, 20 Hz repetition rate, 5 ns pulse width, and 50 mJ pulse peak energy) to generate optical pulses. PA spectra were acquired using a transducer at a wavelength of 690 nm and with an acquisition rate of 5 frames/s. US/PA signals were processed and reconstructed in a work station. The energy supplied by each pulse of the tunable laser was 1.2 mJ/cm², well below the standard set by the American National Standard Institute across the wavelength range. PA imaging in living mice were conducted for 2 h after i.v. injection of CDIA@LMWC NP with excitation at 690 nm.

Histology

The organs were fixed with 4% paraformaldehyde (PFA), dehydrated in ethanol solution, and embedded in paraffin prior to 10 µm section. Histology samples were stained by hematoxylin and eosin under standard protocols. Images were acquired using a slide scanner (NanoZoomer 2.0 RS, Hamamatsu).

Blood Sample Analysis

The blood samples were collected at different time post treatment in the heparin capillaries through orbital venous of mice, and centrifuge for 20 min (4 ℃, 2000 rpm). The supernatant serum were collected and stored at − 80 ℃. BUN and sCr concentration were determined using commercial kits according to the manufacturer’s protocol (5 mice were analyzed for each sample).

Data Analysis

Results were expressed as the mean ± standard deviation unless stated otherwise. Cell imaging data was analyzed and quantified with Image J and ZEN software. Statistical comparisons between groups were determined by t-test and more than three groups were determined by one-way ANOVA with multiple comparisons test. For all tests, P < 0.05 was considered statistically significant. All statistical calculations were performed using GraphPad Prism.

Acknowledgements

This research was supported by National Natural Science Foundation of China (21775166), Natural Science Foundation for Distinguished Young Scholars of Jiangsu Province (BK20180026), and Fundamental Research Funds for the Central Universities (2632023TD02). Haijun Xu thanks Open Research Fund of School of Chemistry and Chemical Engineering, Henan Normal University.

Data Availability

The main data supporting the results in this study are available within the paper and its Supplementary Information. The raw and analysed datasets generated during the study are too large to be publicly shared, yet they are available for research purposes from the corresponding author on reasonable request. Source data are provided with this paper.

Declaration of Interests

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

Ronco, C., Bellomo, R. & Kellum, J. A. Acute kidney injury. Lancet 394, 1949–1964 (2019).
Kirtane, A. J. et al. Serum blood urea nitrogen as an independent marker of subsequent mortality among patients with acute coronary syndromes and normal to mildly reduced glomerular filtration rates. J. Am. Coll. Cardiol. 45, 1781–1786 (2005).
Priem, F. et al. β-trace protein in serum: a new marker of glomerular filtration rate in the creatinine-blind range. Clin. Chem. 45, 567–568 (1999).
Minhas, A. S. et al. Measuring kidney perfusion, ph, and renal clearance consecutively using mri and multispectral optoacoustic tomography. Mol. Imaging Biol. 22, 494–503 (2020).
Shuhendler, A. J., Pu, K., Cui, L., Uetrecht, J. P. & Rao, J. Real-time imaging of oxidative and nitrosative stress in the liver of live animals for drug-toxicity testing. Nat. Biotechnol. 32, 373–380 (2014).
Zhen, X. et al. Macrotheranostic probe with disease-activated near-infrared fluorescence, photoacoustic, and photothermal signals for imaging-guided therapy. Angew. Chem. Int. Ed. 57, 7804–7808 (2018).
Cheng, P. et al. Near-infrared fluorescence probes to detect reactive oxygen species for keloid diagnosis. Chem. Sci. 9, 6340–6347 (2018).
Zhang, J., Ning, L., Huang, J., Zhang, C. & Pu, K. Activatable molecular agents for cancer theranostics. Chem. Sci. 11, 618–630 (2020).
Miao, Q., Lyu, Y., Ding, D. & Pu, K. Semiconducting oligomer nanoparticles as an activatable photoacoustic probe with amplified brightness for in vivo imaging of pH. Adv. Mater. 28, 3662–3668 (2016).
Li, L.-L. et al. Pathological-condition-driven construction of supramolecular nanoassemblies for bacterial infection detection. Adv. Mater. 28, 254–262 (2016).
Pu, K. et al. Semiconducting polymer nanoparticles as photoacoustic molecular imaging probes in living mice. Nat. Nanotechnol. 9, 233–239 (2014).
Chen, N., Aleksa, K., Woodland, C., Rieder, M. & Koren, G. N-Acetylcysteine prevents ifosfamide-induced nephrotoxicity in rats. Brit. J. Phamacol. 153, 1364–1372 (2008).
Zhou, Z. et al. Activatable singlet oxygen generation from lipid hydroperoxide nanoparticles for cancer therapy. Angew. Chem. Int. Ed. 129, 6492–6496 (2017).
Hou, J. et al. Treating acute kidney injury with antioxidative black phosphorus nanosheets. Nano Lett. 20, 1447–1454 (2020).
Jiang, D. et al. DNA origami nanostructures can exhibit preferential renal uptake and alleviate acute kidney injury. Nat. Biomed. Eng. 2, 865–877 (2018).
Blum, A. P. et al. Stimuli-responsive nanomaterials for biomedical applications. J. Am. Chem. Soc. 137, 2140–2154 (2015).
Sahin, K. et al. Epigallocatechin-3-gallate activates Nrf2/HO-1 signaling pathway in cisplatin-induced nephrotoxicity in rats. Life Sci. 87, 240–245 (2010).
Ma, X. et al. Saikosaponin-D reduces cisplatin-induced nephrotoxicity by repressing ROS-mediated activation of MAPK and NF-kappaB signalling pathways. Int. Immunopharmacol. 28, 399–408 (2015).
Reddy, K. P, Madhu, P. & Reddy, P. S. Protective effects of resveratrol against cisplatin-induced testicular and epididymal toxicity in rats. Food Chem. Toxicol. 91, 65–72 (2016).
Palipoch, S. A review of oxidative stress in acute kidney injury: protective role of medicinal plants-derived antioxidants. Palipoch. Afr. J. Tradit. Complement Altern. Med. 10, 88–93 (2013).
Ratliff, B. B., Abdulmahdi, W., Pawar, R. & Wolin, M. S. Oxidant mechanisms in renal injury and disease. Antioxid. Redox Signal. 25, 119–146 (2016).
Paller, M. S., Hoidal, J. R. & Ferris, T. F. Oxygen free radicals in ischemic acute renal failure in the rat. J. Clin. Investig. 74, 1156–1164 (1984).
Baliga, R., Ueda, N., Walker, P. D. & Shah, S. V. Oxidant mechanisms in toxic acute renal failure. Drug Metab. Rev. 31, 971–997 (1999).
Heyman, S. N., Rosen, S., Khamaisi, M., Idee, J. M. & Rosenberger, C. Reactive oxygen species and the pathogenesis of radiocontrast-induced nephropathy. Invest. Radiol. 45, 188–195 (2010).
Pisani, A. et al. Role of reactive oxygen species in pathogenesis of radiocontrast-induced nephropathy. Biomed Res. Int. 2013, 868321 (2013).
Winterbourn, C. C. Reconciling the chemistry and biology of reactive oxygen species. Nat. Chem. Biol. 4, 278–286 (2018).
Fang, X., Schuchmann, H.-P. & Sonntag, C. von. The reaction of the OH radical with pentafluoro-, pentachloro-, pentabromo- and 2,4,6-triiodophenol in water: electron transfer vs. addition to the ring. J. Chem. Soc. Perkin Trans. 2, 1391–1398 (2000).
Aubin, N., Curet, O., Deffois, A. & Carter, C. Aspirin and salicylate protect against mptp-induced dopamine depletion in mice. J. Neurochem. 71, 1635–1642 (1998).
Yuan, L. et al. A unique approach to development of near-infrared fluorescent sensors for in vivo imaging. J. Am. Chem. Soc. 134, 13510–13523 (2012).
Miao, Q. et al. Near-infrared fluorescent molecular probe for sensitive imaging of keloid. Angew. Chem. Int. Ed. 57, 1256–1260 (2018).
Chen, H. et al. reversible ratiometric NADH sensing using semiconducting polymer dots. Angew. Chem. Int. Ed. 60, 2–8 (2021).
Eberhardt, M. K. & Colina, R. The reaction of OH radicals with dimethyl sulfoxide. a comparative study of Fenton's reagent and the radiolysis of aqueous dimethyl sulfoxide solutions. J. Org. Chem. 53, 1071–1074 (1988).
Wasil, M. et al. The specificity of thiourea, dimethylthiourea and dimethyl sulphoxide as scavengers of hydroxyl radicals. Biochem. J. 243, 867–870 (1987).
Yadav, A. & Mishra, P. C. Modeling the activity of glutathione as a hydroxyl radical scavenger considering its neutral non-zwitterionic form. J. Mol. Model. 19, 767–777 (2013).
Agnihotri, N. & Mishra, P. C. Mechanism of scavenging action of N-Acetylcysteine for the OH radical: a quantum computational study. J. Phys. Chem. B 113, 12096–12104 (2009).
Yu, H. et al. The antioxidant mechanism of nitroxide TEMPO: scavenging with glutathionyl radicals. RSC Adv. 5, 63655–63661 (2015).
Oziol, L. et al. In vitro free radical scavenging capacity of thyroid hormones and structural analogues. J. Endocrinol. 170, 197–206 (2001).
Bai, X., Huang, Y., Lu, M. & Yang, D. HKOH-1: a highly sensitive and selective fluorescent probe for detecting endogenous hydroxyl radicals in living cells. Angew. Chem. Int. Ed. 56, 12873–12877 (2017).
Pavkovic, M., Riefke, B. & Ellinger-Ziegelbauer, H. Urinary microRNA profiling for identification of biomarkers after cisplatin-induced kidney injury. Toxicology 324, 147–157 (2014).
Feng, W. et al. Lighting up NIR-II fluorescence in vivo: an activable probe for noninvasive hydroxyl radical imaging. Anal. Chem. 91, 15757−15762 (2019).
Du, K., Ramachandran, A. & Jaeschke, H. Oxidative stress during acetaminophen hepatotoxicity: sources, pathophysiological role and therapeutic potential. Redox Biol. 10, 148−156 (2016).
Qiao, H. et al. Kidney-specific drug delivery system for renal fibrosis based on coordination-driven assembly of catechol-derived chitosan. Biomaterials 35, 7157−7171 (2014).
Ramanery, F. P., Mansur, A. A., Mansur, H. S., Carvalho, S. M. & Fonseca, M. C. Biocompatible fluorescent core-shell nanoconjugates based on chitosan/Bi₂S₃ quantum dots. Nanoscale Res. Lett. 11, 187 (2016).
Perez Bay, A. E. et al. The fast-recycling receptor Megalin defines the apical recycling pathway of epithelial cells. Nat. Commun. 7, 11550 (2016).
Nielsen, R., Christensen, E. I. & Birn, H. Megalin and cubilin in proximal tubule protein reabsorption: from experimental models to human disease. Kidney Int. 89, 58−67 (2016).
Tong, Y., Huang, X., Lu, M., Yu, B.-Y. & Tian, J. Prediction of drug-induced nephrotoxicity with a hydroxyl radical and caspase light-up dual-signal nanoprobe. Anal. Chem. 90, 3556−3562 (2018).
Dolman, M. E., Harmsen, S., Storm, G., Hennink, W. E. & Kok, R. J. Drug targeting to the kidney: Advances in the active targeting of therapeutics to proximal tubular cells. Adv. Drug Delivery Rev. 62, 1344−1357 (2010).
Luo, Y. et al. A programmed nanoparticle with self-adapting for accurate cancer cell eradication and therapeutic self-reporting. Theranostics 7, 1245−1256 (2017).
Yuan, Z. X. et al. Specific renal uptake of randomly 50% N-acetylated low molecular weight chitosan. Mol. Pharmaceutics 6, 305−314 (2009).
Debelle, F. D., Vanherweghem, J.-L. & Nortier, J. L. Aristolochic acid nephropathy: a worldwide problem. Kidney Int. 74, 158–169 (2008).
Bouslimi, A. et al. Cytotoxicity and oxidative damage in kidney cells exposed to the mycotoxins ochratoxin a and citrinin: individual and combined effects. Toxicol. Mech. Methods 18, 341–349 (2008).
Xu, X. et al. Puerarin, isolated from Pueraria lobata (Willd.), protects against diabetic nephropathy by attenuating oxidative stress. Gene 591, 411–416 (2016).
Zheng, Y.-Z. et al. Antioxidant activity of quercetin and its glucosides from propolis: a theoretical study. Sci. Rep. 7, 7543 (2017).
Zhang, X. et al. Chlorogenic acid protects mice against lipopolysaccharide-induced acute lung injury. Injury, Int. J. Care Injured 41, 746–752 (2010).
Weng, Q. et al. Catalytic activity tunable ceria nanoparticles prevent chemotherapy-induced acute kidney injury without interference with chemotherapeutics. Nat. Commun. 12, 1436 (2021).
Chen, Y. et al. A promising NIR-II fluorescent sensor for peptide-mediated long-term monitoring of kidney dysfunction. Angew. Chem. Int. Ed. 60, 15809–15815 (2021).
Luo, C.-F. et al. UDP-glucuronosyltransferase 1A1 is the principal enzyme responsible for puerarin metabolism in human liver microsomes. Arch Toxicol. 86, 1681–1690 (2012).
Do, M. T., Kim, H. G., Choi, J. H. & Jeong, H. G. Metformin induces microRNA-34a to downregulate the Sirt1/Pgc-1α/Nrf2 pathway, leading to increased susceptibility of wild-type p53 cancer cells to oxidative stress and therapeutic agents. Free Radical Bio. Med. 74, 21–34 (2014).
Kulkarni, S. R. et al. Fasting induces nuclear factor e2-related factor 2 and ATP-binding cassette transporters via protein kinase A and sirtuin-1 in mouse and human. Antioxid. Redox Signaling 20, 15–30 (2014).
Zhang, Y. et al. Protective effects of dioscin against cisplatin‐induced nephrotoxicity via the microRNA‐34a/sirtuin 1 signalling pathway. Brit. J. Pharmacol. 174, 2512–2527 (2017).
Denicola, G. M. et al. Oncogene-induced Nrf2 transcription promotes ROS detoxification and tumorigenesis. Nature 475, 106–109 (2011).
Zhao, F. et al. Nrf2 promotes neuronal cell differentiation. Free Radical Bio. Med. 47, 867–879 (2009).
Clements, C. M., McNally, R. S., Conti, B. J., Mak, T. W. & Ting, J. P.-Y. DJ-1, a cancer- and Parkinson’s disease-associated protein, stabilizes the antioxidant transcriptional master regulator Nrf2. PNAS 103, 15091–15096 (2006).
Cuadrado, A. et al. Therapeutic targeting of the Nrf2 and Keap1 partnership in chronic diseases. Nat. Rev. Drug Discovery 18, 295–317 (2019).
Guzman, J. N. et al. Oxidant stress evoked by pacemaking in dopaminergic neurons is attenuated by DJ-1. Nature 468, 696–702 (2010).
Peng, T. et al. Molecular imaging of peroxynitrite with HKGreen-4 in live cells and tissues. J. Am. Chem. Soc. 136, 11728−11734 (2014).
Gao, S. et al. Megalin-mediated specific uptake of chitosan/siRNA nanoparticles in mouse kidney proximal tubule epithelial cells enables AQP1 gene silencing. Theranostics 4, 1039−1051 (2014).
Onishi, H. & Machida, Y. Biodegradation and distribution of water-soluble chitosan in mice. Biomaterials 20, 175−182 (1999).
Huang, L. et al. An artemisinin-mediated ROS evolving and dual protease light-up nanocapsule for real-time imaging of lysosomal tumor cell death. Biosens. Bioelectron. 92, 724−732 (2017).
Colbay, M. et al. Novel approach for the prevention. Exp. Toxicol. Pathol. 62, 81−89 (2010).
Li, M. et al. Cisplatin crosslinked pH-sensitive nanoparticles for efficient delivery of doxorubicin. Biomaterials 35, 3851−3864 (2014).

Supplementary Video S1 is not available with this version.

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SupportingInformation.docx
Supporting Information
VideoS2.m4v
Real-time monitoring of CDIA@FITC-LMWC NP in CDDP stimulated HK-2 cells at λex/em of 488/500–650 nm for FITC and λex/em of 640/650–750 nm for CDIA, respectively.

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A fluorescent/photoacoustic probe for imaging of hydroxyl radical and high-throughput screening of natural products to attenuate acute kidney injury

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