In 2018, 1.9 million cancer-related deaths and 3.9 million new cases were reported in Europe, with an increase in cancer incidence and death rates expected [1
]. The main treatments for cancer include chemotherapy, radiotherapy, and surgery. However, chemotherapy results in toxicity towards normal cells and the development of therapy-resistant cancers. Integrating the above methods is not sufficiently effective [2
]. Therefore, active targeting of cancer cells with anticancer drugs has been explored.
Natural medicines account for 60% of anticancer agents used clinically [4
]. Natural products isolated from plants (also called natural prodrugs)—such as vincristine, taxanes, and camptothecin—have a long history in the treatment and prevention of cancer. Natural prodrugs offer safe, cost-effective, and diverse biological-medicinal activities [5
]. Despite these advantages, only a few natural prodrugs are in clinical use because of several limitations, specifically poor water solubility, low bioavailability, short half-life, and non-specific targeting. Nanomedicine has the potential to offer solutions to circumvent these limitations [7
Among anticancer natural plant-derived prodrugs is piperine (Pip), an alkaloid of special interest. Pip is an amide alkaloid extracted and isolated from the fruits of the black pepper plant (Piper nigrum
Linn). Black pepper has a top position among other spices and kitchen uses due to its unique pungency and flavor, earning it the nickname “the king of spices” [10
]. The average of consumption of black pepper is equivalent to 16 mg to 30 mg of pure Pip per person per day [12
]. Black pepper contains 6% to 9% pure Pip based on the dry weight [11
]. Pip has a long history of use in traditional Chinese, Indian, and Arabic medicine as a remedy for many illnesses (e.g., pain alleviation, indigestion, chills, rheumatism, infection, influenza, fever). At the pharmacological level, Pip improves the bioavailability of drugs, either synthetic or natural, when combined [13
], and has anti-inflammatory [15
], neuroprotective [16
], and anti-oxidant effects [17
]. Anti-tumor effects have been reported in several cancers both in vitro and in vivo, including melanoma, breast, ovarian, colon, lung, liver, and prostate [18
]. Despite these fascinating properties, Pip has not yet been used clinically due to its inherent limitations in regards to solubility, site-specific targeting, and bioavailability.
Recently, many strategies have been developed for Pip delivery systems using different nanoparticles, including chitosan-sodium tripolyphosphate [25
], solid lipids [26
], PLGA [27
], chitosan nanoparticles [28
], and self-emulsifying drug delivery [29
]. However, most of the delivery systems developed for Pip have investigated polymeric or lipid nanocarriers. The above-mentioned systems have limitations as far as long-term release effect is considered. Release of over 90% of encapsulated Pip takes place within few hours. This effect is expected since release depends on degradation of the polymeric drug carrier such as chitosan, which takes place in short time. Inorganic nanocarriers used in drug delivery systems (DDSs) for drugs and therapeutic agents have included mesoporous silica nanoparticles (MSNs), magnetic nanoparticles, zinc nanoparticles, and others to treat various diseases [30
]. Hydroxyapatite is the primary inorganic component of teeth and bone. They exhibit a good biocompatibility compared to other inorganic materials that biodegrade in biological fluids. With such fascinating properties, hydroxyapatite nanoparticles have shown many efforts in the delivery of drug and therapeutic agents with prolonged release [34
]. Hydroxyapatite nanoparticles (HAPs) are of great interest in biomedical applications, such as bone regeneration [40
]. Though HAPs have many applications as vehicles, especially in anticancer therapy, they have not yet designed DDSs for important prodrugs for cancers through active targeting. In the current study, we established a novel DDS comprising HAPs as nanocarriers loaded with Pip, followed by coating with gum Arabic (GA) and conjugation of folic acid (FA) on the surface as active targeting ligands for cancer selectivity (Scheme 1
). The delivery system shows promising and potential anticancer effects against monolayer (two dimensions) and spheroid (three dimensions) HCT116 colon cancer cells. The current study demonstrates a prolonged release effect (over 90 h) for Pip from loaded into agglomerated HAP nanoparticles. It depends on pH conditions and total Pip content and is considerably longer than for organic delivery systems for delivering Pip. Such a long-term release effect is required for cancer therapy. Furthermore, our system shows quite a high drug loading capacity of over 20 wt%, which may be superior over the existing systems. Therefore, the current results show a possible way to deliver Pip to target colon cancer cells with a prolonged release effect.
2. Materials and Methods
2.1. Synthesis and Surface Modification of Hydroxyapatite Nanoparticles
HAPs were synthesized using the hydrothermal method according to Kuśnieruk et al. [42
]. The surface of the HAP was functionalized with phosphonate (P) groups by adding 1.5 g of HAPs to 100 mL of 18.2 MΩ ultra-pure water (Milli-Q®
system, Millipore, Darmstadt, Germany) containing 1.5 mL of 3-(trihydroxysilyl)propyl methylphosphonate monosodium salt solution (Santa Cruz Biotechnology, Dallas, TX, USA) with stirring at room temperature. This solution was left for 24 h, and then filtered, washed several times with deionized water, and dried at 60 °C for 24 h. In this condition, the post functionalization of HAP with P groups is possible via the interaction hydrogen bonding of P to a hydroxyl group.
2.2. Piperine Loading
The Pip loading ratio was 1:3 (drug: nanoparticle); 300 mg Pip (Sigma-Aldrich, St. Louis, MO, USA) was dissolved in 20:80 ultrapure water:ethanol (Fisher Scientific, Loughborough, UK) at room temperature, followed by 900 mg of HAPs and HAP-Ps. The pH adjusted to either 7.2 or 9.3. The mixture solution was stirred for 24 h, and then evaporated at 60 °C (Rotavap; Büchi, Flawil, Switzerland). The dried mixture was resuspended in ultrapure water several times to remove unloaded molecules before drying in a 60 °C oven for 12 h. The obtained product was named HAP-Pip7.2, HAP-Pip9.3, HAP-P-Pip7.2, and HAP-P-Pip9.3. These materials were intended for coating with GA in subsequent steps.
2.3. Gum Arabic Coating and Folic acid Conjugation
For GA coating, GA solution (1%) was prepared by dissolving GA powder (Acros Organics, Geel, Belgium) in 0.1 M of NaOH (Acros Organics, Geel, Belgium) under high speed stirring at 60 °C. Next, 400 mg of HAP-Pip9.3 or HAP-P-Pip9.3 was suspended in GA solution and stirred at 250 rpm for 72 h at room temperature. The solution was centrifuged, washed several times with deionized water until the pH of the solution was neutral (7 to 7.4), and then dried at 60 °C in an oven for 12 h. The resulting products were named HAP-Pip9.3-GA and HAP-P-Pip9.3-GA.
For FA conjugation, we prepared the GA coating using the same procedure but without the drying step. Subsequently, the GA-coated HAPs (not dry) were suspended in the activated FA solution. The FA-activated solution was prepared in a separate step by adding 100 mg of FA (Sigma-Aldrich, St. Louis, MO, USA) and 50 mg of EDC (Acros Organics, Geel, Belgium) in 20 mL of NaOH (0.1 M) with stirring (400 rpm) at 60 °C. Typically, GA-coated particles were suspended in 10 mL of FA solution and stirred at 275 rpm for 40 h at room temperature. The solution was centrifuged to collect the particles, which were then washed several times to remove unbonded FA and EDC molecules. The obtained product was dried at 60 °C in an oven for 12 h and named HAP-Pip9.3-GA-FA and HAP-P-Pip9.3-GA-FA. The materials were stored at room temperature until further use.
Investigations of total drug-loading capacity (TLC) and entrapment efficiency (EE) using UV–vis and thermogravimetric (TG) analysis are detailed in the Supplementary Information
2.4. Characterization Techniques
The following techniques were used to characterize the resulting nanomaterials: field emission scanning electron microscopy (FE-SEM; Ultra Plus, Zeiss, Jena, Germany); powder X-ray diffraction (XRD; X’PertPRO System, PANalytical, Marietta, GA, USA) utilizing CuKα radiation (2θ range of 10–100°); Fourier transform infrared (FTIR) spectroscopy (Bruker Optics Tensor 27, Bruker Corporation, Billerica, MA, USA) equipped with attenuated total reflectance (Platinium ATR-Einheit A 255); simultaneous thermal analysis (STA) coupled with differential scanning calorimetry (DSC; STA 499 F1Jupiter, NETZSCH-Feinmahltechnik GmbH, Selb, Germany); the NOVA automated gas sorption system from Quanta Chrome Instruments (Boynton Beach, FL, USA) to measure pore size distribution; and the Brunauer, Emmett, and Teller (BET) specific surface area analysis (Gemini 2360, Micromeritics, Norcross, GA, USA) according to ISO 9277:2010. The powders were dried at 150 °C (without drug) or 50 °C (with drug and polymer) for 24 h under constant helium flow (FlowPrep 060 desorption station, Micromeritics) before analysis for additional confirmation. In STA-DSC, samples (5–10 mg) were placed in the STA unit’s alumina pan. Before measurement, helium flowed for 30 min through the STA furnace chamber. The heating rate was 10 °C/min in a helium/air mixture to 850 °C. Zeta potential was determined using a NanoZS Malvern ZetaSizer (Malvern, UK) based on a phosphate-buffered saline (PBS) suspension of nanoparticles (adjusted for different pH) at 24 °C.
2.5. In Vitro Drug Release
We studied the dissolution profile of Pip using a bottle method with a cellulose dialysis bag (MWCO 12,000 g/mol, Sigma-Aldrich CHEMIE GmbH, Sternheim, Germany). Each bag contained 3 mL of PBS with Pip-loaded HAPs and was closed from both sides before immersion in a capped glass bottle containing 50 mL of PBS adjusted for different pH: 5, 6.8, or 7.4. Bottles were incubated at 37 °C ± 0.5 °C in a WNB14 Memmert Shaking Water Bath (INDO Gama Pratama, Yogyakarta, Indonesia) with the rotation speed fixed at 150 rpm (Erweka GmbH, Hessen, Germany). We added 0.01% w
sodium azide to the release media as a preservative. At 6, 8, 12, 24, 48, and 72 h, a 2 mL aliquot was withdrawn, filtered through a 0.45-mm Millipore filter, and measured at 342 ± 0.05 nm against the blank cuvette. We maintained sink conditions by adding an equal volume of fresh buffer with each sampling. Each experiment was carried out in triplicate. The drug release data were analyzed using KinetDS3 pharmacokinetics software from Jagiellonian University (Department of Pharmaceutical Technology and Biopharmaceutics, Faculty of Pharmacy), employing both linear and non-linear regression. Release efficiency (RE) and mean dissolution time (MDT) were also determined [43
2.6. Cell Culture
We used MCF 7 human breast adenocarcinoma cells and Caco2 human colon carcinoma cells, with WI-38 human fibroblast cells as the normal reference (American Type Culture Collection [ATCC], Manassas, VA, USA). The cells were maintained and tested at the Confirmatory Diagnostic Unit (VACSERA, Dokki, Giza, Egypt). These three cell lines were cultured in Dulbecco’s modified Eagle medium (DMEM) supplemented with 10% fetal bovine serum, streptomycin (100 μg/mL), and penicillin G (100 U/mL; all from Gibco, Thermo Fisher Scientific) at 37 °C in a humidified atmosphere containing 5% CO2. For in vitro biological evaluations, we used HCT116 human colon cancer cells (Sigma-Aldrich, ECACC, USA) due to the high FA receptor content at the cell surface. The HTC116 cells were cultured in McCoy’s 5a medium supplemented with 10% fetal bovine serum, streptomycin (100 μg/mL), and penicillin G (100 U/mL) at 37 °C in a humidified atmosphere containing 5% CO2 (BioMedical Engineering Laboratory, Warsaw University of Technology, Warsaw, Poland). For spheroids, HCT116 cells (ATCC, Manassas, VA, USA) were cultured in DMEM (1 g/L glucose) supplemented with 10% fetal bovine serum, 2 mM glutamine, 50 μg/mL streptomycin, and 60 μg/mL penicillin. The cells were kept at 37 °C in a humidified atmosphere of 5% CO2. The cells were split twice a week and cell morphology and growth monitored on a weekly basis. Agarose-coated 96-well plates were obtained by preparing and autoclaving (121 °C, 20 min) a solution of agarose (Sigma-Aldrich Chemie GmbH, Darmstadt, Germany) in non-supplemented DMEM (1.5% w/v). The agarose solution was preheated in a water bath and 50 μL added to each well of a 96-well flat-bottom microtiter plate under sterile conditions. The plates were left to cool to room temperature and repacked. Plates could be stored at room temperature for 10 days.
2.7. In Vitro Cytotoxicity
Cytotoxicity and anticancer activity against MCF7, Caco2, and WI-38 cells were assessed as described previously [31
]. We tested HAPs and HAP-Ps for cytotoxicity at 12.3, 37, 111, 333, and 1000 µg/mL. For all prepared nanoformulations and free Pip, anticancer activity was assessed at 2.4, 7.4, 22, 66, and 200 µg/mL. The concentrations of the nanoformulations were designed to obtain an equivalent amount of Pip in each. All stock solutions were prepared in PBS (Gibco/Life Technologies, Thermo Fisher Scientific, Langenselbold, Germany). PBS without nanoformulations or Pip was used as a control. Treated cells were incubated for 48 h and 72 h under the same conditions and the absorbance at 540 nm measured using a Robotnik P2000 ELISA reader (Robotnik India PVT LTD, Thane, India). Assays were performed in triplicate and the data expressed as mean ± standard deviation (SD) in terms of cell viability.
Cytotoxicity and anticancer activity against monolayer HCT116 cells were assessed by the XTT assay. The HCT116 cells (2 × 105 cells/mL per well) were seeded in 96-well tissue culture plates. The cell monolayers were confluent after 24 h at 37 °C in a humidified atmosphere containing 5% CO2. The medium was removed and each well washed with PBS before adding suspensions of sample in culture medium at various concentrations. Each experimental plate contained replicates of negative control (NC, cells in pure medium), positive control (PC, cells treated with 2% Triton X-100), and blank control (BL, no cells). All cultures were incubated for 24, 48, and 72 h under the same conditions. We observed cell morphology under inverted light microscopy to detect the following morphological alterations: cell rounding and shrinking, loss of confluency, and/or cytoplasm granulation and vacuolization). We evaluated the cellular metabolic activity using the 2,3-bis-(2-methoxy-4-nitro-5-sulfophenyl)-2H-tetrazolium-5-carboxanilide (XTT) assay. The XTT reagent (Thermo Fisher, Eugene, OR, USA) solution (1 mg/mL) was prepared in culture medium and PMS (5 mM) added shortly before application to cells. Next, 50 µL of XTT solution with PMS in culture medium was added to each well and incubated at 37 °C in a humidified atmosphere containing 5% CO2 for 4 h. The plates were shaken gently for 5 min and the absorbance read using a BioTek reader at λ1 = 450 nm and λ2 = 630 nm. Each sample was run four times and the data expressed as mean ± standard deviation (SD) in terms of cell viability.
For the formation of HCT116 spheroids, 200 μL of HCT116 cell suspension with 5 × 103 cells was seeded into each well of an agarose-coated 96-well plate. The plates were incubated at 37 °C in 5% CO2. Spheroids formed by day 4, and the diameter of the formed structures was measured using phase-contrast microscopy and Zeiss software (Axio Observer 3, Carl, Zeiss Microscopy GmbH, Germany). Each well was exchanged for 100 μL of fresh medium and sample suspension. HAPs and HAP-Ps were prepared at 12.3, 37, 111, 333, and 1000 µg/mL. Pip, HAP-Pip7.2, HAP-P-Pip7.2, HAP-Pip9.3, HAP-P-Pip9.3, HAP-Pip9.3-GA, HAP-P-Pip9.3-GA, HAP-Pip9.3-GA-FA, and HAP-P-Pip9.3-GA-FA were prepared in supplemented DMEM medium at 2× concentration to obtain the final concentration of 2.4, 7.4, 22, 66, or 200 μg/mL in each well. The suspensions were vortexed before addition to the wells. The 3-day-old spheroids were incubated with prepared suspensions for 48 h and 72 h before measuring cytotoxicity and diameter. The positive control was 10% Triton X-100 in the standard medium. The acid phosphatase assay (APH) was performed to evaluate cytotoxicity. The assay buffer was made up of 3M stock solution (pH 5.2), 0.1 M sodium acetate (Sigma-Aldrich Chemie GmbH, Schnelldorf, Germany), and 0.1% (v/v) Triton X-100 (Sigma-Aldrich Chemie GmbH, Schnelldorf, Germany) in deionized/distilled water. The buffer was stored at 4 °C for up to 4 weeks. Immediately before use, we prepared the substrate solution (final pH 4.8) by supplementing the assay buffer with 2 mg/mL ImmunoPure p-nitrophenyl phosphate (Sigma-Aldrich, Merck KGaA, Darmstadt, Germany). At each time point, the sphericity, spheroid integrity, and diameter were analyzed by phase-contrast imaging before the APH assay. Spheroids were carefully transferred with the entire supernatant into standard flat-bottom 96-well microplates using a manual eight-channel pipettor. The plates were washed with PBS 3× by carefully replacing 160 μL of the liquid above spheroids with PBS, leaving 100 μL of liquid in each well after the washing process. To each well, we added 100 μL of APH assay buffer. After incubating for 90 min at 37 °C, we added 10 μL of 1 N NaOH (Sigma-Aldrich Chemie GmbH, Germany) to each well. Within 10 min, we measured the absorption at 405 nm using an Epoch Biotek Microplate reader (Epoch Biotek, Winooski, VT, USA).
2.8. Anticancer Effects
In 24-well culture plates, HCT116 cells (5 × 105 cells/well) were grown on sterile 12-mm-diameter glass coverslips to sub-confluence and allowed to attach for 24 h. For the cellular uptake study, the cells were treated with HAP, HAP-P, HAP-Pip9.3-GA, HAP-P-Pip9.3-GA, HAP-Pip9.3-GA-FA, and HAP-P-Pip9.3-GA-FA at 100 μg/mL in culture medium for 4, 24, 48, and 72 h. Cells without treatments were used as a control. The treated cells were washed with PBS before being fixed with 4% paraformaldehyde in PBS for 2 h at room temperature. The cells were then dehydrated in a graded ethanol series (30, 50, 70, and 98%; 30 min each) before drying in a laminar chamber overnight.
2.9. Statistical Analysis
Data are expressed as mean ± SD. Significant differences were calculated by analysis of variance. Means were compared by the least significant difference at p
< 0.05. Cytotoxicity towards MCF7, Caco2, and WI-38 cells was evaluated by IRRE STST 2005 software (ERRI Institute). For monolayer HCT116 cells and HCT116 spheroids, two-way ANOVA with Tukey’s multiple comparisons at p
< 0.05 was performed using Prism software (Prism, GraphPad, San Jose, CA, USA). The drug-loading content and EE were evaluated by one way ANOVA [44
Pip exerts anticancer effects via different mechanisms of action. Most of the mechanisms that have been reported are based on the inhibition of proliferation and survival as a result of the modulation of cell cycle progression, antioxidant activities due to detoxification of enzymes and suppression of stem cell self-renewal, and anti-apoptotic actions by many molecular signaling pathways [53
]. Pip has been shown to play an important role in mediating several enzymes and transcription factors that contribute to hindering the process of invasion, metastasis, and angiogenesis during cancer progression [53
]. Importantly, Pip is considered a potent inhibitor of p-glycoprotein (P-GP), which results in the inhibition of multidrug resistance in cancer [55
]. Recently, Pip analogs have been developed to target P-GP and overcome drug resistance in cancer [56
]. Interestingly, Pip has shown selective anticancer effects on cancer cells compared to normal cells [53
]. Therefore, we intended to design a delivery system for Pip as a promising drug candidate derived from natural substances. We show for the first time that delivery of Pip-loaded HAPs using FA ligands for active targeting resulted in significant inhibition against colon cancer cells in monolayer or spheroids.
The HAPs exhibited an aggregation effect with a mix of nearly elongated to spherical shapes. The particle size of HAP powder was an average of 9.7 ± 0.1 nm based on the specific surface area measurements according to Kuśnieruk et al. [42
]. SEM images demonstrated the obvious differences between particles before and after GA coating and FA conjugation, confirming the successful preparation. Upon the addition of HAP powder in the PBS adjusted to various pH, the nanoparticle size increased to micrometer-scale as expected based on the tendency of hydroxyapatite particles to agglomerate in biological media. These effects are related to their stability into aqueous media and zeta potential strength, as weaker potential values can induce electrostatic repulsion between particles, resulting in the induction of agglomeration [57
]. HAPs at pH 7.4, 6.8, and 5.5 carry negative potential up to −15 mV. The plausible reason for negative zeta potential is OH groups [36
]. Particles with zeta potential values < ±10 mV can agglomerate [58
], leading to an increase in the size of particles when investigated by DLS. Thus, our results are consistent with previous reports [57
]. We speculate that the main reason for the increase in HAP size is due to the agglomeration effect and not Pip loading or coating. After injection of the particles in vivo they are distributed in different organs depending on the different routes of in vivo application: sub-cutaneous, intramuscular, and intravenous injection [60
]. Therefore, for future in vivo studies, the selection of intra-muscular, intradermal, or subcutaneous depot will be needed in order to avoid the barrier made by the reticulo-endothelial system (RES). Oral administration seems to be preferred, since Pip would be directly released in colon and targeted to cancerous cells.
The specific surface area, total pore volume, and pore size were greatly decreased when particles were loaded with Pip, coated with GA, and conjugated to FA compared to the starting material for both HAPs and HAP-Ps. This observation is expected and in agreement with previous studies [30
]. Interestingly, the pore size distributions calculated by BET method from N2 adsorption/desorption measurements showed that HAPs had a pore size ranging from 9 nm to 13 nm. There are two possible explanations for this observation. First, the HAPs (10 nm) are mesoporous structures themselves. Second, the aggregation/agglomeration behavior of HAPs leads to free spaces between the particles. The second reason is more logical. This porosity is necessary for drug encapsulation, which decreases the surface area properties.
The thermal stability, drug content, crystalline, and decomposition characteristics of Pip can be explored by thermal analysis. STA showed that HAPs lost about 10 wt % as moisture up to 1000 °C, which in the line with data by Kuśnieruk et al. [42
]. When HAPs were further processed by loading and coating, high percentages of weight loss were measured because of decomposition of the organic compounds Pip, GA, and FA. In this context, we can calculate the drug-loaded content in inorganic nanoparticles [62
]. Pip shifted to the non-crystalline state, as the peak indicating its melting point was not observed in all loaded nanoparticles. This observation can be explained by most Pip molecules being entrapped inside pores on the surface of HAPs. These results are in agreement with reports of loading drugs in mesoporous nanoparticles [30
]. Changing the crystalline state of a water-insoluble drug to the non-crystalline state using mesoporous materials can enhance solubility, increasing the drug therapeutic activity [64
]. The DTG results confirmed the thermal stability and decomposition of Pip, GA, and FA. STA-MS revealed that no toxic substances are present in the materials. All thermal analysis results revealed a good relationship between each other (STA, DTG, DSC, MS) and showed their importance in complementary techniques for characterizing materials.
Using FT-IR analysis in the characterization of materials confirmed the Pip loading on HAPs, with several peaks at 562, 600, 670–975, 1020, 1195, 1253, and 1329–1675 cm−1 corresponding to free Pip, showing the fraction of Pip molecules on the surface. The appearance of shifted peaks increased peak intensities, and new peaks confirmed the GA coating and FA conjugation to loaded nanoparticles. Overall, the FT-IR results verified the successful preparation. Going further, the entrapment of Pip in pores changed to the non-crystalline state as confirmed by DSC and XRD.
The drug loading properties (TLC and EE) are the main factor determining drug delivery either long or short-term for desirable therapeutic actions. Previously, Shen et al. reported that a drug-loading content ≥10% is important [65
]. Therefore, we determined these properties using two different techniques. The first was by thermal properties depending on weight loss data from STA. The total Pip content in HAPs and HAP-Ps ranged from 18 wt % to 20 wt % depending on the nanoformulation and pH. The EE reached 72–81wt % depending on the presence or absence of the GA coating and FA conjugation. The drug content calculated based on TG analysis was reported previously for inorganic nanoparticles used in drug delivery [31
]. The second way was UV–vis spectroscopy, which is a commonly used method in most drug delivery strategies. The Pip content ranged from 16% to 22%, which differs from the TG calculation [62
]. The EE determined by the UV–vis method reached 75–85%. We expect this difference to be due to the principle of both methods, as UV–vis depends on the absorbance of samples, whereas TG analysis depends on the thermal behavior of the carrier and drug. The UV–vis method seems to be more accurate than TG analysis. The Pip content of HAPs has not been reported previously, but our results are comparable with those of other strategies used for Pip loaded (4.7%) with mixed nano-sized micelles for cancer cell delivery [66
]. The results prove the feasibility of HAPs as a Pip delivery vehicle.
The highest solubility of Pip is in ethanol compared to other solvents. The solubility of Pip in PBS is pH-dependent, and the highest solubility was achieved at pH 5 compared to pH 6.8 and 7.4. This reflects the shorter release time at pH 7.4. This solubility characteristic is attributed to its alkaloid structure. Alkaloids are known to be weak bases because of their oxygen and nitrogen termini, with easy solubility in strong acids, making them more stable. Alkaloids are hydrophobic because of the benzene ring and cyclic structures forming their main chemical skeleton. All of these factors make Pip soluble in an organic solvent (e.g., ethanol) and slightly soluble in water. The results align with the Pip solubility data available in the literature [67
]. Another characteristic is the high rate of solubility at pH 5, which suggests that Pip is more stable, allowing diffusion in acidic release media with controlled-release behavior.
Importantly, the GA polymeric coating conjugated to FA prolongs the release effect because it hinders Pip release. The release parameters and kinetics were altered with surface modification and polymer coating of the drug-loaded nanoparticles. These results are similar to our previous results in mesoporous silica nanoparticles loaded with thymoquinone and coated with different polymer mixtures [31
], as well as other studies [68
]. Regarding the release of Pip from other DDSs, Shao et al. reported that Pip release from a self-emulsifying DDS was 80% over 1 h [29
]. In another study, 100% of loaded Pip was lost from chitosan nanoparticles over 24 h [28
]. Laha et al. showed that the release of Pip from electrospun cross-linked gelatin nanofibers was affected by the pH of the medium and >90% was released over 24 h, fitting to Higuchi kinetic model [69
]. Thus, the current study demonstrates a promising prolonged release effect for Pip from agglomerated HAPs depending on the pH conditions.
Based on the solubility test for the components used in preparations, the GA would be the first to dissolve in contact with biological media of acidic or neutral pH. With respect to agglomerated HAPs, we propose that, in suitable release medium, HAP absorbs water and swells. Once it swells, HAP absorbs more water, leading to an increase in the nanoparticle pore size and subsequent release of Pip through these pores or the space between agglomerated nanoparticles. The process may continue until the HAP completely degrades [70
The kinetic model of Pip releases changes from Baker–Lonsdale to Korsmeyer–Peppas for the nanoformulations tested at pH 6.8, HAP, and the coating composed of GA and FA. According to the literature, such kinetic models are characteristic of the release from inorganic or polymeric nanoparticles. However, a change in the in vivo pharmacokinetics would be expected when tested in animals [29
]. The Baker–Lonsdale model is a modification of the Higuchi model. It was derived to describe drug release through an equation utilized for linearization of release data for micro/nano capsules or spheres while considering matrix porosity and pore size [71
]. However, Korsmeyer–Peppas model tends to linearize release data through establishing an exponential relationship between drug release and time. This model is a semi-empirical equation describing drug release from polymeric nanosystems when its release mechanism is not known or when there is superposition of two or more apparently independent mechanisms of drug transport, relaxation, and diffusion within and from the particles [72
]. On relating the presented information and the in vitro release data of Pip depending on HAP spherical to elongated shape, pore volume and size, as well as the polymers employed. Thus, when release of Pip from HAP nanoparticles is according to Baker–Lonsdale model, this would reveal that for such particles, pore size, volume and particle shape is the rate-determining factor over its polymeric coating. Consequently, when Korsmeyer–Peppas is the best fitting, the reverse is true.
The RE and MDT for Pip changed with surface modification and coating. Before coating, RE increased with increasing pH (pH 7.4 > pH of 6.8 > pH 5), whereas after coating, the RE increased with decreasing pH (pH 5 > pH 6.8 > pH 7.4). MDT values significantly increased with surface modification and coating, with greater increases as pH increased (pH 7.4 > pH 6.8 > pH 5). The maximum MDT was measured for HAP-P-Pip9.3-GA-FA. The addition of a polymeric coating forms a sheath around encapsulated Pip, resulting in a decreased RE at pH 7.4 but increased RE at pH 5. The reason for this finding is the inherent gelling property of GA. Gelation of GA occurs because of its oxidation, leading to the formation of highly dense cross-linking. Acting as a barrier, this gel layer retards the rate of Pip diffusion into the release medium. Thus, GA is highly recommended for slow release of water-insoluble drugs. The extent of release in vitro depends on the cross-linking density, as well as drug payload. Thus far, it appears a GA coating can enhance the colloidal stability of nanoparticles [73
]. The RE and MDT are used as model-independent kinetic parameters that can provide insights on drug absorption. Release efficiency is the area under the dissolution curve up to certain time, (t), expressed as percentage of area of rectangle described by 100% dissolution at the selected time [74
]. While MDT is kinetic parameter used to characterize drug release rate and indicate drug release retarding efficiency of polymers used. It depends on dose/solubility ratio [75
The results indicate strong anticancer activity of Pip through targeted delivery compared to free Pip and with less cytotoxicity towards normal cells. These results are in line with data published previously on HT-29 colon cancer cells, in which cell killing was via an apoptotic mechanism [20
]. The effect also depends on the cancer cell line, time, and concentration [53
]. Targeted delivery through HAP-Pip9.3-GA-FA and HAP-Pip9.3-GA-FA results in full inhibition of monolayer HCT116 cells compared to other tested lines treated at 200 µg/mL, showing a selective anticancer effect compared to MC7 breast cancer and Caco2 colon cancer cells. Further exploring the anticancer effects in HCT116 spheroids disclosed a weaker effect of ~50%. The high activity of free Pip strongly inhibits monolayer HT-29 colon cancer cells compared to HT-29 spheroids [20
]. These results agree with previous data on HT-29 spheroids showing that drug-loaded nanoparticles are more efficient than a free drug [76
]. The main reason for the difference is that spheroids are a more complex structure, similar to the solid tumor structure. Thus, spheroids offer a more rational platform for screening anticancer effects than monolayer cells [76
]. Thus, the 3D cell culture models, including spheroids, closely mimic the main characteristics of human solid tumors (e.g., organization of the structure, assembly of the cell layers, hypoxia, and nutrient gradients) [78
]. Therefore, monolayer cancer cells poorly predict the therapeutic efficiency or outcome in in vivo animal studies [76
]. Regarding spheroids, the preliminary results in the current study show that the strong effect can further improve anticancer effects with further developments and investigations.
HAPs and HAP-Ps exhibited no significant accumulation in cells compared to control cells, which is associated with their lack of ability to target cancer cells. Importantly, HAP-Pip9.3-GA-FA and HAP-P-Pip9.3-GA-FA had the strongest anticancer effects because of internalization in HCT116 cells compared to other formulations at 72 h. The significant changes in cell morphology were perforation of the cell membrane and disruption of intercellular adhesion, which indicates a high affinity of the FA-conjugated nanoparticles for the cell surface, probably due to the overexpression of folate receptors. This phenomenon substantially suppressed the viability of cells for reduced contact with the culture medium and nutrients, inhibiting intercellular contact, making cell division and contact with signal proteins difficult. Concerning the GA-coated nanoparticles, HAP-Pip9.3-GA and HAP-P-Pip9.3-GA also exhibited significant changes in cell morphology after 72 h related to the kinetic release data at low acidic conditions (see Figure 8
). Drug release from HAPs is strongly stimulated by a pH drop [36
]. The efficient cancer targeting by FA-conjugated agglomerated HAPs agree with previous studies of targeting colon cancer cells through FA conjugation of nanoparticles [79
With spheroids, HAP-Pip7.2, HAP-P-Pip7.2, HAP-Pip9.3, HAP-P-Pip9.3, HAP-Pip9.3-GA, HAP-P-Pip9.3-GA, HAP-Pip9.3-GA-FA, and HAP-P-Pip9.3-GA-FA had visible changes compared to control and free Pip. The main changes were smaller diameter, shrinkage, and defragmentation of spheroids [81
]. The nano-particulate-system containing FA is efficient for delivering Pip prodrug and accumulates in spheroids compared to free Pip; thus, anticancer effects are enhanced due to internalized of HAP-Pip9.3-GA-FA and HAP-P-Pip9.3-GA-FA in HCT116 cells spheroids compared to other formulations. This effect aggress with nanoparticles loaded with anticancer drug compared to free drug to target spheroids [76
]. To summarize the results obtained from our study, Table 5
shows the findings for the targeted delivery proposed by HAPs compared to free Pip prodrug. The most anticancer effective nanoformulations in monolayer and spheroids are those with FA (HAP-Pip9.3-GA-FA 671 and HAP-P-Pip9.3-GA-FA). Our findings are in the line with using of nanomedicine application for cancer targeting strategies and controlled release of anticancer drugs by various nanostructures [82