DOX inhibitor

Targeted delivery system using silica nanoparticles coated with chitosan and AS1411 for combination therapy of doxorubicin and antimiR-21

Fatemeh Khatami a, Maryam M. Matin a, Noor Mohammad Danesh b, Ahmad Reza Bahrami a,*,Khalil Abnous c,*, Seyed Mohammad Taghdisi d,e,**

A B S T R A C T

Herein, a novel targeted delivery system was developed for intracellular co-delivery of doxorubicin (DOX) as a chemotherapeutic drug, antimiR-21 as an oncogenic antagomiR. In this system, DOX was loaded into meso- porous silica nanoparticles (MSNs) and chitosan was applied to cover the surface of MSNs. AS1411 aptamer as targeting nucleolin and antimiR-21 were electrostatically attached onto the surface of the chitosan-coated MSNs and formed the final nanocomplex (AACS nanocomplex). The study of drug release was based on DOX release under pH 7.4 and 5.5. Cellular toxicity and cellular uptake assessments of AACS nanocomplex were carried out in nucleolin positive (C26, MCF-7, and 4T1) and nucleolin negative (CHO) cell lines using MTT assay and flow cytometry analysis, respectively. Also, Anti-tumor efficacy of AACS nanocomplex was evaluated in C26 tumor- bearing mice. Overall, the results show that the combination therapy of DOX and antimiR-21, using AACS nanocomplex, could combat the cancer cell growth rate.

Keywords: Colorectal cancer Silica nanoparticles Tumor
AS1411 aptamer antimiR-21 Chitosan Doxorubicin

1. Introduction

Colorectal cancer stands as the third most diagnosed cancer with approximately 147,950 new cases and 53,200 deaths were predicted to occur in 2020 in the United States (Siegel et al., 2020). The prevalence of colorectal cancer in Iran at the end of 2020 was estimated to be 11,994 for women and 16,395 for men (Abnous et al., 2018).
Doxorubicin (DOX) as an anthracycline antibiotic is one of the most commonly used anticancer drugs for the treatment of colorectal cancer and is an inductor of programmed cell death. However, because of its non-specificity to cancer tissues, it can cause severe side effects such as mucositis and irreversible cardiomyopathy (Chen et al., 2019; Xia et al., 2020). Targeted therapy is now a safe way to increase the effectiveness of anticancer drugs by a direct attack on tumor cells without affecting normal tissues (Abnous et al., 2018; Chatzisideri, Leonidis, & Sarli, 2018).
MicroRNAs (miRNAs), as single-stranded noncoding RNAs, have a length of 20–24 nucleotides. Oncogenic microRNA-21 (miR-21), located on 17q23.1, is one of the prevalent expression regulators in a wide range of cancers. It is associated with metastasis and tumor progression by targeting tumor suppressor genes (Bautista-Sanchez et al., 2020; Finotti et al., 2019; Tornesello et al., 2020). Antagomir (antimiR-21), a class of small engineered oligonucleotides, is used to silent miR-21 and leads to a decline of proliferation in cancer cells and tumor cell migration (Guo et al., 2017; Soheilifar et al., 2019).
On the other hand, aptamers, with their unique characteristics, are considered as targeting agents and new candidates for the treatment of cancer cells. They are short, chemically prepared, single-stranded oli- gonucleotides that fold into unique 3D structures. They carry intrinsic features, including low immunogenicity, low toxicity, intense and quick tumor tissue penetration and convenience of preparation and subse- quent modification. These are remarkable features that allow aptamers to be used as targeting agents and new candidates for the treatment of cancer cells (Fu & Xiang, 2020). AS1411 aptamer is a synthetic G-rich DNA aptamer (26 nt) that has a high affinity towards nucleolin, a protein which is highly expressed on the surface of several types of cancerous cells but not normal cells (Taghdisi et al., 2018; Yazdian-Robati et al., 2020).
Chitosan (C) is a linear polysaccharide with the chemical formula (C₆H₁₁0₄N)n and scientific name poly-(1,4)-2-amino-2-deoxy-beta-D- glucan. It is closely related to cellulose and is produced by chitin acetylation. Chitosan is a non-toxic polymer with high adsorption capacity. The most important polymer used in drug delivery systems is chitosan. It is a degradable and biocompatible polymer with a high positive charge and useful for targeted delivery of therapeutic agents. Chitosan has the ability to control the release of active agents in nanoparticles. In addi- tion, it is widely accepted that cationic polymers such as chitosan have good properties like endosomal escape ability due to proton uptake by their NH2 group, called proton sponge. Due to the above mentioned characteristics of chitosan, many studies have examined the potential utility of chitosan-modified nanoparticles in siRNA and drug delivery for cancer treatment (Afsharzadeh, Hashemi, Mokhtarzadeh, Abnous, & Ramezani, 2018; Alinejad et al., 2016; Ashrafizadeh et al., 2021; Man- souri, Abnous, Alibolandi, Taghdisi, & Ramezani, 2019; Taghavi, Ramezani, Alibolandi, Abnous, & Taghdisi, 2017).
Mesoporous silica nanoparticles (MSNs) are extensively used in bio- clinical applications as biocompatible nanoparticles (Lin, Zhang, Liu, & Ye, 2018; Zhou et al., 2018). They have low toxicity and also can be easily modified to permit chemical loading and site-specific targeting (AbouAitah et al., 2020; Gisbert-Garzaran & Vallet-Regi, 2020; Narayan, Nayak, Raichur, & Garg, 2018). In the present study, AS1411 and chitosan-modified MSNs were applied for targeted delivery of Dox and antimiR-21 as apoptotic mediators to colorectal and breast cancer cells (AACS nanocomplex). Cationic chitosan, a biodegradable and biocom- patible polymer was also used as a gateway to cover the pores of MSNs (Calzoni et al., 2019; Sung & Kim, 2020). Application of chitosan leads to preventing DOX leakage from MSNs, helps the endosomal escape of the therapeutic agents and induces a pH-responsive targeted delivery system. Previous studies have shown that such polymers can protect pharmaceutical components from enzymatic degradation and provide physicochemical stability (Calzoni et al., 2019).
The antagomiR (antimiR-21) and AS1411 aptamer were attached on the surface of chitosan-coated MSNs. It has been demonstrated that free antimiR-21 is unable to enter the cells because of the electrostatic repulsive force between the negative charge of the cell membrane and the negative charge of antimiR-21. AS1411 also carries negative charge as seen in antimiR-21. Meanwhile a specific 3D structure allows it to be selectively connected to its ligand (nucleolin) on the cells. This aptamer can be internalized into the cancer cells with high levels of nucleolin expression (Reyes-Reyes, Salipur, Shams, Forsthoefel, & Bates, 2015; Zhang, Chen, Chen, Chen, & Wang, 2015). All of these features ensure that the designed targeted delivery system is safe for non-target cells (normal cells). C26, MCF-7 and 4T1 cells (Alinejad et al., 2016; Ashra- fizadeh et al., 2021; Mansouri et al., 2019; Salatin & Jelvehgari, 2017) were selected for the in vitro evaluation of the fabricated delivery system (including MTT assay, fluorescence microscope analysis and flow cytometry), while C26 cells were also applied for in vivo experiments (including mouse weight, tumor size, survival efficacy and fluorescence intensity of organs). These cell lines are known for their high rate of nucleolin expression on their cell surface. CHO cells were also used in our study as non-target cells.

2. Materials and methods

2.1. Materials

The antimiR-21 and AS1411 aptamers were designed and then syn- thesized (Microsynth, Switzerland) (Table S1). Doxorubicin hydrochlo- ride (DOX), (3-aminopropyl) triethoxysilane (APTES, 99%), tetraethyl orthosilicate (TEOS, 98%), dimethylsulfoxide (DMSO), cetyl- (4, 5-Dimethyl-2- thiazolyl)-2, 3-(4,5-Dimethyl-2-thiazolyl)-2,5- diphenyl-2H-tetrazolium bromide (MTT) and phosphate-buffered saline (PBS) were obtained from Sigma-Aldrich (Germany). Sodium hydroxide (NaOH), Ethanol, and hydrochloride acid (HCl) were purchased from Merck (Darmstadt, Germany). 3K and 10K centrifugal devices were provided by PALL (USA). The purity content of the applied reagents was analytical grade. RPMI 1640 medium, fetal bovine serum (FBS), trypsin, and peni- cillin/streptomycin were obtained from Gibco (Darmstadt, Germany).

2.2. Cell lines and culture

Cell lines of C26 (mouse colorectal cancer), MCF-7 (human breast cancer), 4T1 (mouse breast cancer), and CHO (Chinese hamster ovary) were ordered from National Pasteur Cell Bank of Iran. The cells were kept in RPMI 1640 including 1% penicillin/streptomycin and 10% FBS at 37 ◦C in a humid atmosphere with 5% CO2.

2.3. Synthesis of MSNs

MSNs were synthesized in accordance with the method of He et al. (He et al., 2012). A solution containing 100 mg CTAB and 48 mL NaOH (2 M, pH = 11) was prepared and the temperature of the solution reached 80 ◦C. After 30 min, 500 μL of TEOS drops was transferred to the sample. The mixture was then stirred for about 24 h at room tempera- ture. The resulting silica solution was centrifuged and freeze-dried.

2.4. MSNs modifications

2.4.1. Modification of MSNs with NH2 groups

To activate MSNs with NH2 groups, 40 μL of APTES was added to 50 μL of MSN. The reaction continued for another 2 h to reach a white precipitate. The solid product was separated by centrifugation and to remove the surfactant (CTAB), the synthesized nanoparticle was refluxed for 24 h in a solution of HCl (1.00 mL) and methanol (18.00 mL) and then, rinsed repeatedly with deionized water. Next, MSNs-NH2 material was freeze-dried to obtain the dry powder (He et al., 2012; Martinez-Edo, Llinas, Borros, & Sanchez-Garcia, 2019).

2.4.2. Modification of MSNs-NH2 with COOH groups (silica- conjugate)

Succinic acid anhydride was used for the attachment of the carbox- ylic groups on the surface of MSNs-NH2 (Feifel & Lisdat, 2011). 1 g MSN- NH2 (SiO2-NH2) nanoparticles was dissolved in 50 mL anhydrous THF. Then, the suspension was dispersed with ultrasound for 30 min. 3.5 g succinic acid anhydride was added to the suspension and stirred for 2 h at 4 ◦C, followed by stirring for another 24 h at room temperature. 10 mL deionized water was added to the mixture to hydrolyze the residual succinic acid anhydride at room temperature. The resulting product (SiO2-COOH) was dispersed by ultra-sonication for 15 min and centri- fuged for 20 min at 14,000 rpm. The precipitate was then redispersed by anhydrous THF and centrifuged again. Finally, the sediment was dispersed in water and centrifuged for another 20 min at 14,000 rpm. The resulting sediment was freeze-dried.

2.5. Loading of DOX into silica-conjugate

The loading of DOX into the silica-conjugate was performed by treatment of 0–150 μM of DOX with 10 mg Silica-conjugate for 60 min at room temperature. 3K centrifugal device was used to remove the unloaded DOX. The fluorescence spectra of DOX (λEx = 480 nm) was measured with a Synergy H4 microplate reader (BioTeK, USA). The drug loading content and the encapsulation efficiency were evaluated as follows (Yazdian-Robati et al., 2020): (weight of Dox loaded in nanoparticles) trimethylammonium bromide (CTAB), chitosan (CS, low molecular weight, 75–85% deacetylated, Product 448869, 50,000–190,000 Da), 3-

2.6. Capping of DOX-loaded silica-conjugate with chitosan (CS nanocomplex)

To obtain a CS nanocomplex, 2 mg CS was dissolved in PBS (100 μL), and acetic acid (1%) (15 μL) was added dropwise. Then, the mixture was added to 10 mg DOX-loaded silica-conjugate (including 130 μM DOX) in the final volume of 400 μL with PBS. This suspension was incubated for 2 h at room temperature in a dark place.

2.7. Modification of CS nanocomplex with AS1411 aptamer and antimir- 21 (AACS nanocomplex)

The details of the AS1411 aptamer and antimiR-21 sequence is shown in Table S1. 10 μL CS nanocomplex (containing 13 μM DOX), 8 μL AS1411 aptamer (10 μM) and 16 μL antimiR-21 (10 μM) were incubated for 1 h at room temperature in the final volume of 50 μL PBS (10 mM, pH = 7.4). Agarose gel electrophoresis (2.5%) (stained with GelRed, running for 20 min at 80 V) was used for evaluation of the attachment of AS1411 and antimiR-21 to CS nanocomplex.

2.8. In vitro drug release assay

To assay the release profiles of D0X, two pH values of 5.5 and 7.4 were used for simulation of different biological conditions. 400 μL of the AACS nanocomplex was transferred into the microtubes, consisting of a phosphate (pH =7.4) or a citrate buffer (pH =5.5). Then, the samples were put on a shaker (50 rpm) and after certain times (0–96 h), the AACS nanocomplex was detached from the buffer using 3 K centrifugal device. All the measurements were performed three times. The fluorescence reader was utilized to read the concentrations of DOX (λEx = 480 nm, λEm = 600 nm).

2.9. In vitro toxicity assessment (MTT assay)

Dose-increasing studies of DOX showed that IC25 for C26, CHO, MCF- 7, and 4T1 cells were 0.06, 0.15, 0.07, and 0.06 μM (final concentration in the culture medium), respectively. These cells were seeded in 96-well plats (5 × 103 cells/well). After incubation at 37 ◦C for 24 h, the culture medium was substituted by fresh medium consisting of AACS nano- complex (based on the results of the dose-increasing study for DOX), antimiR-21 free nanocomplex (based on the data of dose-increasing study for DOX), AS1411 free nanocomplex (based on the data of dose- increasing study for DOX), free DOX (using the IC25 of DOX), free aptamer, free antimiR-21 and DOX free nanocomplex. After 3 h of in- cubation, the fresh medium was substituted and incubated for further 72 h at 37 ◦C. DMSO and MTT were used to measure light density at 545 and 630 nm on a microplate reader.

2.10. Cellular uptake of the AACS nanocomplex

C26, MCF-7, 4T1, and CHO cells were seeded in 12-well plates (2 × 105 cells/well) and incubated at 37 ◦C for 3 h with 2.5 μM free DOX and AACS nanocomplex (containing 2.5 μM of DOX). Their medium was substituted and the cells were incubated for another 3 h in the fresh medium. The cells were rinsed thrice with PBS (10 mM, pH 7.4), treated with trypsin-EDTA for 1 min, centrifuged at 200g for 5 min, and collected. The fluorescence intensity of the cells was recorded applying a BD Accuri C6 flow cytometer (BD Biosciences, USA) and the FlowJo 7.6.1 software.

2.11. Fluorescence microscopy analysis

C26, MCF-7, 4T1, and CHO cells were seeded in 12-well plates (1 × 105 cells/well) and incubated with 2.5 μM free DOX and the AACS nanocomplex (containing 2.5 μM of DOX) at 37 ◦C for 3 h. The culture medium was refreshed, followed by incubation for 3 h, and an inverted fluorescence microscope (CETI, UK) was used to view the cells (Abnous et al., 2020).

2.12. In vivo toxicity of the AACS nanocomplex

BALB/c mice were purchased from Pasteur Institute, Iran. All animal studies were monitored using protocols approved by the Institutional Ethics Committee and the Research Advisory Committee of Mashhad University of Medical Sciences. The tumor-bearing mice were created by the subcutaneous injection of C26 cells (4 × 105 cells) on the right side of the BALB/c 5–7 weeks old mice. The mice were randomly divided into six groups (n = 4, except PBS n = 3) with a tumor volume of 35–50 mm3. Then, 150 μL of the AACS nanocomplex (1.5 mg/kg DOX), free DOX (1.5 mg/kg DOX), PBS, aptamer free nanocomplex, antimiR-21 free nano- complex and DOX free nanocomplex was injected via the tale vein. The survival rate, tumor size, and body weight measurements were followed up every 2 or 3 days until 21-day post-injection.

2.13. In vivo imaging

C26 cells were injected on the right flank of BALB/c mice (Three groups) via subcutaneous injection (4 × 105 cells). After two weeks, the tumor size increased by 200 mm3. The in vivo binding affinity and the specificity of the AACS nanocomplexes to cellular nucleolin receptors were assayed by the in vivo imaging of C26 tumor-bearing mice, 6 h after the injection of the AACS nanocomplex and free DOX (3 mg/kg). In- jection of PBS through a tail vein mouse was performed for control experiment. For distribution analysis of the injected nanocomplex, free DOX and PBS, the mice were sacrificed and the heart, spleen, liver, kidneys, lung and tumor tissue from each mouse were assayed by the in vivo imaging system (IVIS) (Xenogen, CA; 100 series) (Oroojalian et al., 2018).

2.14. Statistical analysis

Data are shown as means ± standard deviation (SD). The differences between groups were determined using the ANOVA test. In our study, ANOVA test was used for statistical evaluation. Data are reported as means ± SD, n = 5 independent treatments in cancer cells and mice. The effects of the AACS nanocomplex, AACS free aptamer, AACS free anti- miR and AACS free DOX, compared to free DOX with p values < 0.05, were considered as statistically significant. 3. Results 3.1. Synthesis and characterization of MSNs and silica complexes The size of MSNs was assessed as 87 ± 6 nm, measured by a dynamic light scattering (DLS) (Fig. S1). The Zeta potentials of MSNs, MSNs modified with the NH2 group, MSNs modified with the COOH group, MSNs loaded with the DOX, the CS nanocomplex and the AACS nano- complex were 35.9 mV, 6.79 mV, —32.3 mV, —25 mV, 18.5 and 16.5 mV, respectively (Table S2). Fourier transform infrared spectroscopy spectra illustrated a peak at about 1100 cm—1, which corresponds to Si–O–Si bond of MSNs. The peak related to amine-functionalized MSNs (NH2 groups) appeared at about 3410 cm—1, and the peak related to COOH groups appeared at 1638 cm—1 (Naowanon, Chueachot, Klinsrisuk, & Amnuaypanich, 2018) (Fig. 1). Transmission electron microscopy (TEM) (CM120, Philips, Netherlands) was used to evaluate the size of MSNs and CS nano- complex. The results showed that the nanoparticles were well-dispersed with a size of about 75 nm (a) and 130 nm (b) for the MSNs and the CS nanocomplex, respectively (Fig. 2). 3.2. Quantification of DOX loading MSNs have a high capacity as nano-vectors for drug delivery because they have a mesoporous structure and great specific surface area. The encapsulation efficiency for DOX loading was 87.5% and the DOX- loaded was entrapped in silica nanoparticle with loading content of 1.39%. 3.3. Attachment evaluation of the oligonucleotides onto the surface of CS nanocomplex As shown in Fig. S2a, after addition of the AS1411 aptamer and antimiR-21 to CS nanocomplex (AACS nanocomplex formation), its associated band did not migrate on the agarose gel (lane 4), compared to the bands of antimiR-21 (lane 1) and free AS1411 aptamer (lane 2), confirming the successful attachment of the oligonucleotides onto the surface of the CS nanocomplex. 3.4. DOX release from the AACS nanocomplex The in vitro DOX release from AACS nanocomplex was performed in both PBS (pH = 7.4) and sodium citrate buffer (pH = 5.5) at room temperature during 96 h by measuring fluorescence. As shown in Fig. S2b, the cumulative release of the DOX reached 64.28% within 6 h at pH 5.5, while this release was 67.39% within 48 h at pH 7.4. 3.5. Cellular uptake of the AACS nanocomplex Fig. 3 shows the results of the fluorescence microscopic assay. The AACS nanocomplex-treated C26, MCF-7 and 4T1 cells (positive nucle- olin, as target cells) showed strong fluorescence emission compared to the AACS nanocomplex-treated CHO cells (negative nucleolin, non- target cells). Fig. 4 indicates the fluorescence FL2 histograms of C26, MCF-7, 4T1, and CHO cells after 3 h treatments with DOX (2.5 μM) and AACS nanocomplex (2.5 μM DOX). FL2 log fluorescence for C26, MCF-7 and 4T1 (in comparison to the CHO cells after treatment with the AACS nanocomplex) showed stronger fluorescence emission, similar to the results of the fluorescence microscope assay. 3.6. In vitro cytotoxicity assay The MTT method was carried out to assess the toxicity of the AACS nanocomplex for different cells (Fig. 5). The viability of the C26 cells, after the treatments with the AACS nanocomplex, antimiR-21 free nanocomplex, aptamer free nanocomplex, DOX free nanocomplex, free AS1411, free antimiR-21 and free DOX, was 43.64 ± 1.24%, 49.31 ± 2.81%, 52.5 ± 1.3, 94.34 ± 1.91%, 95.56 ± 1.88%, 88.58 ± 1.81% and respectively. The viability of the 4T1 cells after similar treatments was, 85.70 ± 1.5%, and 81.81 ± 4.82%, respectively. The statistical analysis using ANOVA exposed a significant difference in cell viability between target (C26, MCF-7 and 4T1) and non-target (CHO) cells when they were treated with AACS nanocomplex (p < 0.05). It is a sign of targeted entry of the AACS nanocomplex into the target cells compared to the non-target cells, while the free DOX is not able to distinguish between these cells and almost equally enters into other treatments. * represents a significant difference (p < 0.05). both target and non-target cells. 3.7. In vivo survival efficacy The anti-cancer activity of the AACS nanocomplex was assessed by the measurement of the tumor volume in the C26-treated mice (Fig. 6). The tumor volumes following treatment with the AACS nanocomplex, AS1411 free nanocomplex, antimiR-21 free nanocomplex, DOX free nanocomplex, free DOX and PBS (control group) were (after 21 days) 508 ± 249 mm3, 831 ± 208 mm3, 614 ± 287 mm3, 933 ± 232 mm3, 895 ± 199 mm3 and 1285 ± 271 mm3, respectively. One-way ANOVA analysis was performed to compare the tumor sizes using three treatments (AACS nanocomplex, DOX and PBS). The results showed that the tumor size in the AACS nanocomplex-treated group grew significantly slower than those treated with DOX and PBS (p < 0.05). other treatments, a small amount of weight gain was observed. As shown in Fig. 8, it was specified that the intravenous treatments of mice with AACS nanocomplex prolonged their survival rate during 21 days which was longer than the survival rate of mice treated with PBS or other complexes. All mice treated with AACS nanocomplex survived for up to 21 days, while in groups receiving the DOX free complex or PBS, mortality cases were observed. 3.8. Ex vivo imaging analysis The tumor-targeting success of the AACS nanocomplex was assessed by exploring the DOX fluorescent signal to measure the distribution and quantity of AACS nanocomplex and free DOX in tumors and other tis- sues. 6 h after injection, the fluorescent signal of the AACS nanocomplex group was almost close to the free DOX in the tumor, while the accu- mulation of the AACS nanocomplex in liver and kidneys of the mice was much less than that of the free DOX (Fig. 9a). The mean fluorescence intensity in the tumors, liver, kidneys, lung, heart and spleen of the mice after treatment with the AACS nano- complex were 148, 137, 135, 140, 117 and 124, respectively. The mean fluorescence intensity in the same tissues, after treatments with the free DOX, was 166, 166, 175, 134, 122 and 123, respectively. These numbers, after treatments with the PBS, were 119, 119, 119, 117, 116 and 115, respectively (Fig. 9b). 4. Discussion Chemotherapy is among the main medical treatments for cancer. However, the use of these drugs is limited because they cannot differ- entiate between target and non-target cells, leading to severe side effects (Abbas & Rehman, 2018; Mohammad, 2018). Of the chemotherapeutic agents, doxorubicin (DOX) has been widely used against a variety of solid tumors, including clone tumors, while it can cause adverse effects (eg, kidney and liver toxicity) on healthy cells (Siminzar, Omidi, Gol- chin, Aghanejad, & Barar, 2019). Recently, the use of nano-medical methods has been introduced as the preferred approach for treating cancer. So, in this study, a sophis- ticated silica nanostructure was developed for targeted delivery of antimiR-21 and DOX to gain better effects on cancer cells (Bhere et al., 2020; Ganju et al., 2017; Yang et al., 2020). Superficial activation of MSNs can be used to regulate their charac- teristics, such as regulating the response to environmental conditions and the immune system, as well as covering events to control drug release (Devulapally, Sekar, & Paulmurugan, 2015; Ren et al., 2016; Rui et al., 2017). In this research, in order to activate MSNs, the NH2 and COOH groups and the chitosan polymer coating were used to target co- delivery of Dox with antimiR-21 based on the activity of the AS1411 aptamer (Scheme 1). An overview of the chemical synthesis of silica nanoparticles and the steps of silica nanoparticles modification with NH2 and COOH groups is given in Scheme 2, respectively. A polymer in which 100% of its amine groups are acetylated is called chitin, and a polymer without amide groups is called chitosan. Chitosan is degraded over time in biological systems. This property causes drugs to be released in a controlled manner in the body, with increased effectiveness. Free amine groups with positive charges are essential for drug delivery. These positive charges interact with the negatively charged DOX and aptamer molecules. In addition to the above proper- ties, biological properties such as bioadhesion, anti-cancer properties and antimicrobial properties distinguish chitosan from other biological polymers (Ki et al., 2014; Luesakul, Puthong, Neamati, & Muangsin, 2018; Mohammed, Syeda, Wasan, & Wasan, 2017; Richard, Thibault, De Crescenzo, Buschmann, & Lavertu, 2013; Salatin & Jelvehgari, 2017). The TEM image analysis of MSNs and CS nanocomplex displayed that the designed nanoparticles are originally spherical (Fig. 2). The struc- ture of the mesoporous nanoparticle is well defined for silica nano- particles. After addition of CS to MSNs, the size and zeta potential of the nanoparticles were increased, confirming the successful formation of CS nanocomplex using electrostatic interaction. Drug loading was performed as a result of electrostatic interaction between silica and DOX. The loading content and encapsulation effi- ciency of the DOX in the MSNs were 1.39% and 87.5%, respectively. This high loading of DOX is due to the mesoporous structure and so, high capacity of the MSNs. The connection signs of the aptamer and antimir on the surface of MSNs are the result of interaction between the negative charges of the AS1411-antimir 21 and the positive charges of the chitosan on the sur- face of MSN. Agarose gel electrophoresis was used to determine whether the antimiR-21 and AS1411 aptamer could bind to the surface of the CS nanocomplex via electrostatic interaction or not (Fig. S2a). The elec- trophoretic band of the free antimiR-21 proved to carry the shortest sequence compared to other groups. The AACS nanocomplex is trapped in the loading well of the gel due to its heavy weight and the presence of fluorescence light, which is related to the DNA strands of aptamer and antimiR-21. This confirms their connection to the CS nanocomplex and formation of the AACS nanocomplex. The release pattern of DOX from the AACS nanocomplex was eval- uated at pH 7.4 and 5.5 to mimic physiological neutral conditions like blood and acidic cancerous cells environment, respectively. Under acidic conditions, the chitosan coating in the AACS nanocomplex is opened owing to protonation of its NH2 groups. Therefore, DOX is released into the cell. Also, chitosan can induce a proton-sponge effect and help the therapeutic agents to escape from endosome. As shown in the Fig. S2b, in the first hour, only 25.59% of DOX was released from the AACS nanocomplex in pH 5.5, while 12.29% of DOX was released from the AACS nanocomplex in pH 7.4. In the next 6 h, 64.28% of the drug was released in pH 5.5 and 38.10% of the release occurred in pH 7.4, but 67.32% of the release occurred in pH 7.4 after 48 h, confirming the pH- responsive profile of the AACS nanocomplex, especially in the acidic environment (Fig. S2b). Previous reports indicate that the DOX release rate from the AACS in pH 5.5 is higher than other carriers, which shows the superiority of this drug delivery system (Abnous et al., 2018; Ki et al., 2014; Luesakul et al., 2018). The efficient tumor-targeting and tumor accumulation of the AS1411-antimir21 modified Silica nanoparticles (AACS nanocomplex) were confirmed by these results. This shows that the AS1411 modified Silica nanoparticle can enhance tumor cellular uptake. DOX intracel- lular accumulation in various cells was examined applying a fluores- cence microscope. A strong fluorescence was observed in C26, 4T1, and MCF-7 cells treated with AACS nanocomplex but a weak fluorescence was observed in CHO cells treated with AACS nanocomplex (Fig. 3). This difference between target and non-target cells refers to the presence of nucleolin as a target for AS1411 on the surface of these cells. These results also confirmed data from flow cytometry analysis and suggested that AACS nanocomplex can be internalized through the association of AS1411-nucleolin and macropinocytosis in cancer cells (Abnous et al., 2018). Flow cytometry method was also used for quantitative analysis of internalization of the AACS nanocomplex into the cells (Fig. 4). C26, MCF-7 and 4T1 cells, treated with the AACS nanocomplex, showed high fluorescence intensity as well as their treatment with DOX. These target cells had high intracellular uptake capabilities for the AACS nano- complex through attachment of the AS1411 to nucleolin receptor existed on the surface of these cells. However, the intensity of fluorescence in the CHO cells, treated with AACS nanocomplex, was significantly lower than that of the free DOX, owing to the absence of nucleolin as the target of AS1411 on their cell surface. These quantitative results displayed that AS1411 could significantly promote the internalization of AACS nano- complex in the target cells through interaction with nucleolin. The in vitro cell viability was conducted using the MTT assay to evaluate the cytotoxicity of different treatments (Fig. 5). The AACS nanocomplex showed the lowest cell survival rate in the target cells (MCF-7, 4T1 and C26 cells) compared to the other treatments. The AACS nanocomplex induced no remarkable cytotoxicity in non- targeting CHO cells, confirming the major role of the receptor of AS1411 for internalization of the nanocomplex into the cells through macro- pinocytosis. The results of this study showed that free antimiR-21 had negligible effect on the survival rate of the cells due to poor internali- zation into the cells. Moreover, the free AS1411 aptamer did not affect the survival rate of the cells even in the targeted cells, due to its low concentration (μM). Also, the lack of DOX ability to detect nucleolin at the surface of the target cells has made DOX act almost identically on the negative and positive nucleolin cells. There is a significant difference for AACS-treated MCF-7 and 4T1 cells compered to antimiR-21 free nanocomplex-treated MCF-7 and 4T1 cells confirming the importance role of both antimiR-21 and DOX for a better cancer treatment. Furthermore, CHO cells showed lower cell viability after the addition of the free DOX compared to treatment of CHO cells with the DOX-complex (p < 0.05) which is related to the lower internalization and cytotoxicity of the complex in non-target cells. Eventually, the anti-tumor activity of the AACS nanocomplex was investigated using BALB/c mice with colorectal tumors. The Fig. 6, showed that the effect of free DOX treatment on tumor growth is small. While, this effect was recorded relatively high, in the case of treatment with the AACS nanocomplex. It was also found that the effect of AS1411 free nanocomplex on tumor growth was significantly lower than that of the AACS nanocomplex, demonstrating the major part of AS1411 for targeted cancer therapy. AntimiR-21 free nanocomplex could not decrease tumor growth as much as AACS nanocomplex, indicating that the presence of DOX is essential for a better cancer treatment. These results confirm that the AACS nanocomplex, due to the presence of both of DOX and antimiR-21, could be a good candidate in anti-cancer therapy programs. The Fig. 7, shows that the effect of AACS nanocomplex treatment on tumor weight loss was much less than the effect of the free DOX treat- ment (p < 0.05). AS1411 free nanocomplex and antimiR-21 free nano- complex treatment had no effect on tumor weight loss. The normal weight of the mice undergoing AACS nanocomplex treatment confirms its low side effects. The survival rate of the mice treated with the targeted nano- complexes, including AACS nanocomplex and antimiR-21 free nano- complex, was improved relatively. In these treatments the presence of AS1411 aptamer, as targeting agent, reduced DOX toxicity, leading to a significant increase in cellular survival rate compared to other treaments (p < 0.05), attributing to the lower side effects of the AACSnanocomplex (Fig. 8). Finally, in vivo imaging (Fig. 9a) showed that upon the free Dox treatment, the liver and kidneys were saturated with the DOX, while in AACS nanocomplex treatment, the least amount of Dox appeared in these organs (p < 0.05). Fig. 9b indicates that free DOX treatment causes the highest intensity of fluorescent in the kidney, as a sign of its high entry into the kidney, but AACS nanocomplex treatment has a highest intensity of fluorescent in the tumor. This would further confirm the targeting property of the AACS nanocomplex (p < 0.05) (Wang et al., 2020; Zavvar et al., 2020). 5. Conclusion Briefly, a targeted nanocomplex delivery system was presented for combination therapy in cancer treatment. The notion is based on MSNs as carriers, AS1411 aptamer as a targeting agent, and DOX and antimiR- 21 as therapeutic agents. Coating of chitosan on MSNs caused simple binding of aptamer and antimiR-21 to the surface of nanoparticles. Furthermore, chitosan as a gatekeeper plays a very important role in the release of the DOX under acidic conditions of cancer cells. The AACS nanocomplex responded to acidic pH for release of the DOX and facilitated the endosomal escape. The designed targeted delivery system showed better DOX release rate from the AACS in pH 5.5 compared to other systems. 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