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Institute of Smart Biomedical Materials, School of Materials Science and Engineering, Zhejiang Sci-Tech University, Hangzhou, 310018, PR ChinaZhejiang-Mauritius Joint Research Center for Biomaterials and Tissue Engineering, Zhejiang Sci-Tech University, Hangzhou, 310018, PR China
Institute of Smart Biomedical Materials, School of Materials Science and Engineering, Zhejiang Sci-Tech University, Hangzhou, 310018, PR ChinaZhejiang-Mauritius Joint Research Center for Biomaterials and Tissue Engineering, Zhejiang Sci-Tech University, Hangzhou, 310018, PR ChinaSchool of Textile Science and Engineering, Zhejiang Sci-Tech University, Hangzhou, 310018, China
Institute of Smart Biomedical Materials, School of Materials Science and Engineering, Zhejiang Sci-Tech University, Hangzhou, 310018, PR ChinaZhejiang-Mauritius Joint Research Center for Biomaterials and Tissue Engineering, Zhejiang Sci-Tech University, Hangzhou, 310018, PR China
Institute of Smart Biomedical Materials, School of Materials Science and Engineering, Zhejiang Sci-Tech University, Hangzhou, 310018, PR ChinaZhejiang-Mauritius Joint Research Center for Biomaterials and Tissue Engineering, Zhejiang Sci-Tech University, Hangzhou, 310018, PR China
Department of Colorectal Surgery and Oncology, Key Laboratory of Cancer Prevention and Intervention, Ministry of Education, The Second Affiliated Hospital, Zhejiang University School of Medicine, Hangzhou, 310000, PR China
Research Center for Clinical Pharmacy & Key Laboratory for Drug Evaluation and Clinical Research of Zhejiang Province, The First Affiliated Hospital, Zhejiang University School of Medicine, Hangzhou, 310018, PR China
Institute of Smart Biomedical Materials, School of Materials Science and Engineering, Zhejiang Sci-Tech University, Hangzhou, 310018, PR ChinaZhejiang-Mauritius Joint Research Center for Biomaterials and Tissue Engineering, Zhejiang Sci-Tech University, Hangzhou, 310018, PR China
Institute of Smart Biomedical Materials, School of Materials Science and Engineering, Zhejiang Sci-Tech University, Hangzhou, 310018, PR ChinaZhejiang-Mauritius Joint Research Center for Biomaterials and Tissue Engineering, Zhejiang Sci-Tech University, Hangzhou, 310018, PR China
Corresponding author at: Institute of Smart Biomedical Materials, School of Materials Science and Engineering, Zhejiang Sci-Tech University, Hangzhou 310018, PR China.
Institute of Smart Biomedical Materials, School of Materials Science and Engineering, Zhejiang Sci-Tech University, Hangzhou, 310018, PR ChinaZhejiang-Mauritius Joint Research Center for Biomaterials and Tissue Engineering, Zhejiang Sci-Tech University, Hangzhou, 310018, PR China
It is meaningful to find suitable in vitro models for preclinical toxicology and efficacy evaluation of nanodrugs and nanocarriers or drug screening and promoting clinical transformation of nanocarriers. The emergence and development of organoids technology provide a great possibility to achieve this goal. Herein, we constructed an in vitro 3D organoid model to study the inhibitory effect of nanocarriers on colorectal cancer. And designed hydroxyapatite nanoclusters (c-HAP) mediated by polydopamine (PDA) formed under alkaline conditions (pH 9.0), then used c-HAP to load DOX (c-HAP/DOX) as nanocarrier for improved chemotherapy. In vitro, drug release experiments show that c-HAP/DOX has suitable responsive to pH, can be triggered to the facile release of DOX in a slightly acidic environment (pH 6.0), and maintain specific stability in a neutral pH value (7.4) environment. c-HAP/DOX showed an excellent antitumor effect in the two-dimensional (2D) cell model and three-dimensional (3D) patient-derived colon cancer organoids (PDCCOs) model. In addition, c-HAP/DOX can release a sufficient amount of DOX to produce cytotoxicity in a slightly acidic environment, entering efficiently into the colorectal cancer cells caused endocytosis and induced apoptosis. Therefore, organoids can serve as an effective in vitro model to present the structure and function of colorectal cancer tissues and be used to evaluate the efficacy of nanocarriers for tumors.
Colorectal cancer (CRC) is the third most frequent cancer globally, causing millions of death worldwide. Unfortunately, the incidence of CRC has increased among young people in the last decade [
]. Therefore, various nanocarriers (NCs) have been developed to eliminate the toxicity of chemotherapy drugs and improve their tumor targeting. Nevertheless, although a large number of 2D cell model-based researches have demonstrated that NCs could improve the therapeutic effect of chemotherapeutic agents, but the genetic information changes and intera-tumoral heterogeneity of this model have restricted the ability of 2D cell cultures to reflect in vivo effects [
N. Gjorevski, N. Sachs, A. Manfrin, S. Giger, M.E. Bragina, P. Ordóñez-Morán, H. Clevers, M.P.J.N. Lutolf, Designer matrices for intestinal stem cell and organoid culture, 539(2016) 560-564, doi:10.1038/nature20168.
L. Broutier, A. Andersson-Rolf, C.J. Hindley, S.F. Boj, H. Clevers, B.-K. Koo, M.J.N.p. Huch, Culture and establishment of self-renewing human and mouse adult liver and pancreas 3D organoids and their genetic manipulation, 11(2016) 1724-1743, doi:10.1038/nprot.2016.097.
M. Takasato, P.X. Er, H.S. Chiu, B. Maier, G.J. Baillie, C. Ferguson, R.G. Parton, E.J. Wolvetang, M.S. Roost, S.M.J.N. Chuva de Sousa Lopes, Kidney organoids from human iPS cells contain multiple lineages and model human nephrogenesis, 526(2015) 564-568, doi:10.1038/nature15695.
W.R. Karthaus, P.J. Iaquinta, J. Drost, A. Gracanin, R. Van Boxtel, J. Wongvipat, C.M. Dowling, D. Gao, H. Begthel, N.J.C. Sachs, Identification of multipotent luminal progenitor cells in human prostate organoid cultures, 159(2014) 163-175, doi:10.1016/j.cell.2014.08.017.
A.J. Miller, B.R. Dye, D. Ferrer-Torres, D.R. Hill, A.W. Overeem, L.D. Shea, J.R.J.N.p. Spence, Generation of lung organoids from human pluripotent stem cells in vitro, 14(2019) 518-540, doi:10.1038/s41596-018-0104-8.
M.A. Lancaster, M. Renner, C.-A. Martin, D. Wenzel, L.S. Bicknell, M.E. Hurles, T. Homfray, J.M. Penninger, A.P. Jackson, J.A.J.N. Knoblich, Cerebral organoids model human brain development and microcephaly, 501(2013) 373-379, doi:10.1038/nature12517.
]. It has been reported that 3D organoids models allow a better simulation of the tumor microenvironment conditions such as hypoxia and nutrient gradients [
]. Based on the high similarity between patient-derived organoids (PDOs) and original organs in structure, function, and gene expression, PDOs have been widely applied to preclinical drug evaluation, biomarker identification, biological research, and individualized therapy [
]. Moreover, tumor organoids are also known as the ideal "patient surrogate", and developing screening platforms based on human organoids may provide a more cost-effective and precise preclinical setting for drug discovery in the long term [
]. Therefore, PDOs have apparent advantages in the efficacy evaluation of NCs.
It has been demonstrated that an ideal drug carrier should physically or chemically incorporate with potential bioactive agents and protect them in the bloodstream. Furthermore, the carrier complex should be degradable gradually and provide sustained drug release over a prolonged period to increase therapeutic efficiency [
]. Hydroxyapatite (HAP), as the main component of hard tissues such as human bones and teeth, has drawn researchers' attention for various biomedical applications, ranging from regenerative medicine [
] because of its high biocompatibility and bioactivity. HAP nanocarriers (HAP NCs) can be used to deliver drugs, proteins, and genes. Compared to other NCs such as carbon nanotubes, silica nanoparticles, and quantum dots, HAP NCs can participate in the normal metabolism of organisms [
Z.-S. Liu, S.-L. Tang, Z.-L.J.W.J.o.G. Ai, Effects of hydroxyapatite nanoparticles on proliferation and apoptosis of human hepatoma BEL-7402 cells, 9(2003) 1968, doi:10.3748/wjg.v9.i9.1968.
]. However, the optimization of HAP NCs for clinical applications, including large-scale preparation, systemic toxicity evaluation, pharmacokinetic and pharmacodynamic studies, as well as study of the efficacy is still a research topic worthy of great effort [
]. Moreover, PDA is abundant in catechol/quinone moieties as well as imine, which endows PDA surface modification with the capability to anchor drugs, peptides or proteins onto the nanoparticles either by physical binding (π-π stacking or hydrogen binding) or chemical binding (Michael addition or Schiff base reactions), and thereby improving the drug loading efficiency [
S.a. Physicochemical, E. Aspects, Nisin-loaded polydopamine/hydroxyapatite composites: Biomimetic synthesis, and in vitro bioactivity and antibacterial activity evaluations.
]. However, although there are many reports on the mineralization of HAP on the surface of materials induced by PDA, few studies focused on the PDA-mediated synthesis of HAP nanoparticles and explored the improvement of drug loading capacity of HAP [
S.a. Physicochemical, E. Aspects, Nisin-loaded polydopamine/hydroxyapatite composites: Biomimetic synthesis, and in vitro bioactivity and antibacterial activity evaluations.
K. Chen, K. Xie, Q. Long, L. Deng, Z. Fu, H. Xiao, L.J.R.A. Xie, Fabrication of core–shell Ag@ pDA@ HAp nanoparticles with the ability for controlled release of Ag+ and superior hemocompatibility, 7(2017) 29368-29377, doi:10.1039/C7RA03494F.
Controlled-temperature photothermal and oxidative bacteria killing and acceleration of wound healing by polydopamine-assisted Au-hydroxyapatite nanorods.
In this study, PDCCOs model was successfully constructed in vitro to evaluate the therapeutic efficacy of HAP NCs, which were loaded with chemotherapy drug doxorubicin hydrochloride (DOX). HAP nanoclusters (c-HAP) as NCs were synthesized by polydopamine (PDA) and used as a pH-responsive nanocarrier to load DOX for tumor chemotherapy. PDA not only guides the formation of a cluster of HAP, but also modifies the surface potential of the material and combines it with DOX. Meanwhile, the synthesized c-HAP/DOX could respond to the slightly acidic environment and achieve pH-triggered DOX release at the tumor site. The PDCCOs model showed that c-HAP/DOX had significant antitumor effects.
2. Materials and methods
Doxorubicin hydrochloride (DOX) was purchased from Sangon (Shanghai, China). Calcium chloride (CaCl2, 96%), disodium hydrogen phosphate (Na2HPO4, 99%) were obtained from Titan (Shanghai, China). Dopamine hydrochloride and 2-Amino-2-(hydroxymethyl)-1,3-propanediol (Tris) were obtained from Biofroxx (Guangzhou, China). FITC was purchased from Yeasen (Shanghai, China). Bovine serum albumin (BSA), 4′,6-diamidino-2-phenylindole (DAPI), 70 µm cell screen mesh, Live/Dead Viability/Cytotoxicity Kit (Promega), Advanced DMEM/F12 (Gibco), B-27 Supplement(50X), N2-supplement (100X), N-acetyl-L-cysteine, Recombinant Mouse EGF were purchased from Gibco (USA). Puromycin, Zeocin, Plasmocin, trypsin-ethylenediaminetetraacetic acid (trypsin–EDTA) were gained from Invitrogen (USA). Matrigel Matrix was obtained from Corning (USA). CS10 was obtained from Stem Cell (Canada). CellTiter-Glo 3D Cell Viability Assay was purchased from Promega (USA). Rhodamine-phalloidin was purchased from Thermo Fisher (USA).
2.1 Preparation of c-HAP
Briefly, anhydrous CaCl2 (110 mg), dopamine hydrochloride (40 mg), and Tris (60 mg) were dissolved in 30 mL deionized water and stirred for 30 min. It was discovered that the color of the solution changed from colorless to orange and then to black. In the next step, 30 mL (5 mM) Na2HPO4 was added to the solution. The mixture was quickly transferred to the reactor at 120 °C for 6 h, cooled at room temperature, and centrifuged at 10000 rpm for 10 min. Pellet was washed with water and anhydrous alcohol three times. Finally, the obtained c-HAP was stored in ethanol and kept at room temperature for the following use.
2.2 Preparation of c-HAP/FITC
About 20 mg of c-HAP was dissolved in 20 mL of anhydrous ethanol under stirring at 80 °C. Subsequently, 0.025 g of FITC was added, and the reaction was kept for 6 h. The synthesized particles were washed several cycles with anhydrous ethanol and deionized water until the centrifugation supernatant became transparent. Finally, the labeled nanoparticles (c-HAP/FITC) were freeze-dried for 24 h.
2.3 Drug loading and release
For the DOX loading test, different concentrations of DOX (0−60 µg/mL) were dissolved in 5 mL of phosphate-buffered solution (PBS) containing c-HAP (2 mg). The solution was incubated for 24 h at 37 °C to load DOX onto c-HAP. To remove the uncoated DOX, the mixture was centrifuged at 10000 rpm for 5 min, and the pellet was washed three times with PBS. The first supernatant was collected to evaluate nanoparticles drug-loading efficiency by UV-vis spectrophotometer. Loading capacity (LC) was calculated by the obtained standard curve. The DOX LC (η) was calculated by this equation:
The loading efficiency was calculated using the following expression:
Where ADOX is the absorbance of original DOX solution at UV-Vis (480 nm). AC and AW are the absorbance of the supernatant solution after centrifugation and the absorbance of DOX molecules recovered after washing of the c-HAP/DOX, respectively.
The pH-triggered release profiles of DOX from c-HAP/DOX were prepared in PBS (pH 7.4, pH 6.5 and pH 6.0) at 37 °C under mechanical shaking (110 RPM). The c-HAP/DOX (2 mg of c-HAP/DOX containing 140.5 µg of DOX) was dispersed in 2 mL of PBS with different pH (6.0 and 7.4) and analyzed in 5.0 mL of the same pH of PBS (5 mL) in a dialysis bag (MWCO: 10 kDa). The DOX release was monitored by UV-vis spectrometer. Each experiment was repeated three times, and the standard deviation (SD) was calculated.
2.4 Characterization
X-ray diffraction (XRD) was performed by a D8 Focus diffractometer (Bruker) equipped with a Cu Kα radiation source (40 kV, 200 mA, l = 0.154 nm). An AVATAR 360 spectrometer was used for Fourier transform infrared (FT-IR) spectra recording. A TGS-2 thermogravimetric analyzer (PerkinElmer) was used for thermogravimetric analysis (TGA). To record fluorescence spectra, RF-5301PC spectro-fluorophotometer (Shimadzu) was used. Transmission electron microscopy (TEM) and energy dispersive analysis (EDS) experiments were carried out on a JEM-2100 (JEOL, Japan) instrument operating at 200 kV. Dynamic light scattering (DLS) and ζ-potential results were determined on a Zetasizer Nanoseries (Nano-ZS, Malvern Instruments). Flow cytometry was carried out using a NovoCyte Flow cytometer (Agilent, USA).
2.5 Cell culture and cell viability assay
Human colon cancer cells (SW480) were cultured in Dulbecco's Modified Eagle Medium (DMEM) growth medium, with 10% FBS at 37 °C under a humidified atmosphere containing 5% CO2. CCK-8 assay was used to evaluate c-HAP, DOX, and c-HAP/DOX toxicity on SW480 cells. To this aim, 1 × 104 cells/well in a 96-well plate were incubated for 12h and then treated by different concentrations of c-HAP, DOX, and c-HAP/DOX. To assess cell viability teat, CCK-8 reagent (Dojindo, Japan) was added into each well and incubated at 37 °C in the darkness for 1 h. The absorbance was measured at 450 nm using the iMark microplate reader. Data expressed as means ± S.E.M. (n = 3 independent experiments in triplicate).
2.6 In vitro intracellular uptake assay
SW480 cells were cultured in a 2 cm dish (2 × 105 cells/dish) at 37 °C with 5% CO2 for 12 h and then treated with 7 µg/mL DOX and 100 µg/mL c-HAP/DOX for 6 h at various pH (6.0 and 7.4). Then cells were washed with cold PBS and fixed with 4% paraformaldehyde (PFA) solution for 30 min. Cells nuclei were stained by 4′,6-Diamidino-2-phenylindole (DAPI), and a scanning confocal microscope (A1, Nikon, Japan) was used to visualize the presence of DOX (red color) and nuclei (blue color). To evaluate cellular uptake, cells were treated with c-HAP/FITC (dispersed in DMEM) into the cells and incubated for 6 h or 12 h. Then treated cells were fixed with 4% PFA solution for 30 min and stained by DAPI and cell membrane red fluorescent probe 1,1′-dioctadecyl-3,3,3′,3′-tetramethylindocarbocyanine perchlorate (Dil).
2.7 Apoptosis study by flow cytometry
Flow cytometry was used to determine the percentage of apoptotic cells. The SW480 cells were plated in a 6-well plate (4.5 × 105 cells per well) and cultivated overnight. Then treated with 0, 100, and 200 µg/mL of c-HAP/DOX for 24 h, followed by washing and suspending in a binding buffer. The staining of these cells was carried out with Annexin V-FITC and propidium iodide (PI) for 15 min in the darkness at room temperature. These samples were quantified by flow cytometry (FACS Calibar) using Cell Quest software (Novoexpress).
2.8 Construction and culture of human colon cancer organoid model in vitro
Patient tissue samples were obtained from the Second Affiliated Hospital of Zhejiang University, and then the colon cancer organoids were extracted from the colon tumor tissue from the patient through shredding, digestion (45 min) and cell filter (100 µm) screening, embedded in matrix glue and cultured to a certain size in complete medium for subculture or experiment. Tumors were de-identified in accordance with the protocols approved by the Institutional Review Board (IRB) of the Second Affiliated Hospital of Zhejiang University School of Medicine.
2.9 Organoid viability assay
In order to investigate organoids viability, 100 organoids were cultured in 48 well/plate with 500 µL of organoid complete medium plus 50 µL of matrigel and different concentrations of c-HAP (0 µg/mL to 125 µg/mL). After 48 h, the culture medium was removed from the wells and 50 µL of CellTiter-Glo®3D reagent was added. The matrigel and CellTiter-Glo®3D reagent mixture was pipetted up and down several times before shaking vigorously for 15 min on a plate shaker. After incubating the plate at room temperature for 25 min, the contents of each plate were transferred to an opaque walled plate to measure the luminescence using a microplate reader. The viability of organoids was calculated using the luminescence value. Data were expressed as means ± S.E.M. (n = 3 independent experiments in triplicate).
2.10 Organoid live/dead & cytoskeleton staining
For the qualitative assay, the organoids toxicity was analyzed using a live/dead viability kit. Briefly, after 48 h treatment with c-HAP, c-HAP/DOX (0 µg/mL to 200 µg/mL), or free DOX, cells and organoids were washed with PBS, followed by the addition of 2 mM Calcein AM and 4 mM ethidium homodimer (EthD-1). After incubation for 30 min at 37 °C, the cells and organoids were observed under a confocal microscope (A1, Nikon, Japan), with excitation and emission of green (ex/em 494/530 nm for Calcein AM) and red (ex/em 528/645 nm for EthD-1) fluorescence. Similarly, the cytoskeleton of organoids was stained by phalloidin. The organoids were incubated for 48 hours were and cleaned with PBS for 3 times, then fixed in the incubator with 4% paraformaldehyde for 30 min. After PBS cleaning for 3 times, the cytoskeleton and nucleus were stained with phalloidin and DAPI. In the process of changing dyes, PBS cleaning was required 2 or 3 times.
3. Results and discussion
3.1 Preparation and characterization of c-HAP
A Schematic diagram of the c-HAP synthesis processes has been shown in Fig. 1A. PDA can chelate many metal ions to coordination sites within the polycatechol frameworks, such as Fe3+, Mn2+, and Ca2+ [
Synthesis of hollow biomineralized CaCO3-polydopamine nanoparticles for multimodal imaging-guided cancer photodynamic therapy with reduced skin photosensitivity.
]. In addition, PDA has a strong adhesion effect and can attach to almost all material surfaces, including metals, ceramics, semiconductors, and synthetic polymers [
], and thus modify the surface of those materials. In the present study, dopamine hydrochloride was used to synthesize PDA under alkaline conditions (pH 9.0). Meanwhile, PDA can chelate the Ca2+ in the solution. After a certain time, an appropriate concentration of PO43− was added to the solution to react with the Ca2+, which was chelated by polydopamine and formed calcium phosphate outside the polydopamine template. At high temperature, calcium phosphate crystals continued to nucleate and grow and finally formed hydroxyapatite nanoclusters. At the same time, because the system contains excess polydopamine, which did not participate in the reaction, it will adhere to the surface of c-HAP.
Fig. 1Preparation steps and morphologies of c-HAP. (A) synthesis of c-HAP particles. (B) SEM, (C) TEM (inset) images of c-HAP prepared under identical conditions, (D) EDS spectrum.
The key point in targeted drug delivery of tumors with slightly larger nanocarriers (diameter between 100 nm to 2 µm) is that they can be recruited at the tumor site through EPR effect [
Employment of enhanced permeability and retention effect (EPR): Nanoparticle-based precision tools for targeting of therapeutic and diagnostic agent in cancer.
]. According to the Fig. 1B and C, c-HAP has a chrysanthemum-like cluster structure with a diameter of about 250 nm. Moreover, from the insert TEM image of Fig. 2B and C, the morphology of c-HAP is coarser than ordinary spherical or rodlike HAP, and it means that c-HAP has a higher drug loading capacity. In order to investigate whether PDA could control the synthesis of c-HAP, we prepared HAP particles in the same reaction conditions (Fig S1). According to the results, HAP synthesis in the absence of PDA causes to form HAP short nano-rod. Moreover, the TEM image of the Energy Dispersive Spectrometer (EDS) spectra of c-HAP showed characteristic X-ray energies for Ca, O, and P, and the ratio of Ca/P was measured 1.26, implying the molecular structure of c-HAP was similar to hydroxyapatite (Fig. 1D).
Fig. 2Characterization of c-HAP. (A) XRD patterns (Vertical lines represent JCPDS No.090432) and (B) FTIR spectra of HAP and c-HAP. (C) Size distribution of c-HAP. (D) TG curves of HAP and c-HAP.
The wide-angle X-ray diffraction (XRD) analysis of HAP and c-HAP was also performed (Fig. 2A). The XRD results show that c-HAP has a typical hydroxyapatite crystal form with low crystallinity, the characteristic peaks consistent with standard hydroxyapatite (JCPDS, No.090432), at 2θ regions of 26°, 32°, 40°, 47°, and 50°. The absorption bands of OH−1 ions (3430 cm−1) and PO43− groups (567, 604, and 1035 cm−1) of the apatitic structure appeared in the FT-IR spectra of c-HAP (Fig. 2B). Moreover, there was a strong vibrational absorption peak at 2361 cm−1 in c-HAP, which could attribute to electrostatic interactions of amine groups in PDA with phosphoric units of c-HAP nanostructure [
]. Compared with c-HAP, a new characteristic peak of -CH2- (2850 cm−1) emerged and the intensity at 3430, 1035, 604 and 567 cm−1 remarkably decreased, and the peak at 2361 cm−1 disappeared in the spectrum of c-HAP/DOX, further confirmed the loading of DOX [
]. DLS results revealed that the hydrodynamic size of c-HAP (249.9 nm) is consistent with the results observed in the TEM images, and the size distribution was uniform (Fig. 2C). The TGA results (Fig. 2D) indicated that the loading content of PDA and DOX was about 8.77% and 6.92%, respectively. Moreover, the weight loss of c-HAP/DOX became rapidly at about 200-400 °C, which is mainly due to the pyrolysis of DOX. The FTIR spectrum and TG curve of c-HAP confirmed that PDA was abundant in the structure of c-HAP.
3.2 Drug loading and releasing of c-HAP
It was demonstrated that PDA with the surface of sp2-bonded carbon nanostructures had the ability to bind to DOX via π-π stacking [
]. To test the drug delivery ability of the c-HAP, DOX was loaded as a guest molecule by soaking c-HAP with DOX in PBS (pH 7.4) overnight. According to the c-HAP LC amount (Fig.S2), the maximum drug loading capacity is about 70.25 µg/mg, and the loading efficiency was about 70%, which was higher than some studies of loading DOX on HAP surface by crosslinking [
]. This result shows that the rough surface structure and π-π stacking between PDA and DOX endow c-HAP with a higher drug loading efficiency. The surface zeta potentials of HAP, c-HAP, and c-HAP/DOX are shown in Fig. 3A. After dispersed in the deionized water, the surface zeta potentials of HAP and c-HAP were changed from -8.8 mV to -12.8 mV. According to previous studies [
], due to the functional catechol groups, this zeta potential change was attributed to the incomplete reaction of PDA covered the particle surface. After DOX loading, the potentials of c-HAP were increased to -9.8 mV. Because of the excessive PDA, DOX was enabled to be absorbed on the surface of c-HAP. In the UV-Vis spectrum of the c-HAP/DOX (Fig. 3B), the DOX peak was observed at 488 nm. Since the c-HAP/DOX suspension is a reddish color, it can also prove that there is DOX load on the surface of c-HAP compared with the DOX solution with the same concentration. In addition, the fluorescence spectra of DOX were obviously quenched after loading into c-HAP, this might be caused by the π–π stacking interactions from energy transfer and/or electron transfer, giving rise to the quenching of DOX fluorescence [
]. These results demonstrated that DOX molecules have been successfully loaded onto c-HAP nanoparticles. To test the drug release profile of c-HAP/DOX, PBS buffers with different pH were used. As shown in Fig. 3C, the release amount of DOX at pH 7.4 was about 15% during 48 h, indicating the high stability of c-HAP/DOX in biological media. Meanwhile, a burst release of DOX molecules was noticed upon exposure to a lower pH environment, the DOX release rate increasing from about 51% at pH 6.5 to approximately 77% at pH 6.0 within 72 h, which may be attributed to the destruction of intermolecular forces between PDA and DOX at the acidic environment. TEM images also suggest that c-HAP particles were enabled to be degraded slowly at pH 6.0 (Fig. 3D). These results could confirm that a slightly acidic environment can trigger the rapid drug release of c-HAP/DOX.
Fig. 3Drug loading/releasing of c-HAP. (A) Zeta potentials of HAP, c-HAP, and c-HAP/DOX. (B) UV-vis-NIR absorbance spectra of (a) c-HAP, (b) DOX, and (c) c-HAP/DOX. (C) DOX release from c-HAP/DOX overtime in PBS buffers at the different pH values (6.0 and 7.4). (D) TEM images of c-HAP after incubation in PBS with different pH values (6.0 and 7.4) for 24 h.
The effects of the nanocarriers on cell viability were evaluated by CCK8 assay. According to Fig. 4A, c-HAP had no significant toxicity on SW480 cells in the concentration range of 0-125 µg/mg, while DOX or c-HAP/DOX significantly decreased cell viability in a dose-dependent manner. Compared with free DOX, c-HAP/DOX showed relatively higher toxicity in the concentration range of 0 to 125 µg/mg, it might be attributed to more DOX capacity and released by c-HAP into the cells.
Fig. 4In vitro pH-responsive DOX release, cellular uptake, and cytotoxicity of c-HAP/DOX. (A) Viability of SW480 cells after incubate with c-HAP, c-HAP/DOX and DOX at different doses for 24 h, and corresponding concentration of DOX was 0, 1.75, 3.5, 5.25, 7.0, 8.75 µg/mL, respectively (n = 4). *p < 0.05, **p < 0.01, ***p < 0.001, and“ns” indicated no statistical difference. (B) CLSM images for SW480 cells incubated with c-HAP/DOX at different pH values (6.0 and 7.4) and free DOX for 6 h. The concentration of free DOX was 7 µg/mL. Scale bars: 50 µm. (C) Accumulation of c-HAP/FITC in SW480 cells after 6 h and 12 h of incubation. Scale bars: 20µm.
The responsive release of c-HAP/DOX under a slightly acidic environment was further verified by confocal laser scanning microscope (CLSM), and the uptake and distribution of DOX in SW480 cells under different pH values were detected. As shown in Fig. 4B, pH values of the culture medium were adjusted to 7.4 and 6.0. Compared with the neutral medium environment, a stronger DOX fluorescence was observed in the cytoplasm of SW480 cells after incubation for 6 h under pH 6.0, indicating that c-HAP/DOX can achieve faster drug release in the slightly acid environment. DOX was easier to enter and accumulate in the nucleus at the same pH conditions compared with c-HAP/DOX after SW480 cells treatment for 6h. In addition, with the extension of time, c-HAP/DOX could continuously release DOX to inhibit the growth of SW480 cells (Fig.S3). All these results show that c-HAP/DOX can have significant sensitivity to the slightly acidic environment of the tumor and then rapid-release drugs at tumor sites.
It has been shown that the therapy-induced inflammation might cause severe adverse effects, including tumor regeneration, metastatic dissemination, and therapeutic resistance [
]. Therefore, we used FITC as a marker to track c-HAP (c-HAP/FITC). According to Fig. 4C, the green fluorescence was continuously increasing in the cell membrane, which shows c-HAP particles were enabled to be swallowed and degraded by cells. As mentioned in the previous report, the degradation of calcium phosphate often produces many free Ca2+, and the accumulation of excessive Ca2+ in cells causes calcium overload, resulting in cell death [
]. Although this process was insignificant to inhibit the growth of SW480 cells, it has certain enlightenment for further research and development of cell calcium death and chemotherapy synergistic therapy.
DOX and other chemotherapeutic drugs mainly kill cancer cells by apoptosis induction. To quantify the apoptotic effect of c-HAP/DOX on SW480 cells, we used annexin-V/PI standing assay kit. As shown in Fig. 5A and B, SW480 in control group showed oval nuclei morphology with distinct edge, indicating that SW480 cells were healthy. The percentage of apoptotic cells in treated groups was significantly higher than control, which increased from 10.72% (control) to 24.38% and 36.72% after treating SW480 with c-HAP/DOX of 100 µg/mL and 200 µg/mL. Moreover, the necrosis rate of SW480 cells increased from 0.13% of control to 23.56% at 200 µg/mL, indicating that when c-HAP used as DOX carrier, it may accelerate apoptosis after activated by lysosomes in tumor weak acidic environment or ingested by tumor cells. The flow cytometry analysis showed that apoptosis activity was enhanced by increasing the incubation concentration of the c-HAP/DOX.
Fig. 5Determination of c-HAP/DOX apoptotic effects: (A) the cell apoptosis plots. (B) Cells percentages in different stages: viable, early apoptosis, late apoptosis, and necrosis.
]. Therefore, we constructed an in vitro model of patient-derived colon cancer organoids (PDCCOs) to further evaluate the therapeutic effect of c-HAP/DOX on colon cancer. The growth status of the extracted PDCCOs after 0 to 5 days of culture has been shown in Fig. 6A. It was observed that the volume of PDCCO is also expanding with the extension of culture time and presents the structure of 3D cell spherical clusters. The organization structure and status of proliferation marker (Ki-67) and Colon cancer-related markers (β-catenin, CDX-2, and CK20) in organoid lines were detected in Histological and immunohistochemical images (Fig. 6B), which proved that we had successfully constructed colorectal tumor organoids with similar histopathological characteristics with colorectal cancer. In order to evaluate materials and drugs toxicity on CRC organoids, 0, 25, 50, 75, 100, and 125 µg/mL of c-HAP, c-HAP/DOX (DOX load capacity is 70 µg/mg), and DOX were used to treat PDCCOs for 48 h, and the survival rate was detected by CellTiter-Glo®3D reagent. According to Fig. 6C, IC50 value of c-HAP/DOX was approximately 125 µg/mL, while in contrast, the IC50 of c-HAP/DOX on SW480 cells was about 80 µg/mL after 24 hours. Interestingly, c-HAP did not show toxicity to PDCCOs, but promoted the growth of PDCCOs at low concentrations, which was slightly different from the cell experiment. Meanwhile, it can be seen that with the increase of c-HAP concentration, the activity of PDCCOs also showed a certain downward trend, although compared with the activity of PDCCOs in the control group, this toxicity is almost negligible. These results can be attributed to the fact that organoids, as a system containing a variety of stem cells, have more vital anti-interference ability against external stimulation and can more genuinely reflect the actual effect of nano-drug carrier system in the human body.
Fig. 6Construction of PDCCOs in vitro and its application to evaluate the antitumor effects of materials. (A) Extraction and culture of patient-derived colon tumor organoids. Scale bars: 100 µm. (B) Histological and immunohistochemical images showing the organization structure and status of proliferation marker (Ki-67) and Colon cancer-related markers (β-catenin, CDX-2, and CK20) in organoid lines. Scale bar: 50 µm. (C) Quantitative detection of organoid activity after incubation with different doses of c-HAP/DOX for 48 hours, and the corresponding concentration of DOX was 0, 1.75, 3.5, 5.25, 7.0, 8.75 µg/mL, respectively (n = 3). *p < 0.05, **p < 0.01, ***p < 0.001, and“ns” indicated no statistical difference. (D) CLSM images of organoids nuclei and cytoskeleton after incubation with 100 µg/mL c-HAP, c-HAP/DOX, and 7 µg/mL DOX for 48 h. The cytoskeleton was stained by phalloidin (green), and the cell nuclei were stained using DAPI (blue). Scale bars: 200 µm. (E) CLSM images of organoids Live/Dead staining after the incubation with 100 µg/mL c-HAP, c-HAP/DOX, and 7 µg/mL DOX for 48 hours. Scale bars: 200 µm.
Scheme 1Schematic illustration. Using the synthesized c-HAP to adsorb DOX (c-HAP/DOX). The release of DOX can be triggered by the tumor microenvironment (pH=6.0) when c-HAP/DOX is enriched to the tumor site through EPR effect.
To investigate the effects of c-HAP/DOX on the cytoskeleton structure, PDCCOs were treated by c-HAP and c-HAP/DOX at the concentration of 100 µg/mL and DOX at the concentration of 7 µg/mL for 48 h (Fig. 6D). Phalloidin and DAPI staining results showed that c-HAP had little effect on the skeleton structure in PDCCOs. However, some skeleton structure damage was observed in DOX and c-HAP/DOX treated PDCCOs. Moreover, the skeleton integrity of PDCCOs also showed a certain damage trend with the continuous increase of c-HAP/DOX concentration because there is more cell nucleus exposed outside the skeleton in a dose-dependent manner, indicating that the tissue structure of PDCCOs was damaged. (Fig. S4A).
PDCCOs were incubated by different doses of c-HAP/DOX for 48 h and stained with the live/dead Kit (Fig. S4B). The number of dead organoids increased with increasing doses of c-HAP/DOX. In a high concentration (200 µg/mL), the number of dead cells increased (red) in the cavity of PDCCOs, showing a trend toward fragmentation and lysis. Then we compared the effects of c-HAP, DOX, and c-HAP/DOX on the growth of PDCCOs at the same material/drug concentration (Fig. 6E). When the concentration is 100 µg/mL, there are only a small number of dead cells in PDCCOs treated with c-HAP. On the contrary, many dead cells were accumulated in PDCCOs treated with free DOX and c-HAP/DOX, indicating that DOX released by c-HAP/DOX after uptake by PDCCOs can have a significant inhibitory effect on its growth.
5. Conclusion
In this work, we developed an acid-triggered drug release nanoplatform based on PDA regulated synthesis of HAP nanoclusters. PDA can induce the synthesis of HAP with uniform size, modify the surface of HAP, and further improve the loading efficiency of c-HAP to DOX through π-π stacking. Moreover, c-HAP/DOX can respond to tumor slightly acidic environment and release DOX to promote SW480 cells apoptosis. In vitro Cell experiments demonstrated that the material can be absorbed significantly by endocytosis. More importantly, we successfully constructed a patients-derived tumor organoids model and used it to evaluate the toxicity of c-HAP and c-HAP/DOX in vitro. Although it takes longer for drugs and nano-drug carrier systems to play a role in tumor organoids than 2D cell models, the c-HAP/DOX shows a significant tumor inhibition effect in both in vitro 2D cell model and 3D tumor organoids model, the organoids model can better help us to study the tissue behavior and changes in the process of nanocarrier treatment of tumors. Meanwhile, we believe that the patient-derived tumor organoid models will provide a more effective prediction for the efficacy evaluation of the nanocarriers.
Author contributions
Conceptualization, Tianhao Deng; Dandan Luo and Xiangdong Kong; methodology, Tianhao Deng; software, Tianhao Deng and Rui Zhang; validation, Qingwei Zhao, Ruibo Zhao and Shibo Wang; formal analysis, M. Zubair Iqbal; investigation, Ruibo Zhao and M. Zubair Iqbal; resources, Yeting Hu and Qingwei Zhao; data curation, Tianhao Deng; writing—original draft preparation, Tianhao Deng; writing—review and editing, Tianhao Deng; M. Zubair Iqbal and Dandan Luo; visualization, Rui Zhang; supervision, Xiangdong Kong; project administration, Xiangdong Kong; funding acquisition, Xiangdong Kong. All authors have read and agreed to the published version of the manuscript.
Declaration of interests
The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.
Acknowledgments
This research was financially supported by the Key Research & Development Program of Zhejiang Province (2021C01180, 2019C04020) and the National Natural Science Foundation of China (51672250, 51902289).
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