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. 2024 Dec 11;17(1):211–221. doi: 10.1021/acsami.4c13358

Ultrasound-Triggered Nanogel Boosts Chemotherapy and Immunomodulation in Colorectal Cancer

Rui Cui †,‡, Jingwen Zhou †,‡, Wei Yang §, Yao Chen †,‡, Limei Chen †,‡, Lei Tan ∥,*, Feng Zhang †,‡,*, Guangjian Liu †,‡,*, Jie Yu ⊥,*
PMCID: PMC11783521  PMID: 39660733

Abstract

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Chemotherapy is the primary therapy for colorectal cancer. However, its efficacy has been limited by chemoresistance, which is mainly caused by inadequate intratumoral drug accumulation and immunosuppressive microenvironments. To address these limitations, we developed a low-intensity ultrasound (LIU)-controlled and charge-reversible nanogel (R-NG), utilizing conjugated chitosan-polypyrrole polymers linked via thioketal bonds, with TiO2 absorbed onto its surface. Following the loading of oxaliplatin, the Oxa-R-NGs were ultimately synthesized. In the acidic tumor environment, the protonation of the pyrrole ring triggered the conversion of Oxa-R-NG into a positively charged form, thereby enhancing tumor penetration and cellular internalization. Based on the charge conversion, intratumoral accumulating Oxa-R-NG was triggered by LIU to continuously generate reactive oxygen species (ROS), which not only disrupted thioketal bonds to liberate oxaliplatin but also regulated tumor-associated macrophage polarization. Consequently, Oxa-R-NG boosted the chemotherapy for colorectal cancer by improving intratumoral drug accumulation and reversing the local immunosuppressive microenvironment synergistically.

Keywords: nanogel, drug release, charge reverse, low-intensity ultrasound, chemotherapy

Introduction

Colorectal cancer (CRC) ranks as the third most prevalent malignancy worldwide and is the second leading cause of cancer-related mortality.1 In the standard procedures for CRC, systemic chemotherapy plays a pivotal role.2 However, the effectiveness of chemotherapy is significantly compromised by the emergence of chemoresistance. In particular, despite the extensive utilization of oxaliplatin (Oxa) as a first-line chemotherapy agent, its monotherapy response rate remains merely at 24%,3 and prolonged treatment can lead to drug resistance in nearly all patients.4 To date, the etiology of chemoresistance has been mainly attributed to inadequate drug accumulation at tumor sites.5 Furthermore, increasing evidence has supported the involvement of the tumor immunosuppressive microenvironment in chemoresistance.6,7 Specifically, the diminished differentiation of M1-like tumor-associated macrophages has been implicated in Oxa chemoresistance.7 As a result, these factors contribute to a heightened rate of recurrence and a deterioration in overall survival.8−10 Therefore, it is imperative to promptly resolve the clinical bottleneck issues.

Several studies have suggested the use of cancer nanomedicines to address the problem of nonspecific drug accumulation.11−13 However, most types of cancer nanomedicine do not exhibit localized action in clinical tests.14 The use of an external physical stimuli-responsive nanocarrier is considered a promising strategy to overcome this challenge, with stimuli that include temperature, electromagnetism, light, and ultrasound (US).15 Among these, US can penetrate deep tissues while offering spatiotemporal controllability with thermal effects, mechanical effects, or both.16,17 US-responsive nanomedicine is designed to respond to either low- or high-intensity US via thermal or nonthermal effects.18 Low-intensity US (LIU) is an ultrasound that delivers at low intensity and outputs. It generally applies a frequency of 1–3 MHz and an ultrasound intensity of no more than 1 W/cm2.19 Compared with high-intensity focused US (HIFU), LIU exhibits reduced possibilities for biological damage; however, its inferior thermal and mechanical effects render it less potent in facilitating drug release.20,21 Nevertheless, it is worth noting that LIU can effectively activate sonosensitizers to consistently generate reactive oxygen species (ROS).22,23 ROS-responsive nanomedicine, involving selective drug release at specific targets with elevated ROS levels, is an effective drug delivery approach.24 Therefore, we chose to incorporate this functionality into the development of an LIU-controlled drug delivery system (DDS) for the specific release of chemotherapy agents.

Moreover, LIU-controlled DDSs can increase intratumoral ROS levels, suggesting its potential for achieving a similar effect as chemodynamic therapy (CDT).25 The underlying mechanism of CDT involves the generation of •OH within the tumor site through Fenton or Fenton-like reactions. Recently, CDT has garnered considerable attention owing to its distinct advantages, especially its ability to modulate the hypoxic and immunosuppressive tumor microenvironments.25,26 Consequently, this LIU-controlled DDS has a notable potential for remodeling the immunosuppressive tumor microenvironment.

Among the various types of DDS, nanogels (NGs) can maintain their highly hydrated state and dynamic shrinking–swelling properties under different environmental conditions.27 As such, they are suitable for encapsulating hydrophobic chemotherapy agents while concurrently fulfilling multiple functionalities, including prolonged blood circulation time, targeting, stimuli responsiveness, cargo hosting, and cargo delivery.27−29 Consequently, we opted for NGs as the carrier in our LIU-controlled DDS. To minimize drug loss during circulation in the bloodstream and enhance intratumoral drug accumulation,30 we have additionally incorporated charge-reversal characteristics into the nanogel structure. Hence, a novel formulation of chitosan-polypyrrole nanogels (CS-PPy-TK NGs) polymerized by thioketal bonds (TK) with an absorbed TiO2 sonosensitizer was developed. This structural design causes two distinct functions in these NGs. The first function is to facilitate a cascade reaction initiated by the US for the cleavage of the TK, resulting in the disintegration of the NG structure. Second, subsequent treatment of the NGs with an alkaline solution was enabled, generating charge-reversible NGs (R-NGs). Finally, Oxa was physically entrapped in the R-NGs to obtain the Oxa-R-NGs. The functionalized Oxa-R-NGs remained negatively charged in the bloodstream for a prolonged circulation time and were converted to a positive charge within the acidic tumor microenvironment, improving tumor penetration and cellular internalization. Following successful tumor-specific accumulation, under US radiation, the local ROS concentration increased and split the structure of the Oxa-R-NGs to release Oxa. To the best of our knowledge, this is the first report on the design of LIU-responsive NGs based on the chemical cascade effect to obtain specific drug releases.

Results and Discussion

Nanogel Synthesis Process

Oxa-R-NGs were designed to undergo charge reversal in a weakly acidic tumor microenvironment and achieve LIU-responsive ROS production and drug release. To construct ultrafast pH-triggered charge-reversal NGs, we first synthesized chitosan-polypyrrole (CS-PPy) polymers through chemical oxidation polymerization. These were then cross-linked with TK in an aqueous solution and activated by 1-ethyl-3-(−3-(dimethylamino)propyl) carbodiimide hydrochloride (EDC) and N-hydroxysuccinimide to create CS-PPy-TK NGs (P-NGs) (Figure 1A). In the Fourier transform infrared spectroscopy (FTIR) spectra, peaks at approximately 1535, 1170, and 940 cm–1 are attributed to the C=C, C=N, and C–H bonds of the pyrrole (Py) ring, respectively. These findings confirm the successful grafting of Py onto chitosan (CS) to synthesize CS-PPy (Figure 1B, red). A new peak at 1690 cm–1 for P-NGs was attributed to the amide bond (−CO–NH−); this indicated successful cross-linking between the amine groups of CS and the carboxyl groups of TK (Figure 1B, blue). After treatment with a concentration of NaOH solution, the P-NGs could acquire a negative charge because of the selective adsorption of OH– on the Py rings.31 The CS-PPy-TK–OH NGs (R-NGs) exhibited a clear charge conversion from pH 7.4 to 6.5 due to the protonation of the Py ring. This led to the formation of self-adaptive, intelligent CS-PPy-TK–OH NGs (R-NGs) (Figure S1). In thermogravimetric analysis (Figure S2A), the curves of R-NGs remained above those of P-NGs, indicating the successful binding of hydroxide ions. In scanning electron microscopy (SEM) images, the prepared R-NGs in a dehydrated state exhibited a relatively uniform morphology, and the size distribution of most particles was around 200 nm (Figure S2B,C). The biodegradability of the R-NGs was assessed using pancreatin; after 24 h, SEM and dynamic light scattering (DLS) images revealed R-NGs of various sizes (Figure S2B,C), indicating gradual degradation in the physiological environment.

Figure 1.

Figure 1

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Synthesis, and characterization of Oxa-R-NGs. (A) Schematic illustration of the synthetic process involved in the production of CS-PPy-TK NGs (P-NGs). (B) FTIR spectra of CS-PPy polymers and P-NGs. (C) ζ-Potentials of the P-NGs and R-NGs at pH values of 7.4 and 6.5. (D) Protein resistance assay of P-NGs and R-NGs at both pH 7.4 and 6.5. (E) Ultrasound (US)-responsive properties and mechanism of R-NGs. US irradiation can make TiO2 to generate ROS, leading to the cleavage and destruction of the structure of R-NGs. (F) SEM image of R-NGs before and after US irradiation. (G) Cumulative release of Oxa in Oxa-R-NGs at pH 7.4 and 6.5 with or without US.

Characterization of pH-Triggered Charge Reversal

The pH gradient between physiological and tumor microenvironments often serves as a triggering signal for charge reversal. To verify the pH-triggered charge-reversal capabilities of R-NGs, the surface potentials of both P-NGs and R-NGs were measured at pH levels of 7.4 and 6.5, simulating physiological conditions and a tumor microenvironment, respectively. When the pH of the solution was changed from 7.4 to 6.5, the charge of the P-NGs remained positive, whereas that of the R-NGs changed from −9.5 to +10.4 mV (Figure 1C). These results indicate the charge conversion abilities of R-NGs under different pH conditions. Besides, in the presence of bovine serum albumin at the pH levels of 7.4 and 6.5, the P-NGs exhibited a significantly higher protein adsorption rate than the R-NGs under both pH conditions (p < 0.001) (Figure 1D). This suggests that R-NGs resist protein absorption in physiological environments, which enhances their stability during blood circulation.

Capability of US-Responsive Drug Release

Site-specific drug release was accomplished through the incorporation of a reactive oxygen species (ROS)-cleavable thioacetal ketone (TK) group and the sonosensitizer titanium dioxide (TiO2) in the synthesis of Oxa-R-NGs. This configuration facilitated the generation of ROS upon low-intensity ultrasound (LIU) irradiation, thereby initiating an ROS-induced cascade reaction that resulted in the release of Oxa (Figure 1E). To quantify the ROS production in R-NGs after different durations of US exposure, fluorescence spectrophotometry was employed. The results revealed that the ROS concentration increased with the US irradiation time (Figure S2D). To assess the changes in the size and morphology of the R-NGs following US irradiation, SEM and DLS were used. Postirradiation, the R-NGs exhibited an uneven surface morphology and a heterogeneous size distribution, ranging from 7.5 to 190.1 nm (Figures 1F and S2E). These observations indicated the US responsiveness of the R-NGs. To further explore US-responsive drug release, Oxa-R-NGs were obtained by physically mixing Oxa and R-NGs at 25 °C. Then, the concentration of Oxa was measured through ultraviolet–visible (UV–vis) spectroscopy (Figure S3). As shown in Figure 1G, in a neutral environment with a pH of 7.4, Oxa was released slowly with a 24-h cumulative release rate of only 21.5%. However, in an acidic environment with a pH of 6.5, the Oxa release rate was higher, reaching a 24-h cumulative rate of 43.4% (p < 0.001). Remarkably, under US irradiation at a pH of 6.5, the 24-h cumulative Oxa release rate increased to 77.2% (p < 0.001).

These results suggest that Oxa-R-NGs are effective at unloading Oxa in acidic tumor microenvironments and that US irradiation can nearly double the rate of drug release. Previously developed US-responsive DDSs mainly utilized the thermal and cavitation effects of US to achieve drug release.18 However, the utilization of the thermal effect requires high-intensity focused US energy, and the cavitation effect relies on the precise regulation of parameters, such as frequency and intensity. Compared with the direct use of thermal and cavitation effects from the US, drug release induced by ROS produced by LIU has higher accuracy and controllability.

Antitumor Effect of Oxa-R-NGs In Vitro

In vitro, the IC50 for the Oxa-R-NGs-combined-with-US group against CT26luc cells was found to be 6.5 μg/mL according to the Oxa concentration (Figure S4). To check whether combining Oxa-R-NGs with US irradiation could enhance the efficacy of chemotherapy at the cellular level, both cell viability and apoptosis rates were evaluated across different treatment groups. As indicated in Figure 2A, the Oxa-R-NGs group had a significantly higher cell inhibition than the Oxa group (57.0 vs 46.5%, p = 0.0343). Moreover, the percentage of cell inhibition for the Oxa-R-NGs combining US group exceeded that for the Oxa-R-NGs group (69.8 vs 57.0%, p = 0.0061). The apoptosis behavior was further assessed using flow cytometry (Figure 2B,C). The results indicated that the proportion of apoptotic CT26luc cells reached 32.7% for the group treated with Oxa-R-NGs combined with US, which significantly exceeded those of all of the other groups (Figure 2B,C). These results confirm that Oxa-R-NGs can enhance the inhibitory effects of Oxa on CT26 cells and that US irradiation can augment this inhibitory action.

Figure 2.

Figure 2

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Antitumor effect of Oxa-R-NGs in vitro. (A) Cell viability of CT26 cells in different treatment groups. (B, C) Flow cytometry graph of the apoptosis of CT26 cells in the various treatment groups and statistical analysis of the flow cytometric data of apoptosis of CT26 cells in different treatment groups. (D, E) Immunofluorescence staining images of Arg-1 (green) and iNOS (green) of RAW264.7 cells and statistical analysis of the Arg-1 and iNOS staining results of RAW264.7 cells in different groups. Data are expressed as mean ± SD (n = 3); *p < 0.05, **p < 0.01, ***p < 0.001, and ****p < 0.0001.

Immunosuppressive Microenvironment Modulation Effect of Oxa-R-NGs In Vitro

Given that the treatment process entails the generation of ROS at the tumor site, we hypothesized that the utilization of Oxa-R-NGs combined with US irradiation could yield outcomes comparable to those of chemodynamic therapy (CDT)-a known method for modulating the immunosuppressive tumor microenvironment. To validate this hypothesis, RAW264.7 cells were subjected to incubation with phosphate-buffered saline, Oxa-R-NGs, R-NGs combined with US, or Oxa-R-NGs combined with US, followed by immunofluorescence staining of iNOS and Arg1. Quantitative analysis indicated that the combination of Oxa-R-NGs and US resulted in a significantly higher M1/M2 ratio compared with those of the Oxa-R-NGs group and the other experimental groups (Figure 2D,E). Therefore, the combination of Oxa-R-NGs and US induced notable polarization of M2 macrophages toward an M1 phenotype, contributing to the reprogramming of an immunosuppressive tumor microenvironment, which is a main factor for chemoresistance.

Antitumor and Immunomodulation Effect of Oxa-R-NGs In Vivo

In vivo, the biosafety of intravenously administered Oxa-R-NGs at an Oxa concentration of 1.5 mg/kg was first confirmed (Figure S5A,B). Then, a subcutaneous xenograft tumor model was established using nude BALB/c mice to verify the therapeutic effect. The mice were randomly divided into four treatment groups: a saline control group, an Oxa-R-NGs group, an Oxa-P-NGs combining US group (without charge-reversal capabilities), and an Oxa-R-NGs combining US group. The treatment schedules are presented in Figure S6A. Tumor progression across these groups was monitored pre- and post-treatment using an in vivo imaging system (IVIS) kinetic optical system (Figure 3A). The combined treatment of the Oxa-R-NGs and US groups led to significant suppression of tumor growth relative to both the control group and the Oxa-R-NGs group (Figure 3B). The tumor dimensions were measured using a vernier caliper every 3 d, and the findings indicated that the tumors in the Oxa-R-NGs-combined-with-US group grew more slowly than those in the other groups (Figure 3C, all p < 0.001). Following a 14-d treatment period, the therapy was terminated, and the tumors were excised for analysis. As shown in Figure 3D,E, the final tumor volumes in the Oxa-R-NGs (US+) group were significantly smaller than those in the other groups (Oxa-R-NGs (US+)- 828.4 ± 11.3 mm3, Oxa-P-NGs (US+)- 1284.5 ± 117.6 mm3, Oxa-R-NGs- 1226.9 ± 71.7 mm3, control- 1748.9 ± 50.2 mm3, all p < 0.001), confirming that the combined treatment of the Oxa-R-NGs and US effectively and consistently inhibited tumor growth in vivo. Furthermore, immunofluorescence staining indicated that Oxa-R-NGs significantly induced apoptosis relative to that in the control group (Figure 3F). Moreover, this apoptotic effect was substantially enhanced when Oxa-R-NGs were treated with US irradiation (p < 0.001) (Figure 3G), which was attributed to the US-responsive release features of Oxa-R-NGs.

Figure 3.

Figure 3

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Treatment effects of Oxa-R-NGs on subcutaneous xenograft CRC mice. (A) IVIS images of subcutaneous CT26 tumors on Day 7 and Day 21 in each treatment group. (B) Bioluminescence intensity of CT26 tumors in different treatment groups. (C) Time-dependent variation in the subcutaneous tumor volume of mice that underwent different treatments. (D) Digital photos of tumors on Day 21 in each treatment group. (E) Terminal volumes of subcutaneous tumors on Day 21 in various treatment groups. (F) Terminal deoxynucleotidyl transferase dUTP nick end labeling (TUNEL) staining (green) of tumor tissues of mice treated in different groups on Day 21 (The nuclei were stained using 4′,6-diamidino-2-phenylindole, blue). (G) Statistical analysis of TUNEL staining results in CT26 tumor tissues. Data are expressed as mean ± SD (n = 3); *p < 0.05, **p < 0.01, ***p < 0.001, and ****p < 0.0001.

Regarding the safety profile of this treatment, the results from body weight monitoring, hematoxylin and eosin staining of major organs, as well as whole blood components analysis indicate that the combined application of Oxa-R-NGs and US irradiation exhibits no systemic toxicity, even after repeated treatments (Figure S6B–D).

To further assess the tumor microenvironment, the microvascular density (MVD) was evaluated through CD31 endothelial cell marker staining (Figure 4A). Quantitative analysis revealed that the MVD was significantly reduced for the group treated with Oxa-R-NGs combined with US, compared to the other groups (Figure 4B). Besides, Oxa can induce immunogenetic cell death (ICD)—a process that activates damage-associated molecular patterns (DAMPs) such as calreticulin (CRT) and high-mobility group protein B1 (HMGB1) translocation. Therefore, we investigated the potential role of Oxa-R-NGs in inducing ICD. Our findings indicated significant increases in CRT surface exposure (Figure 4A,C) and HMGB1 release (Figure 4A,D) for the Oxa-R-NGs-combined-with-US group compared with those of the other treatment groups. This result suggested that the synergistic effect of Oxa-R-NGs and US induced a more robust ICD response than the use of Oxa or Oxa-R-NGs only. The DAMPs release due to ICD and their subsequent recognition by pattern-recognition receptors lead to the generation of specific CD8+ T lymphocytes that effectively eliminate tumor cells. Therefore, the utilization of Oxa-R-NGs augmented the antitumor immune response of Oxa.

Figure 4.

Figure 4

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Immunomodulation effect of Oxa-R-NGs in subcutaneous xenograft CRC mice. (A) Immunohistochemistry staining of HMGB1, CRT, and CD31 (positive areas, brown) in CT26 tumor tissues after corresponding treatments in different groups. (B–D) Statistical analysis of the CD31, CRT, and HMGB1 staining results in CT26 tumor tissues. (E, F) Immunofluorescence staining images and statistical analysis of Arg-1 (green) and iNOS (green) of tumor tissues in different groups. Data are expressed as mean ± SD (n = 3); *p < 0.05, **p < 0.01, ***p < 0.001, and ****p < 0.0001.

To validate the modulation effect of Oxa-R-NGs on immunosuppressive microenvironments in vivo, we performed immunofluorescence staining on iNOS (M1 marker) and Arg1 (M2 marker) within the tumor tissue sections of mice from various treatment groups (Figure 4E,F). The findings revealed a significantly increased M1/M2 ratio for the Oxa-R-NGs-combined-with-US group compared to the other groups (Figure 4E,F). Therefore, the capability of Oxa-R-NGs to reprogram the immunosuppressive tumor microenvironment was demonstrated in vivo. The Oxa-R-NGs were suitable for application in colorectal cancer (CRC) with metastases, particularly in cases involving liver metastases. Owing to the high rates of metastasis and mortality associated with malignant tumors, colorectal liver metastases (CRLMs) are primarily treated through a combination of systemic chemotherapy and localized therapy. However, the effectiveness of systemic chemotherapy varies between the primary tumors and intrahepatic metastases within the same patient. When liver metastases are unresectable or primary tumors persist after chemotherapy, the planned surgical interventions are severely impacted, deteriorating the treatment outcomes. Oxa-R-NGs can achieve high drug concentrations directly at the specific tumor location, whereas US irradiation can target sites that are relatively unresponsive to chemotherapy, enhancing the overall efficacy of the systemic treatment. Therefore, the Oxa-R-NGs present new avenues for improving CRC chemotherapy. Further, the simple component of Oxa-R-NGs and convenient utilization of LIU facilitate a considerable clinical transformation potential of the proposed method.

Conclusions

In summary, we have developed a method to prepare a low-intensity ultrasound-triggered, charge-reversible nanogel to enhance the efficacy of chemotherapy in colorectal cancer while simultaneously boosting antitumor immunity. This novel drug-delivery system achieves controlled local administration without the need for direct local injections. For patients with multiple and deep-seated metastatic lesions, a single intravenous administration combined with low-intensity ultrasound irradiation enabled targeted treatment at multiple sites. This approach holds particular promise as a potential therapeutic strategy for colorectal cancer, which frequently originates multiple metastases in the liver and abdominal lymph nodes. Furthermore, despite carrying only oxaliplatin as a single drug, Oxa-R-NGs improve the chemotherapeutic efficacy and activate antitumor immunity, offering a new avenue for enhancing the effectiveness of colorectal cancer chemotherapy. However, this study has several limitations. Recent studies have confirmed that ROS, when used alongside chemotherapeutic drugs, can activate antitumor T cell immunity.32 Therefore, the effects of ROS and oxaliplatin within the Oxa-R-NGs formulation on T-cell-mediated antitumor immunity warrant further investigation. Ongoing advancements in this nanogel-based technology will likely facilitate its application in clinical settings, ultimately improving the efficacy of chemotherapy and promoting immune responses in solid tumors.

Methods

Materials

CS, Py, ammonium persulfate (APS), EDC, NHS, cyclohexane, Span-80, hydrochloric acid (HCl), acetic acid, absolute ethanol, NaOH, TiO2, and BSA were purchased from Sigma–Aldrich Co. (St. Louis, MO). TK-COOH was purchased from Aladdin Bio-Chem Technology Co. Ltd. (Shanghai, China). Oxa was purchased from Sa En Chemical Technology Co. Ltd. (Shanghai, China). d-Luciferin potassium salt, 4% formaldehyde; 4′,6-diamidino-2phenylindole (DAPI); Cell Counting Kit-8 (CCK-8); a Hematoxylin and Eosin Stain Kit; an Annexin V–PI/FITC Apoptosis Detection kit; and a terminal deoxynucleotidyl transferase-mediated dUTP nick end labeling (TUNEL) kit were purchased from Beyotime (Shanghai, China).

Nanogel Synthesis

To create the NGs, we initially synthesized a CS-PPy polymer, following the methodology used by Li et al.30 To prepare the CS-PPy-TK nanogels (P-NGs), we mixed CS-PPy polymers (10 mg/mL, 1 mL in 1 M HCl), EDC (4.8 mg), NHS, (1.34 mg), TK-COOH (6.3 mg), and Span-80 (25.8 mg/mL, in 10 mL of cyclohexane). The mixture was ultrasonicated for 10 min using a Misonix Sonicator (SCIENTZ IID, Scientz Biotechnology, Ningbo, China) with a 50% duty cycle and 40% output under ice-cooled conditions. This was followed by stirring at room temperature overnight.

To synthesize the charge-reversible and US-responsive NGs (R-NGs), we first purified the prepared P-NGs through centrifugation at 10,000 rpm for 10 min. The precipitate was redispersed in 10 mL of water and dialyzed using a bag with a molecular weight cutoff (MWCO) of 12,000–14,000 against water (2 L, replaced 6 times) for 3 d. The P-NGs (1 mL) were then treated with a 4 mg/mL concentration of NaOH for 24 h. Subsequently, TiO2 nanoparticles were added to the solution at a concentration of 1 mg/mL. The mixture was sonicated with an ultrasonic processor for 10 min and then stirred for another 10 min. For the synthesis of Oxa-loaded NGs (Oxa-R-NGs), R-NGs or P-NGs (1 mg) and Oxa (1 mg) were dissolved in 1 mL of deionized water and stirred in the dark for 24 h. All free Oxa was removed via centrifugation at 12,000 rpm for 10 min to obtain the final Oxa-R-NGs or Oxa-P-NGs.

Characterization of Nanogels

FTIR spectra of CS, CS-PPy, and CS-PPy-TK were collected using an FTIR spectrophotometer (FTIR-650, Guangdong Technology Co. Ltd., Tianjin, China). The morphological structures of the NGs were examined using a scanning electron microscope (JSM-6390LV, JEOL, Japan), before and after ultrasonic treatment. The size distributions were measured using a DLS instrument (Malvern Instruments, Malvern, U.K.) before and after ultrasonic and pancreatin treatments, the latter to evaluate biodegradability. Briefly, all of the test samples were prepared by ultrasonically dispersing 3 mg of R-NGs in 1 mL of deionized water. To test the protein resistance of the NGs, prepared P-NGs and R-NGs were combined with BSA (2 mg/mL) in 1 mL of PBS at pH levels of 7.4 and 6.5. The mixtures were then incubated at 37 °C for 2 h. Subsequently, the mixtures were centrifuged at 3000 rpm for 5 min to obtain the supernatant. The concentration of adsorbed protein was determined using a BCA protein assay kit as per the manufacturer’s guidelines, and absorbance was measured at 570 nm using a microplate reader. To evaluate the charge characteristics of the P-NGs and R-NGs, they were incubated in PBS at pH 7.4 or 6.5. After 10 s of incubation, their zeta potentials were measured by using a Zetasizer Nano ZS system (Malvern Instruments, Worcestershire, U.K.). Thermogravimetric analysis (TGA) was conducted by weighing 3 mg of prepared P-NGs and R-NGs in an aluminum foil crucible. The samples were then heated from room temperature to 700 °C at a rate of 20 °C/min under a nitrogen atmosphere. Changes in mass relative to temperature were recorded using a thermogravimetric analyzer (Q2000; TA Instruments, New Castle, DE).

In Vitro ROS Generation

The ROS levels were measured by using fluorescence spectrophotometry with 2′,7′-DCFH-DA as the probe. First, 0.01 mL of DCFH-DA in dimethyl sulfoxide was mixed with 0.04 mL of 0.01 M NaOH, and the mixture was incubated in the dark for 30 min to obtain DCFH. The reaction was terminated by adding 5 mL of PBS (25 mM). To compare ROS levels generated by R-NGs under different durations of US exposure, the R-NG solution was mixed with DCFH (10 μM) in a 1:1 ratio. The mixture was then sonicated for various time intervals—0, 1, 3, and 5 min—with the following settings: an intensity of 1 W/cm2, a duty cycle of 50%, and a frequency of 1 MHz. The mixed solution was centrifuged at 12,000 rpm for 10 min, and the supernatants were obtained for ROS quantification using a Mithras LB 943 multimode reader (Berthold Technologies, Bad Wildbad, Germany).

Drug Encapsulation and US-Responsive Release

We prepared 10 mL of an Oxa solution at a concentration of 1 mg/mL and diluted it to create a series of standard solutions with concentrations of 0.016, 0.032, 0.063, 0.125, 0.25, and 0.5 mg/mL. These were used to establish a standard Oxa absorbance calibration curve. The concentrations of free Oxa in the collected supernatants were measured using UV–vis spectroscopy at 250 nm according to the standard curve. The LE was calculated using the following equation

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The in vitro Oxa release kinetics of Oxa-R-NGs for the following groups were tested: pH 7.4, 6.5, and 6.5 (US+). One mg of R-NGs was dissolved in 1 mL PBS at a pH of 7.4 or 6.5. Subsequently, the supernatant was collected at different times (0, 0.5, 1, 2, 4, 6, 10, 14, 18, and 24 h) after centrifugation (12,000 rpm, 5 min) and was analyzed via UV–vis spectroscopy at 250 nm based on the standard curve. For the pH 6.5 (US+) group, US treatment was applied (1 W/cm2, 50% duty cycle, 1 MHz) for 5 min after 4 h. Then, the solution was centrifuged (12,000 rpm, 5 min), and the supernatant was collected and measured as described above.

In Vitro Anticancer Efficacy

A total of 1 × 104 CT26luc cells/well were cultured in a 96-well culture plate and incubated with 100 μL of fresh medium for 24 h. The medium in each well was then replaced with fresh medium containing Oxa-R-NGs at different final Oxa concentrations (0.5, 1.0, 2.5, 5, 10.0, and 20.0 μg/mL). After 48 h of incubation, the cell viability was measured by using a cell counting kit-8 (CCK-8) assay. Then, the half-maximal inhibitory concentration (IC50) of Oxa was calculated. In a separate experiment, 1 × 104 CT26luc cells/well were cultured in a 96-well plate with 100 μL of fresh medium for 24 h. The cells were then treated for an additional 48 h with various solutions: PBS, R-NGs, free Oxa, Oxa-R-NGs, or Oxa-R-NGs with US exposure (US+). The Oxa dosages in the free Oxa, Oxa-R-NGs, and Oxa-R-NGs (US+) groups were based on the previously calculated IC50 values. A CCK-8 assay was employed to detect the inhibitory effects on cell proliferation for each treatment group. For apoptosis rate analysis, 5 × 105 cells were cultured in a 6-well plate and incubated for 24 h. Cells were then treated for an additional 48 h with PBS, R-NGs, free Oxa, Oxa-R-NGs, or Oxa-R-NGs (US+). Apoptosis rates were determined using an Annexin V–PI/FITC apoptosis detection kit and flow cytometry.

In Vivo Biosafety

Female BALB/c mice, aged 6 weeks and weighing 15–18 g, were purchased from Guangdong Ruige Biological Technology Co. Ltd. and housed in accordance with the animal research guidelines set by the National Ministry of Health. Twelve of these mice were randomly divided into four distinct groups. The mice in each group were intravenously administered one of the following treatments via the tail vein: saline, R-NGs, Oxa-R-NGs at a dose of 1.5 mg/kg, or Oxa-R-NGs at a dose of 3.0 mg/kg (n = 3). The concentration of R-NGs matched that of the Oxa-R-NG group dosed at 1.5 mg/kg Oxa. One week postinjection, the major organs were harvested from the mice for histological analysis, which involved staining with H&E. Additionally, blood samples were collected through ocular extraction for standard biochemical examination.

In Vivo Anticancer Efficacy

Four-week-old female nude BALB/c mice weighing 13–15 g were purchased from Guandong Ruige Biological Technology Co., Ltd. and housed in accordance with the National Ministry of Health’s animal research policies. The study protocols for in vivo anticancer and biocompatibility experiments were approved by the Ethical Committee of the sixth affiliated hospital of Sun Yat-Sen University. To establish a subcutaneous xenograft model, 5 × 106 cells suspended in 200 μL of PBS were injected into the right flanks of the mice. After 7 days, the tumors had grown to approximately 100 mm3 in volume. The mice were then divided randomly into four groups (n = 3 per group): (a) saline, (b) Oxa-R-NGs, (c) Oxa-P-NGs (US+), and (d) Oxa-R-NGs (US+). Treatment formulations of 100 μL were intravenously administered through tail vein injection for four cycles on days 7, 14, 17, and 20, according to the respective group. The dose of Oxa for groups (b to d) was set as 1.5 mg/kg. Following injections, groups (c and d) were subjected to ultrasonic irradiation treatment (1 W/cm2, 1 MHz, 50% duty cycles, 5 min). The tumor sizes were measured every 3 days and were calculated using the following formula: 0.52 × L × W2 (L, tumor length; W, tumor width). The weights of the mice were recorded every 2 d. Additionally, fluorescence imaging was performed, and the tumor fluorescence intensity was measured using an in vivo bioluminescence imaging system (AniView100 system, BLT, Guangzhou, China) at 7 and 21 d. D-luciferin potassium salt dissolved in PBS (15 mg/mL) was injected intraperitoneally at a dose of 150 μL for each mouse. Ten minutes after injection, the mice were anesthetized and photographed by the imaging system. Bioluminescence images were then analyzed using AniView 100 Living Imaging software (BLT, Guangzhou, China), and the average radiance in the regions of interest was quantified (photons s–1 cm–2 sr–1). All mice were sacrificed on the 21st day. Major organs and blood samples were collected for histological analysis and routine blood biochemical examinations, as previously mentioned. The tumors were stripped from mice and subjected to immunofluorescence staining and immunohistochemistry (IHC) analysis.

Immunohistochemistry Assay (IHC)

Excised tumors were fixed with 4% formaldehyde, embedded in paraffin blocks, and sectioned. Immunostaining was performed on paraffin sections following the process including deparaffinization, antigen-repaired for 30 min, 3% H2O2 solution block endogenous peroxidase activity, 3% BSA blocked for 30 min, incubated with primary antibodies against CRT (Abcam, Cambridge, U.K.), HMGB1 (Novus Biologicals Littleton, CO), Ki67 (A20018, Abclonal, China), and CD31 (ab9598, Abcam, U.K.), followed by incubation with corresponding secondary antibodies as per the manufacturer’s instructions. Staining was exhibited with DAB solution, and Harris’ hematoxylin was used to restain the nucleus.

Immunofluorescence Staining

Tissue immunofluorescence assay was performed on paraffin section and following the process, including deparaffinization, incubate with proteinase K working solution at 37 °C for 25 min, incubate with membrane breaking working solution at 37 °C for 20 min, add fluorophore-tagged antibodies, including iNOS (18985–1-AP, Proteintech, China) and Arg-1 (66129–1-Ig, Proteintech, China). TUNEL assay was performed using the DeadEnd Fluorometric TUNEL System (Promega, WI). Then, the slides were stained with DAPI and observed and analyzed using a fluorescence microscope.

For cell immunofluorescence staining, RAW264.7 cell lines (M0 macrophages) were used in this experiment and cultured in basic DMEM containing 10% FBS. A total of 5 × 105 cells/well were seeded in a 24-well plate and incubated with 1 mL fresh medium for 24 h. Then, cells were treated for an additional 48 h with PBS, R-NGs, R-NGs with US exposure (US+), Oxa-R-NGs, or Oxa-R-NGs with US exposure (US+), respectively. Then, the cells were fixed, permeabilized, and blocked and then incubated with primary antibodies iNOS (18985–1-AP, Proteintech, China) and Arg-1(66129–1-Ig, Proteintech, China) at 4 °C overnight. After washing with PBS 3 times, cells were incubated with the corresponding secondary antibody for 1h at 37 °C in the dark. Then, the cells were stained with DAPI for 5 min, observed, and analyzed using a fluorescence microscope.

Statistics

The results are presented as the mean ± the standard deviation (SD). Differences between the two groups were assessed using an unpaired, two-tailed t test. For comparisons among three or more groups, a two-way analysis of variance (ANOVA) with Bonferroni’s postnote model was employed. p < 0.05 was considered statistically significant. Statistical analyses were performed using GraphPad Prism software version 9.5.0.

Acknowledgments

Guang Zhou Basic and Applied Basic Research Program (202102020992), Guang Dong Basic and Applied Basic Research Foundation (2021A1515110107), National Science Fund for Distinguished Young Scholars (Grant No. 82325027), and Special Research Fund for Military Health Care (23BJZ31).

Supporting Information Available

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acsami.4c13358.

  • Schematic design of R-NGs; characterization of Oxa-R-NGs; UV–vis spectroscopy of oxaliplatin solution; cell viability of CT26 cells treated with Oxa-R-NGs at varying concentrations of Oxa; and biosafety of R-NGs and Oxa-R-NGs; treatment schedule and biosafety assessment in different treatment groups on subcutaneous xenograft CRC mice (PDF)

Author Contributions

# Conceptualization, R.C., with input from J.Y. and G.J.L.; methodology, R.C., J.W.Z., and W.Y., with input from J.Y.; investigation/experiments, J.W.Z., Y.C., L.T., L.M.C., and W.Y.; formal analysis, J.W.Z. and W.Y. with input from J.Y.; manuscript writing, R.C. and J.W.Z.; funding acquisition, R.C.; supervision, Y.C., L.M.C., and F.Z.; manuscript editing and review, all authors.

The authors declare no competing financial interest.

This paper was published ASAP on December 11, 2024. Additional content was removed from page eight of the document and the corrected version was reposted on December 18, 2024.

Supplementary Material

am4c13358_si_001.pdf (1.4MB, pdf)

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Supplementary Materials

am4c13358_si_001.pdf (1.4MB, pdf)

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