In this work, the effects associated with apoptosis were not exclusive to the chemotherapeutic agent and glutathione; some authors report events in which chitosan has an essential role in apoptosis. In studies carried out on bladder tumor cells, Hasegawa et al. showed that chitosan could induce apoptosis through the activation of caspase-3. However, the mechanism of action through which it carries out this activation is unknown [31]. Lee et al., 2011, showed that a derivative of chitosan, diethylaminoethyl chitosan, could induce apoptosis in Hela cells through the regulation of enzymes (caspase-3, -8, and -9); p53 and BAX expression. Modifying the expression of BCL2 proteins. This would generate a disruption of the mitochondrial membrane and an oxidation–reduction imbalance [32].

Wimardhani et al., 2014, reported that the exposure of Ca9-22 cells with chitosan derivatives induced the appearance of early apoptotic cells, increased caspase-3 activity, and the arrest of G1/S of the cell cycle, suggesting that chitosan could be used as a natural anti-cancer agent [33]. Therefore, the caspase-3 activity increase could be due to the exposure to doxorubicin, GSH of NPs, and chitosan.

#### *3.9. CH-GSH NPs Impair Cell Proliferation by Decreasing Ki67 Levels*

Ki67 levels were measured as a molecular marker of cell proliferation. The Ki67 antigen has a specific expression in the M phase of the cell cycle; it is commonly visualized with the MIB1 antibody [34]. Our data showed that the CH-GSH NPs in both concentrations significantly decreased the percentage of Ki-67 concerning the NT cells in both cell lines (Figure 11). It has been reported that GSH levels can regulate the activity of genes associated with proliferation, differentiation, and apoptosis, and a high level of GSH is essential for normal cellular functions, signal translation, and protection against certain carcinogens [35]. When cells are exposed to both doxorubicin and CH-GSH NPs, the levels remain like those obtained with cells exposed only to doxorubicin. The above suggests that NPs could not affect doxorubicin-induced proliferation but could modify the redox state as observed when ROS levels were estimated.

**Figure 11.** Ki67 levels. The cells were exposed to Dox for 12 h and then 2 h with CH-GSH NPs. (**A**) MCF-7 and (**B**) MDA MB-231 cells. Bars with equal letters indicate no significant differences between the means (Tukey's test, *p* < 0.05).

Furthermore, there is evidence that the decrease in Ki67 levels is related to some chitosan derivatives, which modify various cell-signaling pathways directly or indirectly related to cell proliferation and apoptosis. Chitosan can inhibit pro-inflammatory molecules such as TNF-α, blocking the activation of NFkB and reducing the expression of genes that protect and induce cell proliferation [36]. In addition, it can activate the transcription factor Nrf2 through the PI3/Akt pathway. Rojo de la Vega et al. reported that a decrease in the expression of Nrf2 promotes the activation of CDk2 and CDk4 inhibitor p21 [37]. Another study suggests that some cells treated with chitosan derivatives can increase the expression of TGF-β that activates an intracellular-signaling cascade associated with Smad proteins and thus favors the transcription of genes associated with cell cycle inhibition [38]. MCF-7 cells showed increased sensitivity to NPs' exposure compared to MDA MB-231 cells in the various tests performed. This finding may be due to its metabolism. It has been reported that Dox modifies the metabolism of both cell lines by affecting more the metabolic profile of MDA-MB-231 cells than of MCF-7 cells, showing changes in ketonic bodies, glycolysis, and energetic and lipid metabolism [16,39]. MDA MB-231 cells have a higher consumption of glucose and upregulated redox pathways than MCF-7 [40].

Conversely, the observed effect may be due to, as some authors mentioned, the fact that combined exposure of NPs with various agents enhances cell sensibilization. Zalbielca et al., 2017, and Willmann et al., 2015, showed that combined exposure of NPs of Dox/GSH has higher cytotoxic effects than free Dox in feline fibrosarcoma cell lines. In a study in MCF-7 cells with metallic NPs and radiation, there was a higher effect on the combination exposure, wherein NPs act as nano-sensitizers [6,41]. Uma et al., 2016, suggested that gold NPs act as sensitizing agents in MDA-MB-231 and MCF-7 cells, modifying cell-cycle effects, viability, and DNA damage [42]. Alvandifar et al. used a combined exposition of PLGA and verapamil NPs to improve this chemotherapeutic effectiveness and decrease the dose to have a higher effect [43]. These results suggest that the combined exposure of nanoparticles and doxorubicin may decrease the resistance of MDA MB-231 cells to these types of drugs.

#### **4. Conclusions**

The results obtained in this research suggest that CH-GSH NPs modify Ki67 levels and alter the apoptosis by increasing caspase-3 activity and the cellular redox state, reducing the oxidative stress generated by doxorubicin exposure. It has been documented that GSH may play a dual role in tumor progression or cancer cell death concerning cancer. The above is probably due to the fine biochemical regulation in its synthesis and the relationship of its oxidation–reduction modulating effects, as well as other enzymatic and non-enzymatic mechanisms. The effects of doxorubicin are variable and include the ability to induce resistance to chemotherapy; modify the initiation and progression of cancer; activate cellsignaling pathways related to stressful microenvironments; promote apoptosis in tumor cells; and have the radio-sensitization be induced by this antioxidant. We observed higher sensitivity of MCF-7 cells to CH-GSH NPs than MDA MB-231 cells; this may be due to each cells' genotypic characteristics.

As a GSH delivery entity, CH-NPs attract attention. The results of this work show their capacity to diffuse quickly into cells and exert significant effects to modulate the oxidative stress induced by doxorubicin in breast cancer cells. Considering this, CH-GSH NPs must be studied as a potential designed delivery system that offers a new biomaterial with biomedical opportunities to study the molecular, biochemical, and biological mechanisms related to the cellular redox status. This information will be useful to design better therapies based on antioxidant nanoparticles.

**Author Contributions:** Conceptualization, L.D.L.-B. and P.R.-N.; data curation, L.D.L.-B., R.D.-T. and P.R.-N.; formal analysis, L.D.L.-B., R.D.-T., A.M.S., C.R. and P.R.-N.; funding acquisition, R.D.-T. and P.R.-N.; investigation, L.D.L.-B. and J.R.M.-R.; methodology, L.D.L.-B., J.R.M.-R. and P.R.-N.; project administration, R.D.-T. and P.R.-N.; resources, P.R.-N.; supervision, R.D.-T., A.M.S., C.R. and P.R.-N.; validation, L.D.L.-B. and R.D.-T.; visualization, L.D.L.-B., R.D.-T., A.M.S., C.R. and P.R.-N.; writing—original draft preparation, L.D.L.-B., R.D.-T., J.R.M.-R., A.M.S., C.R. and P.R.-N. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by PAPIIT IN214321 from Dirección General de Asuntos del Personal Académico, Universidad Nacional Autónoma de México.

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** The data presented in this study are available on request for the corresponding author.

**Acknowledgments:** The authors acknowledge the invaluable support of M en C Elizabeth Soria Castro in the TEM analysis. L. D. López-Barrera received a scholarship number 639023 from CONACYT as a doctorate student.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**

## *Article* **Targeting Tumor Cells with Nanoparticles for Enhanced Co-Drug Delivery in Cancer Treatment**

**Wen-Ying Huang 1,†, Chih-Ho Lai 2,†, Shin-Lei Peng 3, Che-Yu Hsu 4, Po-Hung Hsu 5, Pei-Yi Chu 4, Chun-Lung Feng 6 and Yu-Hsin Lin 4,7,8,\***

**Abstract:** Gastric cancer (GC) is a fatal malignant tumor, and effective therapies to attenuate its progression are lacking. Nanoparticle (NP)-based solutions may enable the design of novel treatments to eliminate GC. Refined, receptor-targetable NPs can selectively target cancer cells and improve the cellular uptake of drugs. To overcome the current limitations and enhance the therapeutic effects, epigallocatechin-3-gallate (EGCG) and low-concentration doxorubicin (DX) were encapsulated in fucoidan and D-alpha-tocopherylpoly (ethylene glycol) succinate-conjugated hyaluronic acid-based NPs for targeting P-selectin-and cluster of differentiation (CD)44-expressing gastric tumors. The EGCG/DX-loaded NPs bound to GC cells and released bioactive combination drugs, demonstrating better anti-cancer effects than the EGCG/DX combination solution. In vivo assays in an orthotopic gastric tumor mouse model showed that the EGCG/DX-loaded NPs significantly increased the activity of gastric tumors without inducing organ injury. Overall, our EGCG/DX-NP system exerted a beneficial effect on GC treatment and may facilitate the development of nanomedicine-based combination chemotherapy against GC in the future.

**Keywords:** gastric cancer; nanoparticle; fucoidan; D-alpha-tocopherylpoly (ethylene glycol) succinate; combination chemotherapy

#### **1. Introduction**

The stomach is located in the upper abdomen between the esophagus and small intestine. Its primary function is to store and partially digest food after ingestion [1,2]. Abnormal growth and multiplication of gastric cells can lead to gastric cancer (GC). Most GCs are adenocarcinomas that arise from the mucosal tissue lining inside the stomach. Abnormal cells may penetrate deeper into the stomach wall or spread to nearby organs or tissues [3]. Cell adhesion molecules are the major elements of the metastasis process [4]. In particular, P-selectin is an adhesion molecule that mediates the interaction of platelets

**Citation:** Huang, W.-Y.; Lai, C.-H.; Peng, S.-L.; Hsu, C.-Y.; Hsu, P.-H.; Chu, P.-Y.; Feng, C.-L.; Lin, Y.-H. Targeting Tumor Cells with Nanoparticles for Enhanced Co-Drug Delivery in Cancer Treatment. *Pharmaceutics* **2021**, *13*, 1327. https://doi.org/10.3390/ pharmaceutics13091327

Academic Editors: Marina Santiago Franco and Yu Seok Youn

Received: 27 July 2021 Accepted: 17 August 2021 Published: 25 August 2021

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

and endothelial cells with monocytes, neutrophils, and tumor cells [5].  Cluster of differentiation (CD)44-positive GC cells show self-renewal properties and the ability to produce differentiated offspring, consistent with the cancer stem cell phenotype [8]. CD44-positive GC cells are reported to contribute to increased resistance to radiation- or chemotherapy-induced cancer cell death [9].

Cancer nanotechnology is being vigorously developed for applications in cancer imaging, molecular diagnosis, and targeted therapy [10–12]. Compared with their singleor dual-ligand-targeting nanoparticle (NP) counterparts, non-targeted NPs significantly increase metastasis deposition. Multiligand NPs are obtained by modulating the surfaces of NPs with more than one type of targeting ligand. These facilitate high-precision targeting of different metastatic subpopulations that express different targetable receptors otherwise missed by single-ligand NPs [13,14]. Fucoidan (FD) is the active ingredient of seaweed that induces cell apoptosis, inhibits the proliferation of cancer cells, and is reported to have a strong affinity for P-selectin, which interacts with sulfated oligosaccharides [15–17]. Hyaluronic acid (HA) is a natural linear polysaccharide, composed of repeating disaccharide units of β(1,3)-*N*-acetylglucosamine (GlcNAc) and β(1,4)-glucuronic acid (GlcUA), which forms van der Waals bonds and hydrogen bonds with migrating cells via the CD44 ligand receptors [18,19]. D-alpha-tocopheryl poly(ethylene glycol) (PEG) succinate (TPGS) is a water-soluble derivative of natural vitamin E and prevents P-glycoprotein (P-gp) adenosine triphosphatase (ATPase) from hydrolyzing ATP by blocking the ATP-binding sites, thus inhibiting ATPase activity [20]. Herein, we prepared a TPGS-conjugated HA (TH) copolymer that can simultaneously interact with membrane-bound CD44 and reduce P-gp expression in GC cells.

Doxorubicin (DX), an anthracycline antibiotic, is currently the most effective chemotherapeutic drug used to treat GC, ovarian cancer, and breast cancer. DX acts as a DNA intercalator and topoisomerase II inhibitor. However, it exhibits strong cardiotoxicity [19,21]. The standard dose of 20 mg/m2 is widely used for GC, but an overall response to DX monotherapy is observed in only 17% of GC patients [22–24]. Epigallocatechin-3-gallate (EGCG), a bioactive polyphenol compound in green tea, has antioxidant and anti-inflammatory properties and can be utilized in disease management [25]. EGCG can sensitize the efficacy of DX, enhancing its therapeutic effect in hepatocellular or prostate cancer [26,27]. Combination chemotherapy is a promising method for improving cancer treatment. However, clinical combination therapy is limited by the unique pharmacokinetics of the combined drugs, resulting in uneven drug distribution [28,29]. We established tumor-targeting NPs, combining FD binding to P-selectin-expressing tumors and TH targeting CD44-expressing tumors to reduce P-gp expression. Gelatin is a natural polymer containing residues of glycine, proline, hydroxyproline, and alanine, which are common amino acids. Gelatinpolyphenol interactions primarily occur through hydrogen bonds between hydrophobic amino acids (mainly proline residues) and the phenol ring of polyphenols [30,31]. Thus, in this study, we used PEG-gelatin (PG) to load the anticancer agents EGCG and DX to enhance their loading efficiency and sustained release, thereby attempting to maximize their combined therapeutic effect and minimize systemic toxicity (Figure 1a).

ő

**Figure 1.** (**a**) NPs were prepared by adding EGCG/DX solution into FD/TH/PG solution under gentle stirring at 37 ◦C; the schematic representation of the strategy for using EGCG/DX-loaded FD/TH/PG NPs on carcinoma cells is shown; (**b**) Particle size distributions and zeta potential values of EGCG/DX-loaded FD/TH/PG NPs made with different DX concentrations; (**c**) Fourier transform infrared analyses of FD, TH, PG, EGCG, DX, and FD/TH/PG/EGCG/DX NPs.

#### **2. Materials and Methods**

#### *2.1. Preparation and Characterization of EGCG/DX-Loaded FD/TH/PG NPs*

TH and PG were synthesized according to the protocols suggested by Chen et al. [32]. For TH copolymer preparation, 0.2 mmol dicyclohexylcarbodiimide and 0.2 mmol 4-dimethylaminopyridine were dissolved in 5.0 mL of acetonitrile and added to an aqueous HA solution in deionized (DI) water (20.0 mg/mL in 10.0 mL). The solution was then stirred for 1.0 h to activate the carboxylic groups of HA. Subsequently, 0.1 g of TPGS was dissolved in 2.0 mL of DI water, then slowly added to the above mixed solution and stirred for 24.0 h under a nitrogen atmosphere. The PG copolymer was prepared by dissolving the 2.0 g of gelatin into 20.0 mL of dimethyl sulfoxide (DMSO). The resultant mixture was serviced as solvent for 0.6 g of methoxypolyethylene glycol succinimidyl ester (mPEG-NHS), and then stirred for 4 h. The unconjugated reagents were removed by dialyzing in 5.0 L of DI water, which was replaced five times per day. After the resultant TH and PG were lyophilized in a freeze dryer, Fourier transform infrared (FTIR) spectroscopy was used to detect the presence of purified compound in the sample.

NPs composed of the FD-TH complex with PG containing encapsulated EGCG and DX were prepared as follows. First, the EGCG-loaded FD/TH/PG NPs were formulated by optimizing the relative concentration of FD:TH. A series of FD:TH aqueous solutions (0.00:0.00, 1.20:1.20, 2.40:2.40 and 3.60:3.60 mg/mL in 0.50 mL) were added to aqueous PG solution (15.00 mg/mL in 0.50 mL) and then slowly shaken for 0.5 h at 37 °C. Next, EGCG aqueous solution (5.00 mg/mL in 1.00 mL) was added to 1.0 mL of aqueous FD/TH/PG solutions. The reaction mixture was stirred for 30 min. Then, NPs with different FD:TH:PG:EGCG compositions (0.00:0.00:3.75:2.50, 0.30:0.30:3.75:2.50, 0.60:0.60:3.75:2.50, and 0.90:0.90:3.75:2.50 mg/mL) were completed. After encapsulating DX in these NPs, DX was dissolved in DI water (0.00, 0.10, 0.20, and 0.40 mg/mL in 0.50 mL) and then poured into the EGCG aqueous solution (10.00 mg/mL in 0.50 mL) to

form EGCG/DX aqueous solutions (5.00:0.00, 5.00:0.05, 5.00:0.10, and 5.00:0.20 mg/mL in 1.00 mL). The mixture of EGCG/DX solutions (1.00 mL) and FD/TH/PG solution in DI water (1.00 mL) was stirred at 37 °C for 30 min. When the centrifugation was completed, the particle size distributions and zeta potentials of the obtained NPs were analyzed by a Zetasizer Nano apparatus (Malvern Instruments Ltd., Worcestershire, England, UK). The residual EGCG or DX content in the supernatant was assessed. The drug-loading efficiency of the NPs was determined using a reversed-phase high-performance liquid chromatography (HPLC) equipped with a C18 column and UV detector (230 nm).

#### *2.2. Assessing pH Sensitivity and Evaluating Drug Release Profiles in EGCG/DX-Loaded FD/TH/PG NPs*

The size distributions and morphological changes of NPs at pH 7.4, 6.5, and 5.0 were assessed using a Zetasizer instrument and transmission electron microscopy (TEM, JEOL Ltd., Tokyo, Japan) to evaluate the stability of the NPs. Buffers with pH 7.4, 6.5, and 5.0 were used to simulate the circulatory system, tumor tissue, and endosomal compartment environment, respectively [33]. The pH sensitivity of NPs was determined by using an HPLC system to measure the release of EGCG and DX under different pH environments. Briefly, 2.0 mg/mL of NP solution was immersed in a dialysis bag and dialyzed against different pH values of 5.0 (10.0 mM acetic acid/sodium acetate), 6.5, and 7.4 (10.0 mM phosphate-buffered saline (PBS)), respectively. The released solution was sampled at specific time intervals and replaced with an equal volume of fresh buffer. The amount of drug release under different pH environments was evaluated with a standard calibration curve and the experiment was repeated five times under each condition.

#### *2.3. GC Cell Culture and the Anticancer Function*

Stable luciferase-expressing human GC cells (Luc MKN45) were supplied by the Japanese Collection of Research Bioresources Cell Bank (JCRB No. JCRB1379). Luc MKN45 cells can be easily detected in animals using an in vivo imaging system (IVIS, PerkinElmer lnc., Waltham, MA, USA). The culture was started in a Petri dish at a seeding density of 3 × 105 viable cells/mL in Roswell Park Memorial Institute (RPMI) 1640 medium supplemented with 10% fetal bovine serum (FBS) and 4 μg/mL of puromycin to maintain luciferase expression in a humidified atmosphere of 5% CO2/95% air at 37 °C. The cells were harvested for subculture every three days with 0.25% trypsin plus 0.05% ethylenediaminetetraacetic acid (EDTA) solution and used for the cytotoxicity experiments. In the cell cytotoxicity experiment, Luc MKN45 cells were seeded in 96-well plates at a cell density of 1.0 × 104 cells/well. After overnight incubation, the growth medium was replaced by PBS with 5% FBS containing distinct concentrations of EGCG solution (0–450 mg/L), DX solution (0–9 mg/L), EGCG/DX solution, or EGCG/DX-loaded NPs (EGCG concentration 0–450 mg/L; DX concentration 0–9 mg/L) for 2.0 h. The examined samples were extracted, and the cells were washed with PBS twice. After culturing in the growth medium for another 22.0 h, cells were incubated in growth medium with 1 mg/mL of 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) to evaluate the cell viability. The additive effect of the EGCG/DX combination on cancer cell viability was determined according to the Chou–Talalay method. The combination index (CI) was calculated using the following equation:

$$\text{CI} = \text{D}\_{\text{E}} / \text{D}\_{\text{xE}} + \text{D}\_{\text{D}} / \text{D}\_{\text{xD}} \tag{1}$$

where DE and DD are the concentrations of EGCG and DX, respectively, used to achieve x% drug effect in the combined therapy. DxE and DxD are the concentrations of EGCG and DX, respectively, used alone to achieve x% drug effect.  The association between drug concentration and inhibitory concentration (IC) for cell viability reduction was calculated and an isobologram and a combination index (CI)—fraction affected (Fa) plot were created.

#### *2.4. In Vitro Drug Cellular Uptake and Cell Distribution of NPs*

Luc MKN45 cells carrying fluorescent-dye-conjugated EGCG and DX in NPs were assessed by flow cytometry for the quantification analysis of cellular uptake ratio of EGCG and DX. The cellular uptakes of DX and EGCG were determined by detecting the fluorescence of DX itself (EX470/EM585) and cyanine5 hydrazide (Cy5)-conjugated EGCG (Cy5-EGCG), respectively. The synthesis of fluorescent Cy5-EGCG was prepared by pouring Cy5 solution (1.0 mg/0.1 mL in DMSO) into EGCG aqueous solution (0.2 g/20.0 mL). After continuous stirring for 12.0 h, the fluorescence sample was freeze-dried. The 20.0 mL of DI water was added to the lyophilized Cy5-EGCG sample to precipitate and remove unreacted fluorescent dye. The suspension was centrifuged for several times for separating the precipitated Cy5 dye until no more fluorescent dye precipitation. The prepared Cy5-EGCG solutions were lyophilized with a freeze dryer. Fluorescent Cy5-EGCG/DXloaded FD/TH/PG NPs were prepared as described before. In brief, Luc MKN45 cells were treated with Cy5-EGCG/DX solution and Cy5-EGCG/DX-loaded NPs for 2.0 h. The medium was washed with PBS and suspended and collected in 0.5 mL of PBS. A Becton Dickinson FACSCalibur flow cytometer (Becton Dickinson, Franklin Lakes, NJ, USA) was used to assess the Cy5-EGCG and DX content in 1.0 × 104 cells, and the Cell Quest software WinMDI (Verity Software House, Inc., Topsham, ME, USA) was used for analysis.

The cellular distribution of fluorescent NPs was further observed by confocal laser scanning microscopy (CLSM, Zeiss LSM 880, Jena, Germany). Fluorescent rhodamine 6G-conjugated EGCG (Rh6G-EGCG) was prepared by the procedure of Cy5-EGCG. In addition, fluorescent polymer Cy5 hydrazide-labeled FD (Cy5-FD), Alexa594 hydrazide-labeled TH (Alexa594-TH), and Atto425 *N*-hydroxysuccinimide ester-labeled PG (Atto425-PG) were synthesized. These fluorescent solutions (Cy5, Alexa594, and Atto425; 1.0 mg/0.1 mL in DMSO) were gently poured into aqueous FD solutions, TH and PG (0.2 g/20.0 mL) in DI water, respectively. The reaction mixtures were stirred for 12.0 h and dialyzed against DI water in the dark for detaching the unconjugated fluorescent dye. The residue of Cy5-FD, Alexa594-TH, and Atto425-PG solutions were lyophilized in a freeze dryer to prepare distinct fluorescence samples. In brief, Luc MKN45 cells at a density of 3 × 105 cells/cm2 were seeded on glass coverslips. After preculture for 24.0 h and incubation of fluorescent Rh6G-EGCG/DX solutions and Rh6G-EGCG/DX-loaded Cy5-FD/Alexa594-TH/Atto425-PG NPs for 2.0 h, the medium was washed with PBS. The cells were fixed with 3.7% paraformaldehyde (PFA), and the nuclei were stained with 4,6-diamidino-2-phenylindole (DAPI) and observed by CLSM.

#### *2.5. Evaluation of NP Binding to P-Selectin and CD44s*

To determine the capability of FD and TH in NPs in terms of the binding specificity of P-selectin and CD44, human recombinant P-selectin or CD44 (0.5 μg/50 μL) was added to high-hydrophobicity 96-well ELISA plates and incubated overnight at 4 ◦C. After being washed with PBS, the wells were blocked with 0.1 mL bovine serum albumin (BSA) for 1.0 h. After washing with PBS again, the wells were incubated with different concentrations of fluorescent Rh6G-FD/fluoresceinamine (FA)-TH/PG solution or Rh6G-FD/FA-TH/PG/EGCG NPs for 30 min and gently washed thrice with PBS. Fluorescence images were taken with an EnSpire™ (PerkinElmer lnc., Waltham, MA, USA) Multilabel Plate Reader by averaging 16 reads per well (excitation 525 nm/emission 548 nm for Rh6G-FD; excitation 480 nm/emission 520 nm for FA–TH). Test fluorescent sample solutions (400 μg/mL) and anti-P-selectin or anti-CD44 antibody (2.0 μg/mL) were added to the wells of the previously coated P-selectin or CD44 for the comparisons of binding specificity. The total volume of the resultant mixture in each well was 50 μL. After incubating for 0.5 h, fluorescence measurements were executed. The effect of the prepared NPs on cell surface protein expression was evaluated by culturing 3 mL of Luc MKN45 cells at a density of 3 × 105/mL on glass coverslips for 24.0 h at 37 °C.

The fluorescent Rh6G-FD/FA-TH/PG/EGCG NPs were treated to the cells for 2.0 h. Then, NPs were fixed in 1.0% PFA and stained for P-selectin or CD44 for immunofluorescence experiments. The fixed cells were blocked in PBS with 1.0% BSA and Tween 20 for

1 h, and a mixture solution of primary antibodies (rabbit anti-P-selectin antibody or mouse anti-CD44 antibody) was used for treatment overnight at 4 °C. After washing with PBS, the incubation of cells in a mixture of anti-rabbit Cy5® secondary antibody or anti-mouse Alexa Fluor® 594 secondary antibody was performed for 1.0 h in the dark. Subsequently, the cells were uniformly mounted on slides and examined using CLSM.

#### *2.6. Flow Cytometric Analysis of Cell Cycle and Western Blotting Investigation of Apoptosis-Related Proteins*

The incubation of Luc MKN45 cells in the tested EGCG/DX-loaded NPs (EGCG concentration 0–200 mg/L; DX concentration 0–4 mg/L) for 2.0 h was used to evaluate the efficiency of EGCG/DX-loaded NPs on the cell cycle. Then, the cells were washed with PBS and incubated in cell cultured medium for 22.0 h. The cells were again washed with ice-cold PBS and fixed in ice-cold 70% ethanol. After the cells were washed with PBS for the third time, hypotonic buffer was utilized for suspending cells. Subsequently, cells were incubated for 1.0 h at 37 ◦C. The propidium iodide (PI; 1.00 mg/mL, 0.01 mL) was applied to the buffer, and the cells were again incubated for 0.5 h at 4 ◦C in the dark. The stages of the cell cycle were evaluated by PI and a Becton Dickinson FACSCalibur flow cytometer. After treating Luc MKN45 cells with EGCG/DX-loaded NPs with different drug concentrations, Western blotting analysis was utilized to assess the expression of apoptotic proteins. Cell lysis was performed by using freeze-thaw cycles, and the cell lysates were centrifuged at 6000× *g* at 4 ◦C. Cell protein lysates were separated into the same amounts with electrophoresis using sodium dodecyl sulfate-polyacrylamide gel electrophoresis, and the isolated proteins were resettled to polyvinylidene difluoride (PVDF) membranes. The PVDF membranes were put in 5% (*w*/*v*) nonfat dry milk buffer for 1.0 h and then probed with the following primary antibodies at 4 ◦C overnight: rabbit polyclonal anti-caspase-9 and anti-poly(ADP-ribose) polymerase (PARP), mouse monoclonal anti-caspase-3, and anti-α-tubulin. Finally, the PVDF membranes were incubated with horseradish peroxidaseconjugated secondary antibodies for 1.0 h, and the immune complexes were evaluated using enhanced chemiluminescence.

#### *2.7. Estimation of Anti-Tumor Activity of Bioluminescent Imaging and IVIS Spectrum Detection on Nanoparticles in Tumors*

Animal use and care in this study complied with the 1996 revision of the Guide for the Care and Use of Laboratory Animals prepared by the Institute of Laboratory Animal Resources, National Research Council and published by the National Academy Press. Male six-week-old mice with severe combined immunodeficiency (SCID) and weighing 25 g, purchased from the National Laboratory Animal Center, were placed in the Laboratory Animal Center at Yang Ming Chiao Tung University, Taiwan. The animal care guidelines and all experimental protocols were approved by the Institutional Animal Care and Use Committee (IACUC 1070307). The mice were kept under standard conditions (lights on from 6 AM to 6 PM) and allowed to adapt to local conditions for one week. To establish an orthotopic gastric tumor mouse model, the Luc MKN45 cells were inoculated via the orthotopic implantation method. In brief, after sterilizing the shaved abdominal skin, an incision was made to expose the stomach and the injection of Luc MKN45 cells (1 × 107/100 μL) mixed in matrix gel (Becton Dickinson, Franklin Lakes, NJ, USA) suspension into the gastric wall was performed sub-serosally. Once the stomach was returned to the peritoneal cavity, the abdominal wall and skin were sutured closed.

After the stable detection of an orthotopic gastric tumor was achieved, drug treatment was initialized. The tumor growth was monitored using IVIS spectrum detection by capturing bioluminescence signals from Luc MKN45 cells. The mice were divided into three groups randomly with six mice each, with the injection of FD/TH/PG solution (control group), 20.0 mg/kg of EGCG/0.4 mg/kg DX in EGCG/DX solution, or EGCG/DX-loaded FD/TH/PG NPs via tail vein, respectively. Luciferin was injected intraperitoneally into the mice and waited for 10 min for bioluminescence expression and images were taken by IVIS. The quantification of the bioluminescence signals was used to monitor the tumor

growth. The animals were sacrificed after the final observation and organs were taken for histological examination by hematoxylin and eosin (H&E) or immunohistochemical (IHC) staining. The efficiency of EGCG/DX-loaded NPs on tumor status was assessed by IHC staining of M30 (apoptosis marker) and Ki-67 (proliferation marker). After dewaxing the paraffin, the tissue sections were immersed in a series of concentrations of ethanol and xylene for rehydration. The sections were stained with primary antibodies of M30 CytoDeath (Peviva, Bromma, Stockholm, Sweden) and Ki67 (Thermo Fisher Scientific, Grand Island, NY, USA) after blocking with BSA. Then, secondary peroxidase antibodies were used for the probe. Finally, the distribution expression of M30 and Ki67 and tissue inflammation at different magnifications were observed under a light microscope. Meanwhile, in the in vivo NP distribution study, fluorescent-labeled Rh6G-EGCG/DX-loaded Cy5-FD/Alexa594-TH/VioTag750-PG NPs were injected via tail vein to evaluate their distribution during the treatment. After euthanizing the mice at 24 h, fluorescence images of organs and tissues were obtained in the IVIS. Moreover, the mice also received fluorescent Rh6G-FD/FA-TH/PG/EGCG NPs via 24.0 h intravenous injection. Subsequently, the gastric tumor slides were assessed using immunofluorescence staining with rabbit anti-P-selectin antibody or mouse anti-CD44 primary antibodies, probed with secondary antibodies (anti-rabbit Cy5® or anti-mouse Alexa Fluor® 594), then examined using CLSM.

#### *2.8. Statistical Analysis*

One-way analysis of variance was used to analyze differences in the measured characteristics of the treatment groups. The confidence intervals were determined using Statistical Analysis System, version 6.08 (SAS Institute, Cary, NC, USA). The mean ± standard deviation (SD) was used to represent all data and *p* < 0.05 was considered statistically significant.

#### **3. Results**

#### *3.1. Preparation and Characterization of EGCG/DX-Loaded FD/TH/PG NPs*

To create NPs that can be combined with EGCG and DX, first, the optimal material FD:TH ratio should be determined when preparing EGCG-loaded NPs and then the concentration of DX should be adjusted to create EGCG/DX-loaded NPs. As shown in Table 1, as the amount of FD/TH incorporated in FD/TH/PG/EGCG NPs increased, the mean particle size of the NPs was 214.68 ± 18.78 to 741.79 ± 198.79 nm and their polydispersity index (PDI) was 0.25 ± 0.08 to 0.95 ± 0.26. When the FD:TH:PG ratio was 0.60:0.60:3.75 mg/mL, the NPs formed a dissoluble opalescent suspension after centrifugation. The particle size was 224.68 ± 18.78 nm and the zeta potential −26.14 ± 1.75 mV, with a significantly narrow distribution (0.25 ± 0.08) (Table 1). However, when the FD:TH:PG ratio increased to 0.90:0.90:3.75 mg/mL, the average particle size of the NPs increased, as did their PDI (to 0.4), indicating high heterogeneity. To further load DX into NPs, NPs containing different concentrations of EGCG and DX were prepared using a gelation technique in which the EGCG/DX solution was added to the FD/TH/PG aqueous solution. The HPLC analysis method showed that, at an EGCG:DX ratio of 2.50:0.05 mg/mL, the loading efficiency of EGCG and DX in the NPs was 59.07% ± 7.06% and 63.46% ± 4.59%, respectively. The subsequent experiment required a narrower distribution (PDI = 0.21 ± 0.05) (Table 2 and Figure 1b).

**Table 1.** Effect of different FD/TH proportions on particle sizes, polydispersity indices, zeta potential values, and drug loading efficiency of the prepared EGCG-loaded FD/TH/PG NPs (*n* = 5). - Precipitation of aggregates after centrifugation was observed.

**Table 2.** Effect of varying DX concentration on EGCG/DX-loaded FD/THA/PG particle sizes, polydispersity indices, zeta potential values, and drug loading efficiency (*n* = 5).

In FTIR investigates of EGCG/DX-loaded FD/TH/PG NPs, the representative peaks at 1235 and 1427 cm−1 ascribed to FD were dispensed to sulfate esters (O=S=O) and CH2 (galactose, xylose) scissoring vibrations or CH3 (fucose, O-acetyls) asymmetric bending vibrations. The synthesized TH copolymer was analyzed with FTIR and showed representative peaks at 1363 and 2880 cm−1, conforming to the −CH2 group on PEG and the –CH stretching band of TPGS, and peaks at 1413 and 1626 cm−1, reflecting the contribution of C–O stretches symmetrically and C=O stretches asymmetrically, corresponding to the carboxyl group on HA. In the PEG-conjugated gelatin (PG) spectrum, characteristic peaks at 839 and 1545 cm−1 were observed, corresponding to bending C–C and –NH stretching vibrations on PEG and gelatin. Finally, EGCG/DX-loaded FD/TH/PG complex spectra showed characteristic peaks at 1090 and 1208 cm−1, corresponding to the C–OH deformation of EGCG and DX, respectively. Moreover, the NH bending peak on PG shifted from 1545 cm−1 to 1540 cm−1; the S=O or C=O stretching peaks on FD or TH shifted from 1235 cm−1 or 1626 cm−1 to 1231 cm−1 or 1631 cm−1, respectively. This indicates that the H atoms of EGCG or DX interacted via hydrogen bonds with the N atoms of PG (C−OH···N−C) or the O atoms of FD or TH (C−OH···O=S or O=C) (Figure 1c).

#### *3.2. Characterization of the Morphology and Drug Release Profile of EGCG/DX-Loaded FD/TH/PG NPs*

We evaluated the pH sensitivity of the NPs and the distinct drug release profile of EGCG/DX-loaded NPs using TEM and HPLC, respectively (Figure 2). At pH 7.4 and 6.5 (simulating the physiological fluids and tumor tissues), the NPs shown as stable spheres with a matrix structure with particle sizes of 200–230 nm, because EGCG and DX form hydrogen bonds with PG or TH. The drug release percentage from NPs over 3.0 h incubation was 6.21% ± 0.87% (EGCG) and 9.54% ± 0.15% (DX) at pH 7.4 and 7.94% ± 0.69% (EGCG) and 9.87% ± 1.35% (DX) at pH 6.5. At pH 5.0 (simulating an acidic environment in endosomal of cancer cells), some of the –COO– groups were moderately protonated in TH, affecting the negative electrostatic repulsion of NPs, which become unstable and partially collapse but with increased particle size (418.31 ± 69.75 nm) (Figure 2a). Under those conditions, the EGCG release was 24.58% ± 1.64% and the DX release was 28.17% ± 3.68% within 3.0 h. In addition, the hydrogen bonding between the N or O atom

in PG or TH, and the H atom in the hydroxyl group of EGCG or DX caused slower drug release from EGCG/DX-loaded NPs, reaching 59.74% ± 3.31% and 75.74% ± 2.08% for EGCG and DX, respectively, over 24.0 h (Figure 2b).

**Figure 2.** (**a**) Transmission electron micrographs of nanoparticles in buffer solution with different pH values; (**b**) The release profile of EGCG and DX in varying pH environments at 37 °C. Data is expressed as mean ± SD (*n* = 5).

#### *3.3. Effects of FD/TH Solutions and EGCG/DX-Loaded FD/TH/PG Nanoparticles on Binding of P-Selectin and CD44*

To study the targeting of P-selectin and CD44 by Rh6G-FD/FA-TH/PG solution or Rh6G-FD/FA-TH/PG/EGCG NPs with different FD or TH molecule concentrations (100, 200 and 400 μg/mL), their fluorescence intensity was quantified by spectrofluorometry. The evident fluorescence intensities at Rh6G-FD or FA-TH concentrations of 100 μg/mL to 400 μg/mL were 207.19 ± 24.09 to 659.21 ± 85.31 or 173.39 ± 35.19 to 480.11 ± 51.26 (only FD/TH/PG solutions), respectively and 373.91 ± 59.08 to 911.21 ± 52.61 or 231.39 ± 40.09 to 662.31 ± 60.76 (EGCG/DX-loaded FD/TH/PG NPs), respectively. Additionally, Rh6G-FD or FA-TH fluorescence analysis indicated that the contact degree of EGCG/DX-loaded NPs with P-selectin or CD44 was greater than that of FD/TH/PG solutions. The addition of P-selectin or CD44 reduced the adhesion of targeting proteins to FD or TH by approximately 50% due to blockage by the competing anti-targeting P-selectin or CD44 protein antibodies (Figure 3a). Therefore, we evaluated the benefits of the relative composition of FD and TH in P-selectin- and CD44-targeted NPs. The prepared NPs interact with protein expression on GC cell surface was observed by CLSM. After 2.0 h of incubation with fluorescent Rh6G-FD/FA-HA/PG/EGCG NPs, fluorescence signals (Rh6G-FD (red dot) and FA-TH (orange dot)) of the NPs were observed in the cell cytoplasm and intercellular spaces, and then the NPs could exhibit co-localize and interact with P-selectin (purple dots) and CD44 (green dots) in gastric cancer cells (blue or red arrows indicate superimposed red/purple or orange/green dots) (Figure 3b).

**Figure 3.** (**a**) Binding assays of fluorescence Rh6G-FD and FA-TH to immobilized recombinant P-selectin and CD44 protein. Data is expressed as mean ± SD (*n* = 4). Asterisk symbols \* and \*\* represent statistically significant differences with *p* value < 0.05 and < 0.01, respectively; (**b**) Immunofluorescence staining of P-selectin and CD44 proteins in MKN45 cancer cells incubated with Rh6G-FD/FA-TH/PG/EGCG NPs.

#### *3.4. Evaluating the Anticancer Effect of EGCG/DX-Loaded FD/TH/PG NPs and the Combination Index Determination*

To evaluate the potential inhibitory effect of EGCG or DX alone and the synergy of the EGCG/DX solution, the effects of EGCG/DX-loaded NP treatment on GC cell viability was analyzed. Both EGCG and DX significantly affected cell growth at high concentrations of 450 and 9 mg/L, respectively, and the cell viability decreased to 50.59% ± 3.46% and 50.81% ± 5.11%, respectively (Figure 4). We also compared the efficacy of EGCG/DX solution and EGCG/DX-loaded NP on inhibiting GC growth. The cell survival after EGCG/DX-loaded NP treatment was obviously lower than that after EGCG/DX solution treatment. The anti-cancer activity of EGCG/DX-loaded FD/TH/PG NPs (EGCG concentration, 150–450 mg/L; DX concentration, 3–9 mg/L) on Luc MKN45 cells was significantly higher than that of the EGCG/DX combined solution (*p* < 0.05; Figure 4c). The therapeutic IC50 of EGCG/DX-loaded NPs was 200/4 mg/L, which was half that of the EGCG/DX combined solution (400/8 mg/L).

In addition, the CI-Fa plot and isobologram are described in Figure 4d. EGCG/DX combination treatment had a slightly synergistic effect on the cell viability, with CI of IC10 and IC20 being 0.86 and 0.94, respectively. EGCG/DX-loaded NP treatment showed a better synergistic antiproliferation effect, with CIs of IC10, IC20 IC30, IC40, and IC50 being 0.56, 0.48, 0.70, 0.80, and 0.89, respectively, compared with EGCG/DX combination solution. This result showed that GCG/DX-loaded FD/TH/PG NP targeting cancer therapy has better benefits than EGCG/DX combined drug therapy.

**Figure 4.** The effects of EGCG, DX, EGCG/DX combination solution, and EGCG/DX-loaded NPs on Luc MKN45 cell viability: (**a**) A 2.0 h dose response treatment with EGCG; (**b**) A 2.0 h dose response treatment with DX; (**c**) A 2.0 h dose response treatment with EGCG/DX solution and EGCG/DX-loaded NPs; cell viability was assessed at 24.0 h for a–c; data are expressed as mean ± SD (*n* = 8). Asterisk \* represents statistically significant difference with *p* value < 0.05. (**d**) Combination index–fraction affected (CI-Fa) plot; CI < 1.0 indicates synergistic effects, CI = 1.0 indicates additive effects and CI > 1.0 indicates antagonistic effects of the combined drug action. Dots below the slanted line indicate synergistic effects.

#### *3.5. Internalization of EGCG and DX and Cellular Uptake of Complexes Contained within Fluorescent Nanoparticles*

To confirm the cellular uptake ability, fluorescence-activated cell sorting analysis was made after the cells were treated with both the drug combination and NPs containing Cy5-EGCG and DX. Flow cytometry examination was performed with Luc MKN45 cells after treatment with FD/TH/PG NPs loaded with EGCG/DX. The cellular uptake and fluorescence intensity of Cy5-EGCG were 86.53% ± 3.25% and 27,639.98 ± 254.91, respectively, while those of DX reached 60.42% ± 1.56% and 13,493.23 ± 143.87, respectively, after exposure for 2.0 h. In contrast, after treatment with only EGCG/DX solution, the cellular uptake and total fluorescence intensity of Cy5-EGCG were 56.29% ± 1.32% and 18,783.54 ± 198.40, respectively and those of DX were 28.62% ± 0.97% and 6275.41 ± 60.43, respectively (Figure 5a). This difference in drug absorption capacity is closely related to the presence of EGCG/DX in NPs, which successfully identified the specific targets, namely P-selectin and CD44, on the cell surface.

**Figure 5.** (**a**) In vitro cellular uptake of Cy5-GCG and DX was assessed using flow cytometry after treatment with EGCG/DX combination solution and EGCG/DX-loaded NPs; (**b**) Confocal images of gastric cancer cells viewing cellular distribution of the Rh6G-EGCG/DX combination solution and Rh6G-EGCG/DX-loaded Cy5-FD/Alexa594-TH/Atto425-PG NPs, \*\*\* represent statistically significant differences with *p* value < 0.001.

CLSM was used to compare and evaluate the delivery efficiency of EGCG/DX solution and EGCG/DX-loaded FD/TH/PG NPs after 2.0 h treatment, and DAPI with fluorescence expression was used to stain cell nuclei. Cy5-FD, Alexa594-TH, and Atto425-PG were co-localized in the cells. The internalization of EGCG and DX was evaluated by observing Rh6G-EGCG and DX fluorescence performance signals. Drug content (Rh6G-EGCG and DX) was higher in GC cells treated with the EGCG/DX-loaded NPs than in those treated with the EGCG/DX solution, indicating that the NP system infiltrated cancer cells and improved the delivery of EGCG and DX (Figure 5b).

#### *3.6. Cell Cycle Arrest and Apoptosis Effects of NPs Loaded with EGCG/DX*

To investigate whether the decrease in the growth rate of cells treated with test samples is associated with cell cycle arrest, we analyzed PI-stained Luc MKN45 cells treated with EGCG/DX-loaded NPs containing different EGCG and DX concentrations within 24.0 h by flow cytometry. As shown in Figure 6a, after EGCG/DX-loaded NP treatment, the cells displayed significant accumulation in the G2/M phase compared to the control group.

In particular, EGCG/DX-loaded NPs significantly reduced the G0/G1 population (from 57.28% ± 2.35% to 18.66% ± 4.51%), the number of cells undergoing the S phase slightly decreased (from 16.48% ± 0.87% to 12.11% ± 5.03%), and the accumulation of G2/M in cells (from 25.11% ± 1.73% to 67.86% ± 3.95%) increased in an EGCG/DX dose-dependent manner, indicating that EGCG/DX-loaded NPs affect cell cycle arrest at the G2/M phase in GC cells. To determine whether the EGCG/DX-loaded NPs can induce cell apoptosis, we performed Western blotting and evaluated the ratios of cleaved caspase-9, caspase-3, and poly(ADP-ribose) polymerase (PARP) levels to their total protein expression (cleaved form plus proform). α-Tubulin was used as an internal control. Twenty-four hours after the samples were treated with different drug concentrations, EGCG/DX (200/4 mg/mL) was found to increase the cleaved protein to total protein expression ratios of caspase-9, caspase-3, and PARP by approximately 18%, 45%, and 69%, respectively, compared with that in the untreated group (Figure 6b).

**Figure 6.** (**a**) The proportions of cells in the G0/G1, S, and G2/M phases, after treatment with EGCG/DX-loaded FD/TH/PG NPs are indicated. The cells were stained with propidium iodide and the cell cycle distribution was analyzed using flow cytometry. (**b**) Western blotting analysis of apoptosis-related caspase-9, caspase-3, and PARP in MKN45 cells after incubation with EGCG/DX-loaded FD/TH/PG NPs and α-tubulin was used as an internal control. \* Statistically significant difference of *p* < 0.05 and \*\* *p* < 0.01, respectively as compared with the without sample treatment group.

#### *3.7. Bioluminescent Imaging to Evaluate Anticancer Activity in an Orthotopic Gastric Tumor Mouse Model*

We examined tumor-specific effects of EGCG/DX-loaded NPs in the gastric tumor mouse model of a luciferase-expressing gastric tumor. Male SCID mice were inoculated with EGCG/DX combination solution (20 mg/kg EGCG/0.4 mg/kg DX) and EGCGloaded FD/TH/PG NPs or FD/TH/PG solution as a control by using intravenous tail injection every three days for a total of five injections. The bioluminescence signals of gastric tumor pointedly increased by 3.87-fold ± 0.39-fold (FD/THA/PG solution) and 3.10-fold ± 0.29-fold (EGCG/DX solution) at day 18. Furthermore, EGCG/DX-loaded FD/THA/PG NPs (2.16 ± 0.35 units) better inhibited tumor growth, with a lower relative photon flux compared with mice in other treatment groups (Figure 7a). There was no difference in body weight loss in our EGCG/DX-loaded NPs group and it was also superior to that in the FD/TH/PG control solution, indicating significant antitumor activity (Figure 7a). We also used IVIS to further evaluate the local accumulation of NPs. After fluorescent Rh6G-EGCG/DX-loaded Cy5–FD/Alexa594-TH/VioTag750-PG NPs were used for tail vein injection, fluorescent signals were detected on different tissues by IVIS technology. The gastric tumor was visualized by detecting bioluminescent signals formed by stable luciferase-expressing Luc MKN45 cells. The intensity of the fluorescent signals (Rh6G-EGCG and DX) at the tumor site significantly increased locally within 24.0 h, indicating continuous and sustained drug delivery by NPs targeting P-selectin and CD44 in the tumor. After 24.0 h, the mice were sacrificed to determine the total fluorescent counts in tumor and normal tissues and an increase in the NP distribution in the tumor led to a decrease in their distribution in other organs. Therefore, the tumor-targeting ability of EGCG/DX-loaded FD/TH/PG NPs was effective in the orthotopic gastric tumor mouse model (Figure 7b).

#### *3.8. Histological and Immunohistochemical Analyses of Cancer Biomarkers*

After the experimental mice were sacrificed, H&E staining was used for histological examination of gastric tumors and other organ biopsies (Figure 8). Compared with the EGCG/DX-loaded NP groups, tumor sections from the EGCG/DX combination group had more granular eosinophil infiltration (black arrow). The EGCG/DX-loaded NP group showed increased tumor necrosis, characterized as grade 2 tissue necrosis (necrosis of tumor cells or disappearance of more than 2/3 cells; the right side of the red line) compared with the EGCG/DX combination group (Figure 8a). Additionally, immunohistochemical analysis presented that after EGCG/DX-loaded FD/TH/PG NP treatment, M30 expression in tumors increased, while Ki67 expression diminished (i.e., coffee dots; green arrows, Figure 8a). These findings showed that targeted NPs induce GC cells apoptosis, resulting in a significant increase in anticancer activity with tumor necrosis. In the EGCG/DX combination group, the lung tissue showed inflammatory cell exudation, red blood cells scattered in multiple alveolar cavities, and thickened interstitial pulmonary edema (i.e., black arrows, Figure 8b). In contrast, the pathological damage was evidently minor in the EGCG/DX-loaded NP group, in which the alveolar structures were clear, and signs of inflammation decreased (i.e., black arrows, Figure 8b). Moreover, the hepatocyte swelling density and neutrophil infiltration situation of liver tissue biopsies in the EGCG/DXloaded NP group was less than those in the EGCG/DX combined group (i.e., red arrows, Figure 8b). Therefore, targeting NPs can not only significantly improve the antitumor ability of drugs against GC, but also reduce the inflammatory response in the body.

**Figure 7.** (**a**) The antitumor effects of different samples in orthotopic Luc MKN45 xenograft mice model. Mice were divided into three groups of six mice each and treated with FD/TH/PG solution (-), EGCG/DX combination solution (•), or EGCG/DX-loaded FD/TH/PG NPs (). \* Statistically significant difference of *p* < 0.05 and \*\* *p* < 0.01. (**b**) Distributions of fluorescent Rh6G-EGCG/DX-loaded Cy5-FD/Alexa594–TH/VivoTag750-PG in tumor and organs after treatment. Gastric tumors were detected with a bioluminescent in vivo imaging system. (**c**) Immunofluorescence staining of P-selectin and CD44 proteins in gastric tumor slides treated with Rh6G–FD/FA-TH/PG/EGCG NPs.

**Figure 8.** (**a**) Histological analysis and immunohistochemical analysis of Ki67 (proliferation marker) or M30 (apoptosis marker) of gastric tumor biopsy after treatment with EGCG/DX combination solution or EGCG/DX-loaded FD/TH/PG NPs; (**b**) Histological analysis of different organs in mice bearing gastric tumor after treatment with EGCG/DX combination solution or EGCG/DX-loaded FD/TH/PG NPs.

#### **4. Discussion**

Cell adhesion is the process by which cells interact and attach to neighboring cells through specialized molecules, regulating cell motility and migration [37,38]. Adhesion molecules of the selectin family (E-, L-, and P-selectin) are a class of membrane-bound protein that participate in the initial adhesion of leukocytes to the site of infection or

inflammation [37]. P-selectin expression increases in the vasculature of human cancers and facilitates metastasis by promoting the adhesion of circulating cancer cells to activated platelets and endothelial cells in distant organs [39,40]. P-selectin can also bind with high affinity to some sulfatides and sulfated polysaccharides, such as heparin, dextran sulfate, and FD [41,42]. Park et al. used FD extracted from *Fucus vesiculosus* to cause autophagic cell death of human GC cells and show antiproliferation activity via Beclin1 upregulation and the conversion of microtubule-associated protein (MAP) light chain 3-I (LC3-I) to LC3-II [43]. P-selectin binds to CD44 and promotes the capture of vascular endothelial cells and cancer cells by leukocytes [44]. HA, a major extracellular matrix component, is a physiological ligand that is overexpressed in CD44 on the surface of malignant tumor cells and can be used to selectively activate multiple oncogenic signaling pathways that lead to tumor cell-specific phenotypes [45]. CLSM showed that the fluorescence signals (Rh6G-FD (red dot) and FA-TH (orange dot)) of the prepared NPs were co-localized and interacted with P-selectin (purple dot) and CD44 (green dot) in gastric tumor tissue (white arrows)(Figure 7c). Rh6G-EGCG and DX fluorescence signals emanated by Rh6G-EGCG/DX-loaded Cy5-FD/Alexa594-TH/Atto425-PG NPs were attached to Luc MKN45 cells, and more drugs (EGCG and DX) were released into the cell cytoplasm compared with EGCG/DX combination treatment (Figure 5). Furthermore, the carrier EGCG/DX induced cell cycle arrest in the G2/M phase, inhibited GC proliferation, and enhanced the expression of apoptosis-related proteins (Figure 6).

Tumor cell growth generates a complex physiological environment that is distinct from regions surrounding healthy tissues. Both the extracellular and intracellular pH values of tumor cells are considered to be acidic and lower than those of normal cells [46,47]. Physiological pH is generally believed to be in the range of 7.2–7.4 for normal tissues and the circulatory system [48]. However, the pH value of tumor cells is weakly acidic, between 6.5 and 7.0, and the pH value of endosomes and lysosomes is between 5.0 and 6.0 [49]. Therefore, the pH of the tumor microenvironment and tumor cells plays an important role in the development and treatment of cancer. The NP system adopts a pH-sensitive adjustment strategy to prevent the untimely release of drugs. In the present study, NPs containing EGCG/DX were designed with a matrix structure that minimizes drug release into normal physiological environments, such as nontarget tissues and blood, and promotes the release of EGCG and DX into lysosomes and late endosomes of the tumor cells. The cumulative release of EGCG and DX at pH 5.0 amounted to 59.74% ± 3.31% and 75.74% ± 2.08%, respectively (Figure 2). Specifically, in vivo experiments confirmed that the lower bioluminescence intensity after intravenous injection of EGCG/DX-loaded NP (20 mg/kg EGCG/0.4 mg/kg DX) treatment significantly inhibited tumor growth (Figure 7). Histological analyses showed that the delivery of EGCG/DX to tumors through NPs can induce GC cell apoptosis, as indicated by increased levels of caspase-cleaved cytokeratin 18 (M30, ccK18) as an apoptotic cell death pharmacodynamic biomarker, accompanied by a reduction in inflammatory pulmonary lesions (Figure 8).

#### **5. Conclusions**

In summary, we applied an FD-based and TH-based NP system for the active, targeted co-delivery of EGCG and DX through P-selectin and CD44 ligand recognition to GC cells. Drug release was regulated by the NP system using a pH-sensitive adjustment strategy. In addition, the NPs prepared for the co-drug delivery system significantly improved the synergistic anti-cancer effect of EGCG and DX compared with that of the EGCG/DX combination solution. Targeting enhanced NP distribution in GC cells and decreasing it in major organs in the orthotopic gastric tumor could lead to the development of novel cancer therapies and facilitate their clinical trials.

**Author Contributions:** W.-Y.H. and C.-H.L. contributed equally to this work and should be considered co-first authors. W.-Y.H. and C.-H.L. conceived and designed this work. S.-L.P., C.-Y.H., and P.-H.H. performed the experimental study. P.-Y.C. and C.-L.F. analyzed the data and discussed the

results. Y.-H.L. supervised the research and obtained funding. All authors have read and agreed to the published version of the manuscript.

**Funding:** This work is particularly supported by Ministry of Science and Technology, grant number MOST 107-2221-E-010-004-MY3 and Yin Yen-Liang Foundation Development and Construction Plan of the School of Medicine, National Yang-Ming University. The funders had no role in study design, data collection and analysis, decision to publish or preparation of the manuscript.

**Institutional Review Board Statement:** The study was conducted according to the guidelines of the Declaration of Helsinki, and approved by the Institutional Animal Care and Use Committee of Laboratory Animal Center at Yang Ming Chiao Tung University (IACUC 1070307 and 1 January 2019 of approval).

**Acknowledgments:** We thank the Laboratory of the Leiss LSM 880 confocal Spectral microscopy experiment and BD FACSCalibur flow cytometer provided by core facility of National Yang Ming Chiao Tung University. The authors would like to thank Enago (www.enago.tw) for the English language review.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**

## *Article* **AS1411 Aptamer Linked to DNA Nanostructures Diverts Its Traffic Inside Cancer Cells and Improves Its Therapeutic Efficacy**

**Giulia Vindigni 1,†, Sofia Raniolo 1,2,†, Federico Iacovelli 2, Valeria Unida 2, Carmine Stolfi 1, Alessandro Desideri 2 and Silvia Biocca 1,\***

**Abstract:** The nucleolin-binding G-quadruplex AS1411 aptamer has been widely used for cancer therapy and diagnosis and linked to nanoparticles for its selective targeting activity. We applied a computational and experimental integrated approach to study the effect of engineering AS1411 aptamer on an octahedral truncated DNA nanocage to obtain a nanostructure able to combine selective cancer-targeting and anti-tumor activity. The nanocages functionalized with one aptamer molecule (Apt-NC) displayed high stability in serum, were rapidly and selectively internalized in cancer cells through an AS1411-dependent mechanism, and showed over 200-fold increase in anti-cancer activity when compared with the free aptamer. Comparison of Apt-NCs and free AS1411 intracellular distribution showed that they traffic differently inside cells: Apt-NCs distributed through the endo-lysosomal pathway and were never found in the nuclei, while the free AS1411 was mostly found in the perinuclear region and in nucleoli. Molecular dynamics simulations indicated that the aptamer, when linked to the nanocage, sampled a limited conformational space, more confined than in the free state, which is characterized by a large number of metastable conformations. A different intracellular trafficking of Apt-NCs compared with free aptamer and the confined aptamer conformations induced by the nanocage were likely correlated with the high cytotoxic enhancement, suggesting a structure–function relationship for the AS1411 aptamer activity.

**Keywords:** G-quadruplex; nucleolin; DNA nanocages; intracellular localization; cancer targeting; molecular dynamics simulations

#### **1. Introduction**

Aptamers are short single-stranded DNA or RNA oligonucleotides with stable threedimensional structures capable of binding with high affinity to a variety of molecular targets [1]. A unique class of aptamers is constituted of G-rich sequences that, under physiological conditions, spontaneously fold into non-canonical four-stranded structures named G-quadruplexes. These are characterized by the stacking of two or more successive planes of four guanine residues arranged in a square planar array via Hoogsteen hydrogen bonding (a G-quartet). G-quadruplex structures are highly polymorphic, and a single sequence can fold into several different conformations, depending on the relative orientations of strands and types of loops [2–4]. The 26-base G-rich AS1411 aptamer has been identified as an anti-cancer agent, which specifically recognizes nucleolin on the cancer cell surface with high selectivity and affinity [5,6]. As other G-quadruplex aptamers, AS1411 adopts a mixture of multiple structures in solution, and one of them has been solved through NMR spectroscopy, studying a DNA sequence named AT11, where a single guanine has

**Citation:** Vindigni, G.; Raniolo, S.; Iacovelli, F.; Unida, V.; Stolfi, C.; Desideri, A.; Biocca, S. AS1411 Aptamer Linked to DNA Nanostructures Diverts Its Traffic Inside Cancer Cells and Improves Its Therapeutic Efficacy. *Pharmaceutics* **2021**, *13*, 1671. https://doi.org/ 10.3390/pharmaceutics13101671

Academic Editors: Marina Santiago Franco and Yu Seok Youn

Received: 8 September 2021 Accepted: 7 October 2021 Published: 13 October 2021

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

been substituted by a thymine. This mutated version shows a preferential G-quadruplex conformation and exhibits an anti-proliferative activity comparable to that of AS1411 [4,7].

The AS1411 target molecule, nucleolin, is a multifunctional cyto-nucleoplasmic protein mostly localized in the nucleoli in normal cells and aberrantly overexpressed in many types of cancers, where it is also located on the cell surface [8]. In cancer cells, the surface nucleolin acts as a receptor of oncogenic ligands and shows protumorigenic function regulating the biosynthesis of specific oncomiRNAs [9]. Following its interaction with nucleolin, AS1411 inhibits cancer cells proliferation and induces cell death, with little effects on normal cells. AS1411 exhibits antiproliferative effects by upregulating p53, downregulating Bcl-2, and inhibiting cancer cell migration and invasion in an Akt1-dependent manner [10–12]. Due to its unique properties, AS1411 has been used for its anticancer therapeutic effect and, when linked to nanoparticles, as a selective tumor-targeting ligand [13,14]. Notably, AS1411 aptamer treatment was evaluated in phase II clinical trials in patients with acute myeloid leukemia and renal cell carcinoma [15], but it never reached phase III.

In the last years, we have been involved in the in silico, in vitro, and in cell characterization of a DNA-based truncated octahedral nanocage family [16–21]. These nanocages are made by 12 double helices, obtained by assembling 8 oligonucleotides whose ends are covalently linked in order to produce fully covalent structures, which ensure high stability in biological fluids and inside cells. We have characterized the receptor-mediated cell internalization of DNA nanostructures and their efficacy in selective doxorubicin delivery and miRNA silencing in cancer cells [22–24]. When functionalized with folate molecules and tailored with anti-miR21 complementary sequences, DNA nanocages induce selective miR21 sequestering and cytotoxicity to tumor cells overexpressing the α isoform of the folate receptor [25].

In this work, with the aim of increasing AS1411 stability and therapeutic efficacy, we designed octahedral DNA nanocages harboring one AS1411 molecule (Apt-NC) and studied its efficiency as targeting ligand and cytotoxic molecule against cancer cells. We studied the in vitro and intracellular stability, the time-dependent cell internalization, the targeting selectivity, and the therapeutic efficacy of the new assembled nanocages. Interestingly, the AS1411-functionalized nanocages displayed higher cytotoxic efficacy and different intracellular trafficking than the free AS1411 aptamer. Molecular dynamics simulations of two 3D models of free AS1411 in comparison with the aptamer-functionalized nanocage indicated that the free aptamer sampled a large number of metastable conformations, while the conformations were more localized and confined into a low-energy conformational basin when linked to the nanocage. These results indicate that the nanocage constrains the aptamer in a more defined conformation, likely correlated to the different intracellular localization and higher therapeutic effect.

#### **2. Materials and Methods**

#### *2.1. Preparation of Aptamer-Functionalized DNA Nanocages*

AS1411-functionalized nanocages (Apt-NC) and non-functionalized nanocages (NC) were prepared as described [21]. A biotin molecule was added on one edge of the structure for the detection of nanostructures through the streptavidin–biotin reaction. Briefly, nanocages were assembled by mixing equimolar amounts of 8 oligonucleotides in TAM buffer (40 mM Tris–acetic acid, pH 7.0, 12.6 mM magnesium acetate). The sequences of the oligonucleotides used for the assembly of NCs and the sequence of free AS1411 aptamer are reported in Supplementary Table S1. After assembly, nanocages were incubated for 2 h at 25 ◦C with T4 DNA ligase (New England Biolabs Inc., Ipswich, MA, USA) to covalently link the obtained structures and run on native 5% polyacrylamide gels in TAEM buffer (40 mM Tris–acetic acid, pH 7.0, 1 mM EDTA, 12.6 mM magnesium acetate). The band corresponding to the correctly assembled nanocages was cut out of the gel, eluted, and concentrated by 2-propanol precipitation.

#### *2.2. Cell Cultures*

HeLa cells, derived from human cervix cancer, and CHO, Chinese hamster ovary, cells were grown in DMEM (Dulbecco's modified Eagle medium, Euroclone, Devon, UK) and DMEM/F-12 (Dulbecco's modified Eagle medium/Nutrient Mixture F-12, Euroclone, Devon, UK), respectively, supplemented with 10% FBS (Gibco, Paisleg, UK), 1 mM L-glutamine (Sigma Aldrich, St Louis, MO, USA), 1mM sodium pyruvate (Biowest, Miami, FL, USA), and 100 U/ml penicillin–streptomycin (Euroclone, Devon, UK).

#### *2.3. DNA Nanocages Stability*

Biotinylated cages were incubated in TBS (Tris-HCl 50 mM, NaCl 150 mM, pH 7.8) or in culture medium supplemented with 10% FBS at 37 ◦C for different times. Each sample was then digested with proteinase K (100 μg/mL) for 1 h at 37 ◦C, and analyzed by DNA blot, as previously described [21]. Biotin detection was carried out using streptavidin– HRP (horseradish peroxidase) (Abcam Inc., Toronto, ON, Canada), and visualized by enhanced chemiluminescence (ECL Extend, Euroclone, Devon, UK). For image processing and densitometric analysis, photographic films were digitized by scanning and bands analyzed by ImageJ software.

#### *2.4. Purification of DNA Nanocages and DNA Blot*

Cells were plated in 48-well plates one day before treatments. After incubation with DNA nanocages for different time periods, cells were lysed, centrifuged, digested with proteinase K, and DNA blot was performed as previously described [21].

#### *2.5. Confocal Analysis*

Cells were seeded onto poly-L-lysine-coated glass cover-slides. For binding experiments, cells were incubated with biotinylated nanocages in DMEM 10% FBS for 1 h at 4 ◦C, washed in PBS, fixed in 4% paraformaldehyde, and incubated for 5 min with NaBH4. For uptake experiments, cells were incubated with nanocages at 37 ◦C for different time periods, fixed with 4% paraformaldehyde, and permeabilized for 4 min with Tris/Triton (Tris HCl 0.1 M, Triton 0.1%, pH 7.7). Rabbit polyclonal anti-flotillin-1 antibody (Santa Cruz Biotechnology Inc., Dallas, TX, USA) was used to detect the flotillin-1 protein. Early endosomes were visualized with polyclonal EEA1 antibody (Abcam Inc., Toronto, ON, Canada) and lysosomes with mouse monoclonal anti-LAMP-1 antibody (Abcam Inc., Toronto, ON, Canada). Donkey anti-rabbit IgG and donkey anti-mouse IgG, both Rhodamine Red-Xconjugated AffiniPure (Jackson ImmunoResearch, Cambridgeshire, UK) were used as secondary antibodies [24]. Biotinylated cages were detected by using streptavidin–FITC (Jackson ImmunoResearch, Cambridgeshire, UK). The nuclei were stained with DAPI (Invitrogen, Carlsbad, CA, USA). Images were obtained with a laser confocal fluorescent microscope Olympus FV1000 at 60× magnification, and the fluorescence signal was evaluated with the IMARIS software. Co-localization events were evaluated as previously described [24].

#### *2.6. Cell Viability Assay*

HeLa and CHO cells were plated in 96-well plates at a density of 5 × 103 cells/well and 7 × 103 cells/well, respectively, grown for 24 h, and treated with different concentrations of Apt-NCs, NCs, or free AS1411 aptamer for 24 h at 37 ◦C. The MTS assay (3-(4,5 dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium) (Promega, WI, USA) was used for evaluating cell proliferation. Multiskan Ascent 96/384 plate reader (MTX Lab Systems, Bradenton, FL, USA) was used for measuring the absorbance at 492 nm. Cell viability was normalized to control condition (untreated cells).

#### *2.7. Statistical Analysis*

All experiments were carried out in triplicate and data analyzed using GraphPad Prism. Results were expressed as a mean ± S.E.M and statistical analyses performed using Student's *t*-test. Differences were considered statistically significant when *p* < 0.05 (\*), *p* < 0.01 (\*\*) and *p* < 0.001 (\*\*\*).

#### *2.8. Computational Methods*

Two models of the AS1411 G-quadruplex were generated. The first one (Supplementary Figure S1, AS1411) was generated starting from the PDB structure with ID:2N3M corresponding to the AT11 variant of the aptamer [4] using the fiber module of the 3DNA program [26] to generate the PDB file. The nucleotide sequence of the strand composing the G-quadruplex was modified through the 3DNA mutate\_bases module [26] to fully match the AS1411 oligonucleotide sequence. The PyMol sculpting module [27] was used to generate an additional conformation, imposing guanines 11 as part of the first G-quartet of the aptamer (Supplementary Figure S1, AS1411\*). The structures were minimized using the UCSF Chimera program [28] to remove any clashes and unwanted interactions introduced by the modeling procedure. The octahedral scaffold of the DNA nanocages was built through our Polygen software [16] designing eight oligonucleotide sequences (Supplementary Table S1) based on those previously used to experimentally assemble different truncated octahedral geometries [22,29]. The AS1411-NC and AS1411\*-NC structures were modeled using the SYBYL 6.0 program (TRIPOS, http://www.tripos.com, accessed on 30 June 2021), manually adding one copy of the AS1411 aptamer to the octahedral scaffold, using a 7-thymidine spacer to keep the structure accessible for nucleolin targeting. The system topologies and the coordinates of AS1411, AS1411\*, AS1411-NC, and AS1411\*-NC models, used as input for the AMBER 16 MD package [30], were generated through the AmberTools tLeap module, parametrizing the structures through the AMBER ff19SB force field with the parmbsc1 corrections [31]. The structures were immersed in a rectangular box filled with TIP3P water molecules, imposing a minimum distance between the solute and the box of 14 Å, neutralizing the charges adding Mg2+ counterions to the solvated systems in favorable positions, as implemented in the tLeap program, together with 4 K+ ions for G-quadruplex stabilization [32]. All systems were minimized using 2500 steps of the steepest descent algorithm to remove unfavorable interactions and prevent Mg2+ ions from binding to DNA, applying harmonic restraints of 50 kcal·mol−1·Å−2. The systems were then equilibrated to 300 K in the NVT ensemble for 500 ps using the Langevin thermostat, with a coupling coefficient of 1.0 ps and including cartesian restraint of 15 kcal·mol−1·Å−2 on nucleotide atoms. After the equilibration phase, the systems were subjected to an equilibrium simulation of 500 ps removing all constraints and to relax all atoms. The systems were then simulated using the NPT ensemble for a period of 10 ns, applying periodic boundary conditions, a 2.0 fs time-step, and the PME method [33] for the long-range electrostatic interactions with a cutoff of 9.0 Å for the evaluation of short-range nonbonded interactions. The SHAKE algorithm [34] was used to constrain covalent bonds involving hydrogen atoms. The temperature was fixed at 303 K using the Langevin dynamics [35], while pressure was kept constant at 1 atm through the Langevin piston method [36]. Atomic positions were saved every 1000 steps (2.0 ps) for the analyses.

#### *2.9. Gaussian Accelerated Molecular Dynamics Simulations*

For each system, 100 ns dual-boost Gaussian accelerated molecular dynamics (GaMD) simulations [37] were performed, saving the atomic positions every 1000 steps. GaMD is an enhanced sampling method in which a harmonic boost potential is applied to smooth the potential energy surfaces and reduce the systems energy barriers [37]. Using this approach, it is possible to sample a large conformational space, which cannot be normally accessed by classical MD simulations. Moreover, by constructing a boost potential following Gaussian distribution, using cumulant expansion to the second order, GaMD simulations can be correctly reweighted to recover the original biomolecular free energy profiles [37]. GaMD simulations were entirely performed using an NVIDIA TITAN XP GPU using the default parameters suggested for the method (http://miao.compbio.ku.edu/GaMD/, last accessed 30 June 2021).

#### *2.10. Trajectory Analysis*

The GROMACS 2020.3 analysis tools [38] were used to compute the root mean square fluctuations (RMSFs) and principal component analysis (PCA) over the entire 100 ns trajectories. A clustering analysis was performed on all the saved configurations through the gmx cluster module of GROMACS using the GROMOS algorithm [39], forcing a cut-off of 0.12 nm for the geometrical clustering procedure. In order to recover the original freeenergy profiles of the simulated structures, the GaMD trajectories were reweighted using PyReweighting, a toolkit of Python scripts designed to facilitate the GaMD simulation analyses [37]. The UCSF Chimera program [28] was used to generate the pictures of the representative structures.

#### **3. Results**

#### *3.1. Design, Assembly, and In Vitro Stability of AS1411 Functionalized DNA Nanocages*

AS1411 has been shown to adopt multiple conformations characterized by very similar hydrodynamic and electrophoretic properties [7]. A single G-quadruplex conformation, named AT11, can be obtained by mutating a guanine in thymine in the position 10 [4]. The AT11 structure was obtained by NMR spectroscopy, and we started from this PDB structure to build the AS1411 atomic 3D model, introducing the original AS1411 sequence through the 3DNA mutate\_bases module (Supplementary Figure S1) [26]. A second 3D model of the aptamer, called AS1411\*, was generated replacing guanine 10 with guanine 11 in the formation of the first G-quartet of the aptamer (Supplementary Figure S1). A 3D model with one AS1411 or AS1411\* aptamer (Apt) attached to a DNA nanocage (NC) was built to form the Apt-NC structure (Figure 1A). The octahedral nanocage was obtained from 8 oligonucleotide sequences (OL1–OL8, Supplementary Table S1) to form a scaffold made up of 12 double stranded B-DNA helices forming the edges of a covalently closed octahedron, connected by short single-stranded five-thymidine linkers, corresponding to the square truncated faces [21,23,25]. AS1411 aptamer was connected to the nanocage scaffold through a seven-nucleotide linker made by thymidine, using a modified oligonucleotide (OL8Apt) (Supplementary Table S1).

The assembly of AS1411-octahedral nanocages was performed as described [21], using the oligonucleotides reported in Supplementary Table S1. In electrophoresis, the assembled nanocages ran in the gel as a single well-defined product with a molecular weight of about 530 bp (Supplementary Figure S2). As depicted in Figure 1A, Apt-NCs were functionalized with a biotin molecule on one edge of the structure. The presence of biotin enabled detection of the assembled nanocages after transfer from the gel to a membrane, through the streptavidin (HRP)–biotin reaction in the DNA blot, a system with a sensitivity up to 100 times higher than the ethidium bromide staining, as we calculated in a comparative test.

Figure 1B shows the stability of biotinylated Apt-NCs incubated at 37 ◦C in DMEM cell culture medium supplemented with 10% FBS for different time points. After incubation, each sample was treated with proteinase K (100 μg/mL), run in 5% polyacrylamide gel, and analyzed in DNA blot using streptavidin–HRP [21]. Apt-NCs were stable and fully intact for at least 5 h of incubation with serum, then they started to be slowly degraded as a function of time and were still detectable after 48 h (Figure 1B, top panel). In detail, the half-life of Apt-NCs in DMEM–10% FBS was 25 h, calculated by the relative intensity of each band visualized through DNA blot (Figure 1B, bottom panel). Since K+ ions act as stabilizing agents for G-quadruplex structures [40], we tested the in vitro stability of Apt-NCs in serum in the absence or in the presence of 5 mM KCl. DNA blot analysis showed that the presence of only K+ ions increased the resistance of Apt-NCs to serum nucleases, extending their half-life from 16 to 22 h (Supplementary Figure S3). Notably, the presence of both monovalent and bivalent cations in DMEM culture medium guaranteed excellent stability to Apt-NCs (Figure 1B).

**Figure 1.** (**A**) Schematic representation of the Apt-NC nanostructure model. (**B**) DNA blot analysis of Apt-NCs incubated in DMEM supplemented with 10% FBS at 37 ◦C for different times, as indicated. Apt-NCs before incubation with serum proteins are indicated as time 0. Biotinylated Apt-NCs were detected with streptavidin–HRP. The graph shows the densitometric analysis of three different experiments, performed using the ImageJ software. The relative intensity of each band was normalized to the intensity of the band corresponding to time 0 and reported in the graph. Values are expressed as an average ± SEM.

#### *3.2. Intracellular Stability and Targeting Efficiency*

The targeting efficiency and intracellular stability of AS1411-modified NCs were studied by DNA blot in HeLa cells, a human cervical cancer cell line with high surface nucleolin expression [41]. Figure 2 shows HeLa cells incubated with 30 nM biotinylated Apt-NCs or non-functionalized NCs (NC) for 1 and 24 h at 37 ◦C. After incubation, nanostructures were purified from cell lysates and analyzed by DNA blot, using streptavidin–HRP [21]. Lanes 1, 4, 7, and 10 show the band corresponding to DNA nanocages before incubation with cells (time 0). After purification from cell extracts, Apt-NCs ran as one band in the gel, with a mobility comparable to the input (time 0), confirming that we were visualizing only Apt-NCs that were intact inside cells. Interestingly, a similar band intensity was detected at 1 and 24 h of incubation (compare lanes 2–3), suggesting that Apt-NCs were rapidly internalized inside cells, but they did not accumulate in a time-dependent manner. Notably, NCs without aptamer functionalization were almost non-detectable (lanes 5–6), demonstrating that efficient cell uptake occurred only through a nucleolin-mediated AS1411-dependent mechanism. As a control, non-cancer CHO cells, which express very low amount of membrane nucleolin [42], did not show any significant entry of nanocages after 1 and 24 h of incubation with either biotinylated Apt-NCs (lanes 8–9) or non-functionalized NCs (lanes 11–12).

**Figure 2.** DNA blot of Apt-NC (lanes 2–3 and 8–9) and non-functionalized NC (lanes 5–6 and 11–12) purified from HeLa and CHO cell extracts after 1 and 24 h incubation. In lanes 1, 4, 7, and 10, 30 ng of nanocages before incubation with cells (time 0) are shown. Biotinylated nanostructures were detected with the streptavidin (HRP)–biotin reaction.

#### *3.3. Intracellular Trafficking of Apt-NCs*

The membrane binding and cellular internalization of AS1411-functionalized NCs were studied by confocal microscopy in HeLa cells. To distinguish between membranebound and internalized nanostructures, cells were incubated at 4 ◦C or at 37 ◦C, respectively. At 4 ◦C, nanostructures can only enter the cell through direct translocation, because endocytosis, an energy-dependent process, is inhibited at this temperature. Moreover, for visualizing the membrane-bound nanostructures, cells were only fixed, whilst for studying the internalized NCs, cells were incubated for different time periods, then fixed and permeabilized. Representative confocal images of biotinylated-Apt-NC binding at 4 ◦C and uptake at 37 ◦C are shown in Figure 3A. Apt-NCs efficiently bound to HeLa cells and appeared as many small green fluorescent dots on the plasma membrane (Figure 3A, panel a). At 37 ◦C, after 30 minutes, most of the green fluorescent dots were already visible inside cells (Figure 3A, panel b) and remain visible after 4 h of incubation (Figure 3A, panel c), showing a vesicular distribution. In line with DNA blot experiments shown in Figure 2, very low fluorescent signal is observed in HeLa cells treated with NCs (Figure 3A, panel d), confirming that nanocages are entering in cells through a AS1411-dependent mechanism.

The nucleolin-dependent internalization mechanism was clarified by performing a double fluorescence staining of Apt-NCs and Flotillin-1, a lipid raft marker protein which associates with nucleolin in membrane lipid rafts [43]. As shown in Figure 3B (panels a–e), after 30 min of incubation, the green punctate staining of Apt-NCs (panel b) significantly overlapped with the red fluorescence of flotillin-1 (panel c), as confirmed by merged images of green and red fluorescence in panel d. The correlation of the intensity values of the green and red pixels is reported in the scatter plot (panel e) used to calculate the Pearson's coefficient (PCC) [44]. The PCC was 0.47 ± 0.03, confirming that Apt-NCs were internalized through a nucleolin-mediated mechanism involving flotillin endocytosis.

The intracellular distribution was further studied by colocalization analysis of Apt-NCs with the early endosomal antigen (EEA1) after 1 h incubation or the lysosomalassociated membrane protein 1 (LAMP-1) [24] after 4 h incubation. Figure 3B shows representative images of double immunofluorescence of Apt-NCs, shown as green dots inside cells (panels g and l), and EEA1 and LAMP-1 (panels h and m) as red dots. The intracellular distribution was only partially overlapping, as visualized in merged images (panels i and n) and the corresponding scatter plots (panels j and o). The correlation of the intensity values of the green and red pixels evaluated by the Pearson's coefficient (PCC) [44], analyzing the scatter plot, is shown in panels e and j. In detail, the PCC was 0.20 ± 0.02 with the EEA1 marker after 30 min of internalization and 0.27 ± 0.01 with the LAMP-1 marker after 4 h, indicating that Apt-NCs partially trafficked through the endo-lysosomal pathway.

**Figure 3.** (**A**) Confocal analysis of membrane-binding and intracellular distribution of Apt-NCs. HeLa cells were incubated with 60 nM of biotinylated Apt-NCs (panels **a**–**c**) or non-functionalized NCs (panel **d**) for 1 h at 4 ◦C (panel **a**) or for 30 min and 4 h at 37 ◦C (panels **b**–**d**), as indicated. (**B**) Co-localization analysis of Apt-NCs (panels **b**,**g**,**l**) with flotillin-1 (panel **c**), endosomal marker EEA1 (panel **h**), and lysosomal marker LAMP-1 (panel **m**).  Cell nuclei are visualized in (panels **a**,**f**,**k**). The correlation of the intensity values of the green and red pixels was performed using the IMARIS software and scatter plots are reported in (panels **e**,**j**,**o**). Biotinylated NCs were visualized using streptavidin–FITC and nuclei were stained with DAPI. Scale bar: 20 μm.

Guided by these observations, we compared the intracellular distribution of free AS1411 and Apt-NC in HeLa cells and verified whether the presence of a large excess of free AS1411 was able to impair the uptake and the intracellular distribution of Apt-NCs. HeLa cells were incubated with Apt-NCs (Figure 4, panels a and b) or AS1411-Cy5 (Figure 4, panels c and d) for 1 h at 37 ◦C and analyzed by confocal microscopy. Apt-NCs appeared as green fluorescent punctate staining in the cytoplasm (Figure 4, panel b), whilst free AS1411-Cy5 appeared as red dots mostly localized in nucleoli and in the perinuclear region (Figure 4, panel d). It was evident that Apt-NCs and free AS1411 aptamer trafficked inside cells differently and that the AS1411-functionalized nanocages were never found in the nuclei.

**Figure 4.** Comparison of intracellular distribution of Apt-NCs and Cy5-AS1411. Cells were incubated with 60 nM biotinylated Apt-NCs (panels **a**,**b**) or 3 μM free Cy5-AS1411 (panels **c**,**d**) for 1 h at 37 ◦C. (panels **e**,**f**) show cells incubated with Apt-NCs in the presence of 500 molar excess of free AS1411. Biotinylated Apt-NCs were visualized using streptavidin–FITC, and nuclei were stained with DAPI. Scale bar: 20 μm.

Interestingly, incubation of Apt-NCs in the presence of 500 times molar excess of free AS1411 (Figure 4, panel f) did not lead to significant reduction of the green signal (compare panels b and f), as also confirmed by green fluorescence intensity profiles of the confocal images (Supplementary Figure S4), indicating that cell uptake of Apt-NCs was not impaired by the presence of a large excess of free aptamer and demonstrating that the receptor-mediated uptake of AS1411-linked to the nanocages in HeLa cells had a higher efficiency compared with the free aptamer. All together, these results suggest that AS1411-NC functionalization may facilitate AS1411-nucleolin or other membrane proteins interactions, activating a different entry pathway.

#### *3.4. Cytotoxic Effect of Apt-NCs*

The cytotoxic activity of Apt-NCs was tested in comparison with non-functionalized NCs and free AS1411 in cancer HeLa and non-cancer CHO cells. Cells were treated for 24 h with NCs at concentrations ranging from 3.75 nM to 60 nM and with free AS1411 at concentrations ranging from 3.75 nM to 30 μM (Figure 5). Figure 5A shows that incubation of HeLa cells with free AS1411 aptamer was not cytotoxic up to micromolar concentrations. On the other hand, we observed a dose-dependent reduction of cell viability of HeLa cells when treated with Apt-NCs already in the nanomolar range, indicating more than 200-fold increase in cytotoxicity, in comparison with the free aptamer (Figure 5A). No significant reduction in cell viability was observed when treating HeLa cells with NCs not functionalized with the aptamer. As a control, we analyzed the cytotoxic effect of Apt-NCs and free AS1411 on the non-cancer cell line CHO (Figure 5B). No reduction in cell proliferation was observed in CHO cells at all tested concentrations, demonstrating that AS1411 linked to DNA nanocages is not toxic in non-cancer cells. In agreement with this result, free AS1411 induced low cytotoxicity in CHO as in other non-cancer cells [42]. Flow cytofluorimetry was performed for detecting apoptotic cells by a double staining with annexin V–FITC/propidium iodide (PI) (Supplementary Figure S5). In detail, after 24 h

treatment with Apt-NCs the percentage of apoptotic cells reached 33.2 ± 1.2% at 45 nM, while in cells treated with pristine NCs at the same concentration, the percentage was 9.8 ± 1.9%, similar to that observed in untreated control cells (9.9 ± 0.3%). The analysis confirmed the MTS results, showing that Apt-NCs induced a significant dose-dependent apoptosis in HeLa cells while no effect was observed in non-cancer CHO cells.

**Figure 5.** Cytotoxic effect of Apt-NCs in HeLa (**A**) and CHO cells (**B**). Cell proliferation after 24 h treatment with Apt-NCs, non-functionalized NCs, and free AS1411 at different concentrations was assessed by MTS. The values are the means of six replicates normalized to untreated cells. Statistical significance: (\*\*\*) *p* < 0.001 (Student's *t*-test) compared with NCs treated cells in the case of Apt-NCs and untreated cells in the case of AS1411.

#### *3.5. GaMD Simulations*

G-quadruplex-forming sequences are remarkably polymorphic. A single sequence may reach a different 3D structure depending on the physicochemical conditions, such as different folding conditions or presence of a different counterion bound in its core. The high-resolution 3D structure of AS1411 has never been reported due to the high complexity resulting from the simultaneous presence of several conformers that has been demonstrated through multiple spectroscopic techniques [7]. We performed four 100 ns long GaMD simulations, two for each model generated for the free aptamers (AS1411 and AS1411\*) and two for the models of the aptamers linked to the octahedral nanocage (AS1411-NC and AS1411\*-NC), in the presence of potassium ions. Figure 6 shows that the RMSF values, describing the time-averaged deviation of C2 atom positions of the nucleotides, were higher for the free AS1411 (black line) than for the NC-linked aptamer (AS1411-NC) (red line), suggesting a lower conformational space sampling for the latter structure. A similar result was obtained comparing the RMSF of the free AS1411\* and of the AS1411\*-NC models (Supplementary Figure S6). A PCA analysis of the motions, coupling the projection of the first two main motions to the reweighting of the GaMD simulations, was carried out

to recover the original free energy profile of the molecules (Figure 7 and Supplementary Figure S6). In this representation, the conformational space sampled by the structure is directly proportional to the number of points plotted on the graph, while the color indicates the energy of the sampled conformations, whose values increase from blue to red. The free AS1411 and AS1411\* structures (Figure 7A and Supplementary Figure S7A) sampled several metastable conformations, characterized by small energy differences of about 1–2 kcal/mol. On the other hand, in the AS1411-NC and AS1411\*-NC structures, the aptamer sampled a more localized conformational space, confined into low-energy basins (Figure 7B and Supplementary Figure S7B). Indeed, by clustering the different frames obtained from the four trajectories, we found two main representative clusters for the AS1411-NC and AS1411\*-NC (Supplementary Figure S8, green and yellow bars), while the two free aptamer structures were characterized by more than 40 low-populated different clusters (Supplementary Figure S8, red and blue bars), confirming a much lower conformational variability for the aptamer linked to the nanocage scaffold.

**Figure 6.** RMSF values calculated for the C2 atoms of the AS1411 aptamer. Black and red filled circles indicate the values calculated for the free and cage-linked aptamers, respectively.

**Figure 7.** Free energy principal component projection of the free (**A**) and cage-linked (**B**) AS1411 aptamers.

#### **4. Discussion**

Here we report a marked enhancement of the G-quadruplex AS1411 therapeutic efficiency due to its functionalization over octahedral DNA nanocages. Aptamer-linked nanocages (i) are stable both in serum and inside cells, similarly to other previously characterized covalently linked DNA nanostructures [21,22,29], (ii) are selectively and efficiently internalized by cancer cells, and (iii) increase the cytotoxic efficacy by more than two orders of magnitude compared with a free aptamer. At least two main explanations can be conceived for the enhanced AS1411 cytotoxicity observed in cancer cells when linked to DNA nanostructures: (i) a more defined conformation adopted by the cage-linked AS1411 and (ii) a different intracellular entry pathway in cancer cells followed by the cage-linked AS1411 compared with free aptamer.

Analysis of the conformational variability of two 3D models of the free aptamers (AS1411 and AS1411\*) and of the aptamers linked to the octahedral nanocage (AS1411-NC and AS1411\*-NC) through GaMD simulations showed that the free AS1411 or AS1411\* aptamers generated more than 40 conformational clusters, indicating that the free aptamer samples many metastable conformations, characterized by small energy differences. On the other hand, MD simulations of Apt-NC pointed out that the nanocage scaffold constrains the aptamer to adopt a more defined conformation, independently of the starting structure, mainly described by two representative clusters (Supplementary Figure S6). Consequently, the structure of the free aptamer appears to be more floating, as confirmed by the RMSF analyses. These results are likely correlated to the higher efficiency of the cage linked AS1411 in targeting nucleolin and/or other membrane proteins in cancer cells and in improving cell toxicity compared with the free aptamer. Indeed, this may represent an example of structure–function relationship, since the enhanced AS1411 cytotoxicity appears correlated to a confinement of the aptamer in a more defined conformation than that of the free aptamer.

AS1411 has been linked to various types of nanoparticles due to its selective binding to nucleolin, a protein overexpressed on the cell surface and inside cancer cells [45–49]. Notably, a marked improvement of cytotoxicity has been previously obtained by increasing the loading (more than 100 molecules) of AS1411 on the surface of gold nanostars [50]. However, the here described inhibition of cancer cell proliferation by AS1411 linked to NCs (up to two orders of magnitude greater than that by the free aptamer) is the most powerful reported so far.

We demonstrated that the aptamer AS1411 confers a selective targeting activity to Apt-NCs, when compared with pristine nanocages, showing a clear targeting preference for the nucleolin-positive HeLa cancer cells compared with the nucleolin-negative control cells. Apt-NCs are efficiently taken up by cancer cells using a flotillin-dependent endocytosis mechanism, traffic through the endo-lysosomal pathway, and never reach the nuclei, whilst AS1411 traffics both to the cytoplasm and to the nuclei, accumulating in nucleoli. By competition experiments, we also demonstrated that aptamer-linked nanocages display a higher internalization efficiency than free AS1411. The different intracellular traffic and localization may be responsible for the higher cytotoxicity, changing the mechanism of action of AS1411, although further studies are required to definitely confirm it and to identify the pathway.

The proposed AS1411-functionalized DNA nanocages are very stable and, beyond the marked increase of cytotoxicity to cancer cells, present the advantage of being a potential multifunctional scaffold. In fact, the octahedral DNA nanocages can be tailored with sequestering units for selective oncomiR inhibition [25] and/or loaded with DNA intercalating drugs, such as doxorubicin [23,25], and used as multipurpose delivery vehicles.

**Supplementary Materials:** The following are available online at https://www.mdpi.com/article/ 10.3390/pharmaceutics13101671/s1, Table S1: Sequences of the oligos for the assembly of octahedral DNA nanocages, Figure S1: Schematic representation of G-quadruplex folding topology of AS1411 and AS1411\* in K+ solution, Figure S2: Gel electrophoresis analysis of pristine (NCs) and AS1411-functionalized nanocages (Apt-NCs), Figure S3: Stability of Apt-NCs in the absence or presence of K+ ions, Figure S4: Intensity profiles of streptavidin–FITC signal, Figure S5: Annexin V-FITC/propidium iodide assay in HeLa and CHO cells, Figure S6: RMSF values calculated for the C2 atoms of the AS1411\* structures, Figure S7: Free energy principal component projection of the isolated (A) and cage-linked (B) AS1411\* aptamers, Figure S8: Bar chart representing the distribution of the clustered AS1411, AS1411-NC, AS1411\*, and AS1411\*-NC conformations.

**Author Contributions:** G.V., S.R. and V.U. designed and performed experiments, analyzed data, and helped writing the manuscript; F.I. and C.S. designed experiments and analyzed data, A.D. designed experiments and helped writing the manuscript, and S.B. designed experiments, analyzed and interpreted data, and wrote the manuscript. All authors have read and agreed to the published version of the manuscript.

**Funding:** This work was supported by PRIN 2017 "Progetto di Rilevante Interesse Nazionale (PRIN) grant (to S.B.) and Associazione Italiana per la Ricerca sul Cancro FIRC-AIRC fellowship for Italy (to S.R. and F.I.).

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** Not applicable.

**Acknowledgments:** We thank E. Romano from the Centre of Advanced Microscopy "Patrizia Albertano" for the skillful assistance in confocal analysis.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**

## *Article* **Antigen-Capturing Mesoporous Silica Nanoparticles Enhance the Radiation-Induced Abscopal Effect in Murine Hepatocellular Carcinoma Hepa1-6 Models**

**Kyungmi Yang 1,2,†, Changhoon Choi 1,†, Hayeong Cho 3, Won-Gyun Ahn 1, Shin-Yeong Kim 1, Sung-Won Shin 1,2, Yeeun Kim 1, Taekyu Jang 3, Nohyun Lee 3,\* and Hee Chul Park 1,2,\***

**Citation:** Yang, K.; Choi, C.; Cho, H.; Ahn, W.-G.; Kim, S.-Y.; Shin, S.-W.; Kim, Y.; Jang, T.; Lee, N.; Park, H.C. Antigen-Capturing Mesoporous Silica Nanoparticles Enhance the Radiation-Induced Abscopal Effect in Murine Hepatocellular Carcinoma Hepa1-6 Models. *Pharmaceutics* **2021**, *13*, 1811. https://doi.org/10.3390/ pharmaceutics13111811

Academic Editors: Marina Santiago Franco and Yu Seok Youn

Received: 5 October 2021 Accepted: 26 October 2021 Published: 29 October 2021

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**Abstract:** Immunomodulation by radiotherapy (RT) is an emerging strategy for improving cancer immunotherapy. Nanomaterials have been employed as innovative tools for cancer therapy. This study aimed to investigate whether mesoporous silica nanoparticles (MSNs) enhance RT-mediated local tumor control and the abscopal effect by stimulating anti-cancer immunity. Hepa1-6 murine hepatocellular carcinoma syngeneic models and immunophenotyping with flow cytometry were used to evaluate the immune responses. When mice harboring bilateral tumors received 8 Gy of X-rays on a single tumor, the direct injection of MSNs into irradiated tumors enhanced the growth inhibition of irradiated and unirradiated contralateral tumors. MSNs enhanced RT-induced tumor infiltration of cytotoxic T cells on both sides and suppressed RT-enhanced infiltration of regulatory T cells. The administration of MSNs pre-incubated with irradiated cell-conditioned medium enhanced the anti-tumor effect of anti-PD1 compared to the as-synthesized MSNs. Intracellular uptake of MSNs activated JAWS II dendritic cells (DCs), which were consistently observed in DCs in tumor-draining lymph nodes (TDLNs). Our findings suggest that MSNs may capture tumor antigens released after RT, which is followed by DC maturation in TDLNs and infiltration of cytotoxic T cells in tumors, thereby leading to systemic tumor regression. Our results suggest that MSNs can be applied as an adjuvant for in situ cancer vaccines with RT.

**Keywords:** mesoporous silica nanoparticles; radiotherapy; immunotherapy; tumor microenvironment; abscopal effect

#### **1. Introduction**

In recent years, immunotherapy has gained interest as an option in cancer treatment. Immune checkpoint inhibitors targeting cytotoxic T lymphocyte-associated protein 4 (CTLA-4), programmed cell death protein 1 (PD1), and programmed cell death ligand 1 (PD-L1) have been approved by the United States Food and Drug Administration (FDA) and are considered promising systemic therapies based on clinical trials for various types of cancers [1]. Radiotherapy (RT) is one of the major types of cancer treatments and is known to modulate cancer immunity. Preclinical studies and some clinical cases have reported surprising results of the abscopal effect of combining immunotherapy with RT [2]. However, this has not yet become standard treatment in the clinic, and not all patients have experienced this positive outcome. Clinical trials to determine the optimal combination of RT and immunotherapy are ongoing.

Immunotherapy has been tested clinically for hepatocellular carcinoma (HCC), which is the second leading cause of cancer-related deaths worldwide [3–5]. Studies have shown positive results, with objective response rates of up to 20% [5]. Despite these encouraging results, immunotherapy does have clinical limitations, such as a relatively low response rate and acquired resistance [6]. In particular, immune checkpoint inhibitors are known to be less effective for HCC because of the lack of tumor-infiltrating lymphocytes and the immunosuppressive tumor microenvironment [7]. Thus, there is an unmet need to find innovative ways to enhance such treatment effects [8]. RT, one of the strategies to overcome this problem, has been considered as an option to boost the efficacy of immunotherapy for HCC [9]. Clinically, in a study investigating the role of RT in advanced HCC patients treated with nivolumab, patients with previous or concurrent RT showed better clinical outcomes than those without RT [10]. We previously studied a syngeneic murine HCC model and confirmed the abscopal effect and immunological mechanisms of the combination of RT and anti-PD1 antibody [11]. However, the mechanism by which RT stimulates the immune system is not yet clear, and effective immune modulation by RT itself needs to be tested to better understand the synergy between RT and immunotherapy. Researchers have suggested that RT induces immunogenic cell death and that this property could be useful for in situ vaccination [12].

Various types of engineered nanomaterials have been developed for medicinal purposes. Among them, mesoporous silica nanoparticles (MSNs) have attracted growing attention as nanocarriers for drug delivery or antigen targeting for vaccination. Owing to their large surface area and porous structure, MSNs possess the absorbing property of small biomaterials, such as proteins [13]. They effectively deliver absorbed biomaterials to target cells [14,15]. Researchers have made progress in the development of this property of MSNs for cancer treatment. In addition to chemotherapeutic drug delivery, MSNs have been studied for cancer vaccination, which could elicit an anti-cancer immune response by delivering cancer antigens to antigen-presenting cells [16].

In this study, we used a syngeneic murine HCC model to investigate anti-cancer immune stimulation by using MSNs in combination with RT. We tested the hypothesis that MSNs effectively deliver RT-releasing cancer antigens to local immune cells and investigated whether MSNs can enhance local tumor control and the abscopal effect.

#### **2. Materials and Methods**

#### *2.1. Synthesis and Characterization of Nanoparticles*

MSNs were synthesized via the sol-gel reaction of silane agents in the presence of structure-directing agents [17]. We dissolved 3 g of cetyltrimethylammonium chloride (CTAC, 25% solution, 12 mL, Sigma-Aldrich, St. Louis, MO, USA) and 60 mg of triethanolamine (Sigma-Aldrich, St. Louis, MO, USA) in 120 mL of deionized water. After heating to 95 ◦C, 2.25 mL of tetraethyl orthosilicate (TEOS, Acros Organics, Fair Lawn, NJ, USA) was added. After 2 h, the solution was cooled to room temperature, and the nanoparticles were collected by centrifugation at 11,000 rpm for 30 min and washed with ethanol three times. To extract residual structure-directing agents from the pores of MSNs, 1.3 mL of hydrochloric acid was added to the solution, which was refluxed for 3 h. For amine functionalization, 2.8 mL of (3-aminopropyl) triethoxysilane (APTES, 99%, Sigma-Aldrich, St. Louis, MO, USA) was added to the pore-extracted MSN solution and reacted for 3 h at 80 ◦C.

Fluorescently labeled MSNs were prepared by the co-addition of a fluorescence dyeconjugated silane agent with TEOS. The fluorescence dye-conjugated silane agent was synthesized by reacting 5 mg of fluorescein isothiocyanate (FITC, Sigma-Aldrich, St. Louis, MO, USA) with 44 μL of APTES in 1 mL of ethanol for 12 h in the dark. Immediately after the addition of TEOS (2.25 mL) to a solution containing CTAC (12 mL) and triethanolamine (60 mg), 250 μL of pre-conjugated FITC-APTES solution was added. After 2 h, the solution was cooled to room temperature, and the nanoparticles were collected by centrifugation at 11,000 rpm for 30 min and washed with ethanol three times. To extract residual structuredirecting agents from the pores of MSNs, 1.3 mL of hydrochloric acid was added to the solution, which was refluxed for 3 h. For amine functionalization, 2.8 mL of APTES was added and reacted for 3 h at 80 ◦C.

The morphology of the MSNs was analyzed using a transmission electron microscope (JEM-2010, JEOL, Mitaka, Tokyo, Japan) at an accelerating voltage of 200 kV. A transmission electron microscopy (TEM) sample was prepared by dropping a diluted solution containing MSNs on a TEM grid, which was followed by air drying. TEM images were obtained without staining. The hydrodynamic sizes and zeta potentials of MSNs were measured by dynamic light scattering (Zetasizer ZS90, Malvern Instrument, Malvern, Worcestershire, UK). The Brunauer-Emmett-Teller (BET) surface area and pore volume of MSNs were measured by nitrogen (N2) adsorption using a BELSORP-mini II (BEL, Toyonaka, Osaka, Japan).

#### *2.2. Measurement of Antigen-Capturing Capacity*

Fluorescein-conjugated ovalbumin (F-OVA, Invitrogen, Carlsbad, CA, USA) was used as a model antigen to evaluate the antigen-capturing capacity of MSNs. MSNs (1 mg) were incubated with 0.2 mg of F-OVA at 4 ◦C for 24 h. Next, the mixture was centrifuged at 11,000 rpm for 15 min. The F-OVA content was measured using a spectrophotometer (RF-6000, Shimadzu, Nakagyo, Kyoto, Japan). The amount of adsorbed MSNs was calculated from the change in the amount of F-OVA before and after adsorption.

#### *2.3. Cell Culture*

Murine hepatoma Hepa1-6 cells were purchased from the American Type Culture Collection (ATCC, Manassas, VA, USA). Hepa1-6 cells were cultured in Dulbecco's modified Eagle's medium (DMEM, Gibco, Carlsbad, CA, USA) containing 10% fetal bovine serum (FBS, Gibco) at 37 ◦C under a humidified atmosphere of 5% CO2.

#### *2.4. Measurement of Tumor Growth in Mice Co-Treated with MSNs and Radiation*

Five-week-old male C57BL/6 mice were purchased from Orient Bio (Seongnam, Gyeonggi, Korea). All animal procedures were conducted in accordance with the appropriate regulatory standards under the study protocol (ID:<PHONE_NUMBER>1). To observe the abscopal effect, a bilateral syngeneic HCC model was established as previously described [11]. Briefly, Hepa1-6 cells (1 × 106 cells) were injected into the right hind leg of the mice, and the same number of Hepa1-6 cells was injected into the left leg of the same mice 3 days after the first injection. The mice were randomized into four groups (*n* = 10 per group): (i) sham treatment, (ii) radiation treatment, (iii) MSN treatment and (iv) radiation + MSNs. The right leg bearing a Hepa1-6 tumor was given with sham treatment or irradiated with 8 Gy of X-ray 14 days after cell inoculation. Irradiation was performed using a linear accelerator (Varian Medical System, Palo Alto, CA, USA), as previously described [8]. The as-synthesized MSNs (400 μg per tumor) were intratumorally injected into the irradiated tumors twice on days 14 and 15 after cell inoculation. The size of the tumors in both legs was measured every 2–3 days using calipers. The tumor volume was calculated using the formula: volume (mm3) = (W2 × L)/2 (W, width (mm); L, length (mm)), as previously described [11,18]. The mice were sacrificed 42 days after cell inoculation.

#### *2.5. Flow Cytometry Analysis*

Tumors were harvested from both legs of the mice on days 20 and 42 after cell injection. A single-cell suspension was prepared as described previously [11]. Briefly, after the removal of red blood cells, the cells were fixed with Cytofix (BD554655, BD Biosciences, San Jose, CA, USA) for 30 min at 4 ◦C and resuspended in staining buffer (BD554656). The cells were permeabilized using the Fix/Perm kit (00-5523, eBioscience, San Diego, CA, USA). For T cell analysis, cells were stained with anti-Foxp3 (BD560408) and anti-IFNγ (BD557724) antibodies for 30 min at 4 ◦C. After washing, the cells were stained with

antibodies specific for CD4 (BD552051), CD8 (BD560469), CD25 (BD551071), and CD45 (BD559864). Data were acquired using a BD FACSVerse flow cytometer (BD Biosciences) and analyzed using FlowJo software version 10.6.1 (Three Star Inc., Ashland, OR, USA).

For dendritic cell (DC) analysis, inguinal lymph nodes of the irradiated tumor side were harvested four days after irradiation, and DCs were isolated using the EasySep mouse plasmacytoid DC isolation kit (STEMCELL Technologies, Vancouver, BC, Canada). The DCs were stained with anti-CD11c (BD553801), anti-CD80 (BD560016), anti-CD86 (BD560582) and anti-MHC II (BD562363), and subjected to flow cytometry.

#### *2.6. Pre-Incubation of MSNs with Irradiated Cell Conditioned Medium*

Hepa1-6 cells were seeded in a 100 mm dish and irradiated with 100 Gy of γ-rays using an IBL437C blood irradiator (CIS Bio International, Gif-sur-Yvette, Essone, France). After 48 h of incubation without FBS, the conditioned medium (CM) was collected and centrifuged at 1500× *g* for 10 min to remove cell debris. The MSNs were resuspended at a concentration of 2.5 mg/mL in the cell culture supernatant and incubated for 72 h at 4 ◦C. After centrifugation at 11,000 rpm for 15 min, CM-incubated MSNs (CM-MSNs) were collected and washed twice with PBS. CM-MSNs were resuspended in PBS at a final concentration of 8 mg/mL.

#### *2.7. Cellular Uptake of MSNs*

The immortalized immature DC line JAWS II was purchased from ATCC. JAWS II cells were seeded onto a cover slip (Paul Marienfeld GmbH & Co. KG, Lauda-Königshofen, Baden-Württemberg, Germany) and incubated with 250 μg/mL of rhodamine-preloaded MSNs for two days. Cells were fixed with 4% formaldehyde and permeabilized with 0.01% Triton X-100. Cells were stained with Alexa-Fluor488-conjugated phalloidin (Life Technologies, Eugene, OR, USA) and DAPI (Sigma-Aldrich, St. Louis, MO, USA). Fluorescence images were acquired using a Zeiss Observer D1 fluorescence microscope (Carl Zeiss, Oberkochen, Baden-Württemberg, Germany). The activation of JAWS II cells by CM-MSNs was determined by flow cytometry. JAWS II cells (5 × 106 cells) were seeded in 12-well plates and incubated with 100 or 200 μg/mL of CM-MSNs for two days. Cells were stained with anti-CD40 (BD562846), anti-CD80, anti-CD86, anti-MHC I (BD742859), anti-MHC II and anti-PD-L1 (12-5982-82, eBioscience, San Diego, CA, USA). Stained cells were analyzed by a BD FACSVerse flow cytometry.

#### *2.8. Measurement of Tumor Growth in Mice Co-Treated with CM-MSNs and Anti-PD1*

To test the immune-boosting effect of CM-MSNs, a single Hepa1-6 tumor model was established by injecting into the right hind leg of C57BL/6 mice. When the tumors were palpable, the mice were randomly divided into four groups (*n* = 4 per group): (i) MSNs + isotype IgG, (ii) MSNs + anti-PD1, (iii) CM-MSNs + isotype IgG and (iv) CM-MSNs + anti-PD1. Isotype IgG and anti-PD1 (BE0089 and BE0146; Bio X Cell, West Lebanon, NH, USA) were intraperitoneally administered at a dose of 2 mg/kg twice per week. MSNs or CM-MSNs (8 mg/kg) were subcutaneously injected into the left hind leg on the same day as the antibody injection. Tumor size was measured every 2–3 days using calipers, as described above. The mice were sacrificed 28 days after cell inoculation.

#### *2.9. Immunohistochemistry*

Formalin-fixed paraffin-embedded tumor tissue specimens were prepared as described previously [18]. For immunohistochemistry (IHC), the tumors were sliced into 4-μm-thick sections, deparaffinized in xylene, rehydrated in graded alcohol, and washed with 0.01 M PBS, pH 7.4. Heat-induced epitope retrieval was performed using citrate buffer (pH 6.0; Dako, Carpinteria, CA, USA), followed by blocking with a blocking buffer (Dako). The tissue sections were stained with anti-CD4 and CD8 antibodies, followed by incubation with horseradish peroxidase-conjugated secondary antibodies and 3,3 -diaminobenzidine substrate chromogen solution (DAB, Dako). IHC images were obtained using an Aperio

ScanScope AT slide scanner (Leica Biosystems Inc., Buffalo Grove, IL, USA) and analyzed using ImageScope software 12.4.3 (Leica Biosystems, Buffalo Grove, IL, USA).

#### *2.10. Statistical Analysis*

All quantitative data are presented as the mean ± standard deviation (SD) or standard error of the mean (SEM). Statistical analyses were performed using Prism 9.2 (GraphPad Software, San Diego, CA, USA). *p*-values were calculated using one-way analysis of variance (ANOVA), Kruskal-Wallis test, or unpaired two-tailed *t*-test. Statistical significance was set at *p* < 0.05.

#### **3. Results**

#### *3.1. Characterization of MSNs*

MSNs were synthesized via the sol–gel reaction of TEOS in the presence of CTAC as a pore-directing agent. TEM images showed that the MSNs had a spherical mesoporous structure (Figure 1A). The average size of MSNs was 100.82 ± 13.41 nm (Figure 1B). To optimize the antigen-capturing capacity, the surface of the MSNs was modified using APTES. The zeta potential of the strongly negative as-synthesized MSNs changed to a positive value after surface modification, confirming successful amine functionalization (Figure 1C). The hydrodynamic diameter of MSNs measured by dynamic light scattering was 104.22 nm (Figure 1D). The BET surface area and pore size determined via N2 adsorption/desorption experiments were 325.31 m2/g and 4.21 nm, respectively (Figure 1E). The antigen-capturing capacity of MSNs was estimated by the adsorption of F-OVA. Poly(lactic*co*-glycolic acid) (PLGA) was employed as a control because its antigen-capturing capacity was recently reported [19]. After incubation of F-OVA with PLGA and MSNs, only 9% and 7% of F-OVA was adsorbed, respectively (Figure 1D). Surface modification with APTES significantly increased the affinity to F-OVA and 45% of F-OVA was captured by aminefunctionalized MSNs. The interaction between MSNs and F-OVA can be attributed mainly to electrostatic interactions. After the adsorption of F-OVA, the zeta potential and hydrodynamic size of MSN were changed to −11.1 mV and 191 nm, respectively (Figure 1C,D). Owing to the enhanced capturing capacity, subsequent experiments were performed using amine-functionalized MSNs.

#### *3.2. Effects of Intratumorally Injected MSNs on Radiation-Induced Abscopal Tumor Growth*

Given that MSNs with a tumor microenvironment can capture biomaterials released after RT, we tested the effect of intratumoral injection of MSNs on radiation-induced tumor growth inhibition. To observe the abscopal effect, the bilateral Hepa1-6 tumor model was used, as in a previous study [11]. Hepa1-6 cells were injected into both legs 3 days apart. X-rays (8 Gy) were delivered to the primary tumor site in the right hind leg, and MSNs were directly injected into the same primary tumor (Figure 2A). Radiation alone significantly decreased the growth of primary tumors but not secondary tumors (*p* < 0.001; Figure 2B–E). Intratumoral administration of MSNs did not affect the growth of unirradiated primary and secondary tumors, but it significantly decreased the growth of both primary and secondary tumors when injected into irradiated primary tumors (Figure 2B–E) compared with sham treatment. The difference in tumor volume between radiation and radiation plus MSNs was not statistically significant. These data indicate that MSNs in combination with RT inhibited the growth of distal and primary tumors.

**Figure 1.** Synthesis of mesoporous silica nanoparticles (MSNs) with antigen-capturing capacity. (**A**) Transmission electron microscopy image of MSNs and (**B**) diagraph showing size distribution. (**C**) Zeta potentials of MSN, MSN-NH2 and MSN-NH2 after capturing fluorescein-conjugated ovalbumin (F-OVA). (**D**) Hydrodynamic diameter of MSN-NH2 before and after capturing F-OVA. (**E**) N2 absorption/desorption isotherms of MSNs. (Inset: pore size distribution from the adsorption branch). (**F**) Amount of F-OVA captured by poly-lactic-*co*-glycolic acid (PLGA), MSNs and MSN-NH2.

**Figure 2.** Intratumoral administration of MSNs enhances the radiation-induced abscopal effect in a bilateral Hepa1-6 tumor model. (**A**) Schedule of MSN or radiation treatment. (**B**) Growth curves of irradiated primary Hepa1-6 tumors implanted into C57BL/6 mice. (**C**) Growth curves of unirradiated secondary Hepa1-6 tumors in the same mice receiving RT. (**D**,**E**) Comparison of tumor volumes between treatment groups 42 days after tumor inoculation. \*\* *p* < 0.01; \*\*\* *p* < 0.001.

#### *3.3. Effects of Intratumorally Injected MSNs on Tumor-Infiltrating Lymphocytes*

To further understand how MSNs facilitate radiation-induced abscopal effects, we evaluated the immunophenotype of tumor-infiltrating T lymphocytes. Primary and secondary tumors were harvested 6 and 28 days after irradiation (Figure 3A). Flow cytometric analysis revealed that in the tumors harvested on day 6, radiation increased the percentage of CD4+IFN*γ*+ T cells (*p* < 0.05) and CD8+IFN*γ*+ T cells (*p* < 0.05) in the primary tumors but not in the secondary tumors compared to the sham treatment (Figure 3B,C). Injection of MSNs into the irradiated primary tumors, but not the unirradiated tumors, further augmented the percentage of activated T cells, although this increase did not reach statistical significance. Instead, MSNs significantly decreased the number of CD4 + CD25 + Foxp3 + regulatory T cells (Tregs) in primary tumors (Figure 3B). In the tumors harvested on day 28, the number of activated CD4 + or CD8 + T cells significantly increased in the secondary tumors as well as the primary tumors co-treated with MSNs and radiation compared to that in sham or single treatments (Figure 3D,E). This correlates with a greater reduction in tumor size (Figure 2D,E). Radiation increased the number of Tregs in

the unirradiated secondary tumors, which was suppressed by co-treatment with MSNs (Figure 3C,E). These data suggest that MSNs may enhance RT-induced tumor regression at the distal site via modulation of the T cell response.

**Figure 3.** Direct injection of MSNs into irradiated primary tumors increases the number of cytotoxic T cells infiltrating the distal tumors. (**A**) Illustration of the treatment schedule and timing of tumor harvesting. (**B**,**C**) Profiling of activated T cells and Treg cells from primary (**B**) and secondary tumor tissues (**C**) harvested 20 days after tumor inoculation. (**D**,**E**) Profiling of activated T cells and Treg cells from primary (**D**) and secondary tumor tissues (**E**) harvested 42 days after tumor inoculation. \* *p* < 0.05; \*\* *p* < 0.01; \*\*\* *p* < 0.001.

#### *3.4. Effects of MSNs on Dendritic Cell Activation*

It is likely that the systemic anti-tumor effect of MSNs is linked to the activation of DCs residing in lymph nodes. To verify this, we tested whether MSNs could activate DCs in vitro. To recapitulate the in vivo situation, conditioned medium (CM) was prepared from Hepa1-6 cells irradiated with 100 Gy of γ-rays and incubated with MSNs to generate CM-MSNs (Figure 4A). First, to test whether DCs can take up CM-MSNs, immortalized immature dendritic JAWS II cells were incubated with rhodamine-preloaded CM-MSNs. Fluorescence imaging showed an efficient uptake of nanoparticles into the cytoplasm of JAWS II cells without visible toxicity (Figure 4B). Flow cytometry showed that CM-MSNs increased the surface expression of DC maturation markers, including CD86, MHC I and MHC II, in a dose-dependent manner (Figure 4C). CM-MSNs also upregulated PD-L1 expression in JAWS II cells. In contrast, MSNs without preconditioning in medium did not activate JAWS II cells at 200 μg/mL (Figure 4D).

Next, we examined whether MSNs injected into irradiated tumors facilitated DC maturation in tumor-draining lymph nodes (TDLNs) collected from mice. In mice bearing bilateral Hepa1-6 tumors, the right hind legs were exposed to X-rays (8 Gy) and then injected with FITC-preloaded MSNs or PBS. TDLNs were collected 3 days after irradiation and subjected to flow cytometry (Figure 4E). DCs obtained from mice co-treated with radiation, and MSNs showed an increase in the FITC-positive cell population compared to that in mice treated with radiation alone (*p* < 0.001; Figure 4F), suggesting uptake of MSNs by DCs. Injection of MSNs into irradiated tumors increased the expression of DC maturation markers such as CD80, CD86 and MHC II in DCs from TDLNs (Figure 4F), which is consistent with the in vitro results.

#### *3.5. Effects of CM-MSNs on Hepa1-6 Tumor Growth in a Syngeneic Mouse Model*

To test the function of CM-MSNs as a cancer vaccine therapy in vivo, we inoculated Hepa1-6 cells into the right hind legs and then evaluated the combined effect of the nanomaterials and anti-PD1 on tumor growth (Figure 5A). Treatment with CM-MSNs delayed tumor growth compared to treatment with the as-synthesized MSNs (Figure 5B,C). CM-MSNs augmented anti-PD1-mediated tumor growth inhibition to a greater extent than the as-synthesized MSNs (*p* < 0.05; Figure 5B,C). Immunohistochemical analysis of CD4 and CD8 in Hepa1-6 tumor tissues showed that CM-MSNs and anti-PD1 greatly increased the infiltration of both CD4- and CD8-positive cells within the tumor tissues (Figure 5D,E). These data suggest that CM-MSNs could be applied in cancer vaccine therapies to boost cancer immunotherapy.

**Figure 4.** MSNs pre-incubated with the irradiated cell conditioned medium activates dendritic cells in vitro and in vivo. (**A**) Schematic diagram for the preparation of conditioned medium-incubated MSNs (CM-MSNs). (**B**) Uptake of CM-MSNs into JAWS II cells. Representative images of CM-MSNs labeled with rhodamine. (**C**) CM-MSNs activated JAWS II, an immortalized immature dendritic cell line. (**D**) MSNs without preconditioning did not activate JAWS II cells. (**E**) Schedule of timing of harvesting tumor-draining lymph nodes. (**F**) Flow cytometry data showing the MSN-increased population of activated DCs in TDLNs. \* *p* < 0.05; \*\* *p* < 0.01; \*\*\* *p* < 0.001.

**Figure 5.** MSNs pre-incubated with irradiated cell conditioned medium augments the anti-tumor effect of programmed cell death protein 1 (anti-PD1) in a syngeneic hepatocellular carcinoma (HCC) model. (**A**) Schedule of MSN or anti-PD-1 treatment. (**B**) Growth curves of Hepa1-6 tumors implanted into C57BL/6 mice. (**C**) Comparison of tumor volume 27 days after tumor inoculation. (**D**) Representative immunohistochemistry (IHC) images and quantitative data of CD4 staining in Hepa1-6 tumor tissues. (**E**) Representative IHC images and quantitative data of CD8 staining in Hepa1-6 tumor tissues. \* *p* < 0.05; \*\* *p* < 0.01; \*\*\* *p* < 0.001.

#### **4. Discussion**

MSNs have a large surface area and porous structure that aid in absorbing nano-level materials such as drugs, proteins, and DNA fragments [15]. Based on this ability, numerous studies on MSNs have been conducted for various therapeutic purposes, including drug delivery, diagnosis and imaging [13,15]. MSNs have also been explored for cancer vaccination because MSNs that absorb tumor antigens increase the anti-tumor adaptive

immune response [20]. Hong et al. [21] showed that the encapsulation of ovalbumin in MSNs enhanced DC internalization, lysosomal degradation escape, and cross-presentation, resulting in tumor growth inhibition. The efficacy and clearance of MSNs depend on their physical characteristics, such as surface charge, diameter, and pore size [13]. In terms of size, MSNs with diameters of 30–200 nm are efficient for drug loading, and those less than 100 nm are sufficient for lymph node drainage [21]. Larger pore sizes have shown better antigen-presenting functions [22]. In this study, we designed MSNs that can capture antigens released after RT and tested their efficacy in combination with RT. PLGA particles have been shown to improve the abscopal effect by capturing RT-releasing antigens [19]. The measurement of antigen-capturing capacity revealed that the as-synthesized MSNs adsorbed a similar amount of F-OVA as the PLGA particles. However, the amine functionalization of MSNs increased the absorption capacity by more than 6-fold. Since the interaction between MSNs and biomolecules depends on physicochemical properties such as electrostatic interactions, hydrogen bonding, and hydrophobic interactions, we expect that the antigen-capturing capacity can be further improved using various silane agents.

Based on the finding that MSNs efficiently captured F-OVA, we speculated that MSNs directly injected into tumors could absorb biomolecules from irradiated tumor cells, thereby eliciting anti-tumor immunity. In the bilateral Hepa1-6 tumor model, MSNs plus RT showed better regression of both primary and secondary tumors. Since MSNs alone did not show any significant effect on tumor growth, it is likely that the capturing process of RT-released antigens by MSNs is necessary for boosting systemic anti-tumor effects. CM-MSNs were prepared to mimic the MSN-absorbing antigens released during RT treatment in vivo. As expected, CM-MSNs showed better tumor control and boosted the anti-cancer effect of the anti-PD1 antibody compared with the as-synthesized MSNs. The action of CM-MSNs may be related to the increase in the number of CD4+ and CD8+ T lymphocytes infiltrating the tumor tissues, which was further enhanced by co-treatment with anti-PD1. These data suggest that MSNs could work as an effective nanocarrier for RT-released antigens and exert a systemic anti-tumor effect. Finally, we expect that CM-MSNs or a combination of MSNs and RT will make the outcome of immunotherapy much stronger and more sustainable.