Biological evaluation of dimethylpyridine–platinum complexes with potent antiproliferative activity

Abstract

This study investigates the effect of three new platinum complexes: Pt2(2,4-dimethylpyridine)4(berenil)2 (Pt14), Pt2(3,4-dimethylpyridine)4(berenil)2 (Pt15) and Pt2(3,5-dimethylpyridine)4(berenil)2 (Pt16) on growth and viability of breast cancer cells and their putative mechanism(s) of cytotoxicity. Cytotoxicity was measured with MTT assay and inhibition of [3H]thymidine incorporation into DNA in both breast cancer cells. Results revealed that Pt14–Pt16 exhibit substantially greater cytotoxicity than cisplatin against MCF-7 and MDA-MB-231 breast cancer cells. In the case of human skin fibroblast cell, cytotoxicity assays demonstrated that these compounds are less toxic to normal cells than cisplatin. In addition, the effects of Pt14–Pt16 are investigated using the flow cytometry assessment of annexin V binding, analysis of mitochondrial potential, markers of apoptosis such as caspase-3, caspase-8, caspase-9, caspase-10 and defragmentation of DNA by TUNEL assay. These results indicate that Pt14–Pt16 induce apoptosis by the mitochondrial and external pathway.

Keywords:

• Apoptosis
• anticancer drugs
• cytotoxicity
• platinum complexes
• flow cytometry

Introduction

Breast cancer, the leading cause of death among women in the world, representing 23% of all cancer cases, is a heterogenic disease at cellular and molecular levels1,2. Therefore, there is an urgent need for novel drugs with an improved efficacy against breast cancer cells with minimal toxicity on normal cells. The cornerstone treatment for numerous malignancies, including breast cancer is cisplatin3,4. However, in spite of their high activity, the applications of cisplatin and similar compounds are limited by the side effects5,6. Therefore, despite large progress in anticancer drug development, there are still premises for search of new platinum antitumor complexes that will be both effective and safe. Of the nonclassical platinum compounds, the multinuclear complexes appear to offer the greatest potential as they are generally highly cytotoxic and maintain their activity in platinum resistant cell lines. Dinuclear platinum complexes that are structurally different from cisplatin are developed to circumvent the cellular resistance arising toward the mononuclear compounds, primarily decreased uptake, increased efflux and increased DNA repair.

Previously, we reported on a series of the dinuclear berenil–platinum complexes of general formula [Pt(berenil)₂X₄], where X = amine. These compounds bind to the minor groove of duplex DNA in A/T-rich regions, where they are thought to exert their biological activity through the inhibition of DNA-associated enzymes, such as DNA topoisomerases I and/or II, or possibly by direct inhibition of transcription7,8. The rationale for the design of these and related new DNA minor-groove binding drugs is that DNA-affinity plays an essential role in determining ultimate biological responses, even though by itself it is not the sole factor involved. The kinetic inertness of the berenil–platinum complexes also offers the advantage of reduced side effects in vivo and a significant decrease in drug loss through reactions with plasma proteins or other low-molecular-weight biomolecules. In order to broaden the pharmacological screening of these types of compounds, we synthesized the new dimethylpyridine–platinum complexes (Figure 1). The aim of our study was to see whether modification the compounds in terms of alkyl substitution in the pyridine ring improves anticancer activity in the title group of compounds (Figure 1).

Figure 1. Structure of compounds Pt14–Pt16.

One of the most effective strategies of cancer treatment is to induce cancer cells to apoptosis by using anticancer agents. Therefore, it is important that promising anticancer compounds should be able to induce apoptosis in cancer cells having different cellular and molecular characteristics. There are known two basic apoptotic pathways in mammalian cells: the extrinsic (receptor) and intrinsic (mitochondrial) pathways. The extrinsic pathway of apoptosis is initiated by FasL binding to the extracellular receptor and results in the formation of DISC that activates caspase-8. In some cells, caspase-8 interacts with intrinsic apoptotic pathway by cleaving Bid and leading to the release of cytochrome c9–11. The intrinsic apoptotic pathway is activated by various intracellular stimuli, including DNA damage, growth factor deprivation and oxidative stress12. The mitochondrial pathway is associated with an increased permeability of the mitochondrial membrane and cytochrome c translocation to the cytoplasm. Then, cytochrome c binds to the Apaf-1 and procaspase-9, forms the apoptosome and catalyzes the activation of caspase-9. The activation of both pathways activates effector caspase-3, caspase-6 and caspase-7, which are responsible for irreversible changes in cells.

Pt14–Pt16 cytotoxic effects were demonstrated by MTT assay and inhibition of [³H]thymidine into DNA. The objective of this study was also to investigate whether Pt14–Pt16 have any apoptotic effect in human breast cancer cells. It was assessed whether they cause apoptosis or not via Annexin V-FITC/propodium iodide staining procedure. The concentration of selected apoptotic markers involved in both apoptotic pathways, such as change of mitochondrial membrane potential, caspase-3, caspase-8, caspase-9 and DNA degradation, was measured in cell lysates of breast cancer cells using flow cytometry analysis and TUNEL assay.

Materials and methods

Materials

Dimethylformamide, K₂PtCl₄, KI, acetone, 2,4-dimethylpyridine, 3,4-dimethylpyridine, 3,5-dimethylpyridine, diethyl ether, methanol, ethidium bromide, cisplatin, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) were purchased from Sigma Chemical Co. (St. Louis, MO). Stock cultures of fibroblast cells, human MCF-7 breast cancer cells and human MDA-MB-231 breast cancer cells were purchased from the American Type Culture Collection (Manassas, VA). Dulbecco’s minimal essential medium (DMEM) and fetal bovine serum (FBS) used in a cell culture were products of Gibco (San Diego, CA). Glutamine, penicillin and streptomycin were obtained from Quality Biologicals Inc. (Gaithersburg, MD) [³H]Thymidine (6.7 Ci/mmol) was purchased from NEN, and Scintillation Coctail “Ultima Gold XR” from Packard (Waltham, MA). Acridine orange and ethidium bromide were provided by Sigma Chemical Co, as most other chemicals and buffers were used. FITC Annexin V Apoptosis Detection Kit II, JC-1 MitoScreen Kit, APO-Direct Kit were product of BD Pharmigen; FLICA Caspase 3 Kit, FLICA Caspase 8 Kit, FLICA Caspase 9 Kit, FLICA Caspase 10 Kit were obtained from ImmunoChemistry Technologies (Bloomington, MN).

Physical measurements

The structure of a synthesized compound was confirmed by ¹H NMR and ¹³C NMR spectra recorded on Brucker AC 200F (Germany) apparatus (¹H – 200 MHz and ¹³C – 50 MHz) in deuterated dimethylsulfoxide (d₆-DMSO). Chemical shifts are expressed in δ value (ppm). Multiplicity of resonance peaks are indicated as singlet (s), doublet (d), triplet (t), quartet (q) and multiplet (m). Infrared spectra were recorded on Perkin-Elmer Spectrum 100 FT-IR spectrometer as KBr pellets (4000–450 cm⁻¹). Mass spectra were recorded using a Mariner mass spectrometer (PerSeptive Biosystems, Houston, TX). Melting points were determined on Buchi 535 (New Castle, DE) (GER) melting point apparatus and were uncorrected. Elemental analysis of C, H and N was performed on a Perkin-Elmer 240 analyzer (Waltham, MA) and satisfactory results within ± 0.4% of calculated values were obtained.

Chemistry

General preparation of platinum complexes (Pt14–Pt16)

K₂PtCl₄ (0.72 mmol) was dissolved in 40 ml of deionized water. KI (7.2 mmol) was added and the reaction mixture was stirred for 30 min. Then, the corresponding dimethylpyridine (1.44 mmol) was added dropwise to the reaction mixture while stirring, to obtain a precipitate, cis-[Pt(dimethylpyridine)₂I₂]. The stirring was continued for 60 min and the precipitate was then collected by filtration. This compound was filtered, and then, it was washed with 30 ml deionized water and 5 ml acetone and dried in vacuum. Cis-[Pt(dimethylpyridine)₂I₂] (1.22 mmol) was suspended in 5 ml of an aqueous solution of silver nitrate (AgNO₃) (2.44 mmol). The reaction mixture was stirred for 24 h at room temperature in the dark. The AgI precipitate was filtered off. Berenil (1.22 mmol) and a solution of 10% NaCl (5 ml) were added to the filtrate and stirring until a precipitate of the corresponding dimethylpyridine–platinum complex formed. Afterwards, the product was filtered off and washed with a small amount of diluted HCl, deionized water, methanol, acetone and ethyl ether and dried under vacuum.

[Pt₂(2,4-dimethylpyridine)₄(berenil)₂]·4HCl·2H₂O (Pt14): Yield: 47.2%; yellow powder; mp 230–233 °C; ¹H-NMR (DMSO-d₆) δ (ppm): 9.35 (br s, 4H, amidine), 9.00 (br s, 4H, amidine), 8.55 (d, J = 7.5 Hz, 8H, Ar), 7.84–7.73 (m, 8H, Ar), 7.22 (t, J = 7.5 Hz, 8H, Ar) 7.02–7.15 (m, 16H, Ar), 2.50 (s, 3H, CH₃), 2.30 (s, 3H, CH₃). ¹³C NMR (DMSO-d₆) δ (ppm): 164.1 (amidine), 150.7 (Py), 148.6 (Ar), 147.2 (Py), 147.0 (Py), 133.2 (Py), 129.5 (Ar), 125.2 (Py), 122.0 (Ar), 118.0 (Ar), 19.2 (CH₃), 15.8 (CH₃); IR (KBr, cm⁻¹): 3356 (C=NH imine), 1675 (NCN/C=N imine), 1606 (CN pyridine/triazene), 1484 (CH₃), 1257 (triazene), 1173 (triazene), 830 (1,2,4-trisubstituted aromatic), 486 (Pt-N). MS (ES, HR) m/z (M⁺) calcd. for C₅₆H₆₈N₁₈Pt₂Cl₄ 1522.3924, found 1522.3920. Anal. calcd. for C₅₆H₆₄N₁₈Pt₂ 4HCl·2H₂O: C, 43.06; H, 4.65; N 16.15. Found: C, 42.99; H, 4.63 N, 16.06.

[Pt₂(3,4-dimethylpyridine)₄(berenil)₂]·4HCl·2H₂O (Pt15): Yield: 42.4%; lemon powder; mp 240–242 °C; ¹H NMR (DMSO-d6) δ (ppm): 9.35 (br s, 4H, amidine), 9.00 (br s, 4H, amidine), 8.55 (d, J = 7.5 Hz, 8H, Ar), 7.84–7.73 (m, 8H, Ar), 7.22 (t, J = 7.5 Hz, 8H, Ar) 7.02–7.15 (m, 16H, Ar), 2.51 (s, 3H, CH₃), 2.49 (s, 3H, CH₃). ¹³C NMR (DMSO-d6) δ (ppm): 164.1 (amidine), 158.5 (Py), 149.9 (Py), 149.2 (Ar), 147.3 (Py), 129.5 (Ar), 124.9 (Py), 122.6 (Py), 122.0 (Ar), 118.0 (Ar), 19.2 (CH₃), 15.8 (CH₃); IR (KBr, cm ⁻¹): 3351 (C=NH imine), 1686 (NCN/C=N imine), 1606 (CN pyridine/triazene), 1486 (CH₃), 1257 (triazene), 1173 (triazene), 850 (1,3,4-trisubstituted aromatic), 525 (Pt-N). MS (ES, HR) m/z (M⁺) calcd. for C₅₆H₆₈N₁₈Pt₂Cl₄ 1522.3924, found 1522.3920. Anal. calcd. for C₅₆H₆₄N₁₈Pt₂ 4HCl·2H₂O: C, 43.06; H, 4.65; N 16.15. Found: C, 42.89; H, 4.60 N, 16.10.

[Pt₂(3,5-dimethylpyridine)₄(berenil)₂]·4HCl·2H₂O (Pt16): Yield: 57.6%; lemon powder; mp 252–254 °C; ¹H NMR (DMSO-d6) δ (ppm): 9.35 (br s, 4H, amidine), 9.00 (br s, 4H, amidine), 8.55 (d, J = 7.5 Hz, 8H, Ar), 8.51 (s, 2H, Py), 7.84–7.73 (m, 6H, Ar), 7.22 (t, J = 7.5 Hz, 8H, Ar) 7.02–7.15 (m, 16H, Ar), 2.24 (s, 6H, CH₃). ¹³C NMR (DMSO-d6) δ (ppm): 164.2 (amidine), 147.8 (Py), 149.2 (Ar), 137.1 (Py), 133.3 (Py), 129.5 (Ar), 124.9 (Py), 122.0 (Ar), 118.0 (Ar), 18.5 (CH₃); IR (KBr, cm^− 1): 3317 (C=NH imine), 1686 (NCN/C=N imine), 1606 (CN pyridine/triazene), 1486 (CH₃), 1256 (triazene), 1172 (triazene), 697 (1,3,5-trisubstituted aromatic), 462 (Pt-N). MS (ES, HR) m/z (M⁺) calcd. for C₅₆H₆₈N₁₈Pt₂Cl₄ 1522.3924, found 1522.3920. Anal. calcd. for C₅₆H₆₄N₁₈Pt₂ 4HCl·2H₂O: C, 43.06; H, 4.65; N 16.15. Found: C, 42.89; H, 4.61 N, 16.17.

Biological activity

Cell culture

The MCF-7 breast cancer cells, MDA-MB-231 breast cancer cells and fibroblasts skin cells maintained as a monolayer in DMEM supplemented with 10% FBS, 50 U/ml penicillin, 50 μg/mL streptomycin at 37 °C in atmosphere 5% CO₂. Cells were cultured in Costar flasks and subconfluent cells were detached with 0.05% trypsin and 0.02% EDTA in calcium-free phosphate-buffered saline (PBS), counted in hemocytometers and plated at 5 × 10⁵ cells/well of 6-well plates (Thermo Scientific, New York, NY) in 2 ml of growth medium (DMEM without phenol red with 10% CPSR1). Cells reached about 80% of confluency at day 2 and in most cases such cells were used for the assays.

Cell viability assay

The growth inhibitory effects of the test compounds against MCF-7 and MDA-MB-231 breast cancer cells and human skin fibroblast cells were measured by using MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) assay as previously described13. Compounds were dissolved in dimethyl sulfoxide (DMSO) and screened at a range of concentrations against cancer and normal cells. Just before the experiments, stock solutions were diluted with the supplemented medium to obtain final concentrations of 5, 10, 20, 30, 40, 50, 75 and 100 μM. The cytotoxic activity was determined after 24 h and 48 h. All experiments were repeated at least three times.

[³H]thymidine incorporation assay

To examine the effect of the study compounds on cells proliferation, MCF-7 cells, MDA-MB-231 cells and human skin fibroblasts cells were seeded in 6-well plates and grown as described previously. Cells culture were incubated with varying concentrations (5, 10, 20, 30, 40, 50, 75 and 100 μM) of Pt₂(2,4-dimethylpyridine)₄(berenil)₂, Pt₂(3,4-dimethylpyridine)₄(berenil)₂, Pt₂(3,5-dimethylpyridine)₄(berenil)₂ or cisplatin and 0.5 μCi of [³H]thymidine for 24 h and 48 h at 37 °C. The cells were harvested by trypsinization and washed several times in the cold PBS (10 min/1500 g) until the dpm in the washes were similar to the reagent control. Radioactivity was determined by liquid scintillation counting. [³H]thymidine uptake was expressed as dpm/well.

Flow cytometry assessment of annexin V binding

To characterize the mode of cell death induced by Pt14–Pt16, a flow cytometry analysis was performed using Apoptosis Detection Kit II (BD Pharmingen, San Diego, CA) according to the manufacturer’s instruction. Cells (10 000 cells measured) were analyzed in a flow cytometer (BD FACSCanto II flow cytometer, CA). Annexin V bound with high affinity to phosphatidylserine and thus could be used to identify cells in all stages of the programed cell death. Propidium iodide (PI) stained cells with a disrupted cell membrane and it could be used to identify late apoptotic and dead cells14. Cells cultured in a drug-free medium were used as controls. Optimal parameter settings were found using a positive control (cells incubated with 3% formaldehyde in buffer during 30 min on ice). Results were analyzed with FACSDiva software (BD Bioscences Systems, San Jose, CA).

Fluorescent microscopy assay

The cell viability was estimated 24 h after the addition of the study compounds to assess apoptosis. The cell suspension (250 μl) was stained with 10 μl of the dye mixture (10 μM acridine orange and 10 μM ethidium bromide), which was prepared in PBS. Acridine orange (fluorescent DNA-binding dye) intercalated into DNA, making it appear green and bound to RNA, staining it red/orange. Ethidium bromide was only taken up by nonviable cells; its fluorescence predominated that of the acridine orange, making the chromatin of necrotic cells appear orange. Cells cultured in a drug-free medium were used as controls. Analysis was performed using Nikon Eclipse Ti inverted microscope, and results were analyzed with NIS-Elements software (both from Nikon Instruments Inc., Melville, NY).

Analysis of mitochondrial membrane potential

Disruption of the mitochondrial membrane potential (MMP) was assessed using the lipophilic cationic probe 5,5′,6,6′-tetrachloro-1,1′,3,3′-tetraethylbenzimidazolcarbocyanine iodide (JC-1 MitoScreen kit; BD Biosciences) as described previously. Briefly, unfixed cells were washed and resuspended in PBS supplemented with 10 μg/ml JC-1. Cells were then incubated for 15 min at room temperature in the dark, washed and resuspended in PBS for immediate BD FACSCanto II flow cytometry analysis. The percentage of cells with disrupted MMP was calculated in the FACSDiva software (both from BD Bioscences Systems, San Jose, CA).

Caspase-3 enzymatic activity assay

Caspase-3 activity was measured using FLICA Caspase 3 Assay Kit (ImmunoChemistry Technologies, Bloomington, MN) according to the manufacturer’s instructions. Briefly, cultured MDA-MB-231 breast cancer cells (1 × 10⁶) were washed with cold PBS twice and resuspended in buffer. Added 5 μl diluted FLICA reagent and 2 μl Hoechst 33342 to 93 μl of cell suspension and mixed by pipetting. The cells were incubated for 60 min at 37 °C. After incubation, cells are washed twice in 400 μl apoptosis wash buffer and centrifuged at 300 × g. After last wash, resuspended cells in 100 μl apoptosis wash buffer are supplemented with 10 μg/ml PI. Analysis was performed using BD FACSCanto II flow cytometer, and results were analyzed with FACSDiva software (both from BD Bioscences Systems, San Jose, CA).

Caspase-8 enzymatic activity assay

Caspase-8 activity was measured using FLICA Caspase 8 Assay Kit (ImmunoChemistry Technologies) according to the manufacturer’s instructions. Briefly, cultured both MCF-7 and MDA-MB-231 breast cancer cells (1 × 10⁶) were washed with cold PBS twice and resuspended in buffer. About 5 μl of diluted FLICA reagent and 2 μl of Hoechst 33342 are added to 93 μl of cell suspension and mixed by pipetting. The cells were incubated for 60 min at 37 °C. After incubation, washed cells twice in 400 μl apoptosis wash buffer and centrifuged at 300 × g. After last wash, resuspended cells in 100 μl apoptosis wash buffer are supplemented with 10 μg/ml PI. Analysis was performed using BD FACSCanto II flow cytometer, and results were analyzed with FACSDiva software (both from BD Bioscences Systems, San Jose, CA).

Caspase-9 enzymatic activity assay

Caspase-9 activity was measured using FLICA Caspase 9 Assay Kit (ImmunoChemistry Technologies, Bloomington, MN) according to the manufacturer’s instructions. Briefly, cultured both MCF-7 and MDA-MB-231 breast cancer cells (1 × 10⁶) were washed with cold PBS twice and resuspended in buffer. About 5 μl of diluted FLICA reagent and 2 μl of Hoechst 33342 are added to 93 μl of cell suspension and mixed by pipetting. The cells were incubated for 60 min at 37 °C. After incubation, washed cells twice in 400 μl apoptosis wash buffer and centrifuged at 300 × g. After last wash, resuspended cells, in 100 μl apoptosis wash buffer, are supplemented with 10 μg/ml PI. Analysis was performed using BD FACSCanto II flow cytometer, and results were analyzed with FACSDiva software (both from BD Bioscences Systems, San Jose, CA).

Caspase-10 enzymatic activity assay

Caspase-10 activity was measured using FLICA Caspase 10 Assay Kit (ImmunoChemistry Technologies, Bloomington, MN) according to the manufacturer’s instructions. Briefly, cultured both MCF-7 and MDA-MB-231 breast cancer cells (1 × 10⁶) were washed with cold PBS twice and resuspended in buffer. About 5 μl of diluted FLICA reagent and 2 μl of Hoechst 33342 are added to 93 μl of cell suspension and mixed by pipetting. The cells were incubated for 60 min at 37 °C. After incubation, cells are washed twice in 400 μl apoptosis wash buffer and centrifuged at 300 × g. After last wash, resuspended cells in 100 μl apoptosis wash buffer are supplemented with 10 μg/ml PI. Analysis was performed using BD FACSCanto II flow cytometer, and results were analyzed with FACSDiva software (both from BD Bioscences Systems, San Jose, CA).

DNA fragmentation assay

DNA fragmentation associated with apoptosis was examined by the terminal deoxynucleotidyltransferase (TdT)-mediated dUTP nick end labeling (TUNEL) method using a commercial assay kit (APO-Direct Kit; BD Pharmingen, San Diego, CA). After treatment, cells were fixed with 1% paraformaldehyde in PBS (4 °C, 30 min), washed in PBS and permeabilized with ice-cold 70% ethanol. The APO-Direct Kit TUNEL assay was performed as described by the manufacturer. Briefly, fixed cells were washed twice using the kit wash buffer, and after centrifugation, the supernatant was discarded. The DNA-labeling solution (containing TdT enzyme and FITC-dUTP) was added to the cell pellet, and the resuspended mixture was incubated for 1 h at 37 °C with occasional shaking. At the end of incubation time, rinse buffer was added, and after centrifugation, the supernatant was discarded. Cell rinsing is repeated with rinse buffer and then suspended the cell pellet in PI/RNase Staining Buffer. Cells were incubated for 30 min at room temperature and immediately analyzed in a BD FACSCanto II flow cytometer. In total, 10 000 events were collected per test sample. The results were analyzed in FACSDiva software (both from BD Bioscences Systems, San Jose, CA). The percentage of cells with distinctive apoptotic DNA strand breaks and distinguished by a green fluorescent emission was calculated.

Statistical analysis

All numerical data are presented as mean ± standard deviation (SD) from at least three independent experiments. Statistical analysis was conducted using the Origin 7.5 software (OriginLab, Northampton, MA). Statistical differences in multiple groups were determined by one-way analysis of variance (ANOVA) followed by Tukey’s test. p < 0.05 was considered statistically significant.

Results

Cytotoxicity assay

To evaluate the effect of Pt14–Pt16 on the viability of breast cells, MCF-7 and MDA-MB-231 breast cancer cells and fibroblasts skin cells were exposed to increasing concentrations of Pt14–Pt16 for 24 h (Figure 2) and 48 h (Figure 3), and the percentage of viable cells were determined in vitro using the MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide assay. Pt14–Pt16 decreased the cell viability in concentration-dependent manner in both MCF-7 and MDA-MB-231 breast cancer cells. Results revealed that Pt14–Pt16 exhibit substantially greater cytotoxicity than cisplatin against MCF-7 and MDA-MB-231 cells. The results demonstrated that the after 24 h of incubation IC₅₀ of Pt14–Pt16 was 45 ± 2 μM, 23 ± 3 μM and 72 ± 2 μM of MCF-7 cells and 42 ± 3 μM, 19 ± 3 μM and 67 ± 2 μM of MDA-MB-231 cells, respectively. After 24 h of incubation IC₅₀ values for the cisplatin alone in MCF-7 and MDA-MB-231 cells were 93 ± 2 and 82 ± 2 μM, respectively. Although, Pt14–Pt16 reduced the viability of breast cancer cells in a concentration-dependent manner, the viability of fibroblast cells was higher than breast cancer cells after exposure to Pt14–Pt16, indicating that breast cancer cells were more sensitive to Pt14–Pt16 than normal cells. For instance, IC₅₀ of Pt15 (the compound most active) of fibroblast cells was 70 ± 3 μM after 24-h incubation (Figure 2) and 34 ± 3 μM after 48-h incubation (Figure 3).

Figure 2. Viability of MCF-7 breast cancer cells, MDA-MB-231 breast cancer cells and fibroblast cells treated for 24 h with different concentrations of Pt14–Pt16 and cisplatin. Mean values ± SD from three independent experiment (n = 3) done in duplicate are presented.

Figure 3. Viability of MCF-7 breast cancer cells, MDA-MB-231 breast cancer cells and fibroblast cells treated for 48 h with different concentrations of Pt14–Pt16 and cisplatin. Mean values ± SD from three independent experiment (n = 3) done in duplicate are presented.

In case of incubation test, compounds by 48 h Pt14–Pt16 also exhibit substantially greater cytotoxicity than cisplatin against MCF-7 and MDA-MB-231 cells (Figure 3). Pt14–Pt16 and cisplatin decreased the number of viable cells in both breast cancer cell lines MCF-7 and MDA-MB-231 in dose-dependent manner. The most cytotoxic agent was Pt15 for which IC₅₀ was 14 ± 3 μM (MCF-7) and 12 ± 4 μM (MDA-MB-231). Pt14, Pt16 and cisplatin IC₅₀ were as follows: 25 ± 2 μM, 44 ± 2 μM, 78 ± 1 μM (MCF-7) and 23 ± 2 μM, 42 ± 2 μM and 72 ± 1 μM (MDA-MB-231), respectively. After 48 h of incubation, IC₅₀ value for the Pt14–Pt16 in fibroblast cells was 52 ± 2 μM, 34 ± 2 μM, 59 ± 2 μM and for cisplatin was above 100 μM.

Cytotoxic interaction of the test compounds may be the result of impaired biosynthesis of DNA. All tested compounds showed concentration-dependent activity, yet with different potency. Furthermore, the obtained profiles of DNA synthesis were similar in MCF-7 and MDA-MB-231 (Figures 4 and 5). The concentrations of compounds Pt14–Pt16 needed to inhibit [³H]thymidine incorporation into DNA by 50% (IC₅₀) after 24-h incubation in MCF-7 were found to be 72 ± 2 μM, 33 ± 3 μM, 67 ± 2 μM and in MDA-MB-231 52 ± 3 μM, 31 ± 3 μM, 59 ± 2 μM, respectively. That suggesting similar cytotoxic potency compared to cisplatin (IC₅₀ 98 ± 1 μM in MCF-7 and 86 ± 1 μM in MDA-MB-231 cells). After 48-h incubation, the concentrations of compounds Pt14–Pt16 and cisplatin needed to inhibit [³H]thymidine incorporation into DNA by 50% (IC50) in MCF-7 were found to be 30 ± 2 μM, 17 ± 3 μM, 34 ± 2 μM, 78 ± 2 μM and in MDA-MB-231 28 ± 3 μM, 19 ± 3 μM, 36 ± 2 μM, 68 ± 2 μM respectively. Tests on human fibroblast cells showed (both 24-h and 48-h incubation) that the compounds of Pt14–Pt16 are lower tendency to inhibition of DNA synthesis in human fibroblast cells compare to the breast cancer cells.

Figure 4. Antiproliferative effects of Pt14–Pt16 and cisplatin (treated for 24 h) in MCF-7 breast cancer cells, MDA-MB-231 breast cancer and fibroblast cells as measured by inhibition of [3H]thymidine incorporation into DNA. Mean values ± SD from three independent experiment (n = 3) done in duplicate are presented.

Figure 5. Antiproliferative effects of Pt14–Pt16 and cisplatin (treated for 48 h) in MCF-7 breast cancer cells, MDA-MB-231 breast cancer and fibroblast cells as measured by inhibition of [3H]thymidine incorporation into DNA. Mean values ± SD from three independent experiment (n = 3) done in duplicate are presented.

Pt14–Pt16 compounds induce apoptosis

Apoptosis was evaluated by staining with Annexin V, a phospholipid binding protein commonly used to detection of apoptosis. To characterize the mode of cell death induced by Pt14–Pt16, flow cytometric analysis was performed using annexin-V and propidium iodide (PI). Dual staining for annexin-V and PI permits discrimination between live cells (annexin-V–/PI–), early apoptotic cells (annexin-V+/PI–), late apoptotic cells (annexin-V+/PI+) and necrotic cells (annexin-V–/PI+) (Figure 6). As depicted in Figure 6, after 24 h of treatment with Pt14–Pt16 at different concentrations, we observed a significant, accumulation of annexin-V-positive cells in a concentration-dependent manner. All test compounds induced a dose-dependent apoptotic cell death. We have found that the apoptotic effect of Pt14–Pt16 was stronger than evoked by cisplatin. We noticed that the compound Pt15 in the broadest extent induced apoptosis in both cancer cell lines.

Figure 6. Flow cytometric analysis of MCF-7 and MDA-MB-231 breast cancer cells after incubation with Pt14–Pt16 and cisplatin (20 μM and 50 μM) for 24 h and subsequent staining with Annexin V and propidium iodide (PI). Dots with Annexin V−/PI − (Q3), Annexin V+/PI − (Q4), and Annexin V+/PI + (Q2) feature represent intact, early apoptotic, and dead cells, respectively. Mean percentage values from three independent experiments (n = 3) done in duplicate are presented.

Furthermore, fluorescence microscopy using double staining acridine orange and ethidium bromide staining also showed condensation of chromatin, fragmented nuclei and nuclear shrinkage (Figure 7). In this case, green intact nuclei indicated normal cells, while as orange staining indicated apoptosis. Bright spots in the Pt14–Pt16-treated cells indicated nuclei undergoing chromatin condensation, strongly suggesting that all breast cancer cells underwent apoptosis. Figure 7 shows that the acridine orange stained cells were evident in both breast cancer cells treated with Pt14–Pt16, as compared to untreated controls. The results showed that the compounds Pt14–Pt16 and cisplatin induces apoptotic cell death, which is consistent with the results observed in the above annexin-V and PI.

Figure 7. Induction of apoptosis in MCF-7 and MDA-MB-231 breast cancer cells treated for 24 h with 20 μM and 50 μM of Pt14–Pt16 and cisplatin evaluated by a fluorescent microscopy assay after acridine orange and ethidium bromide staining. Mean percent ± SD from three independent experiments are presented. Apoptotic and necrotic cells were differentiated according to the criteria described in Results.

Pt14–Pt16 decrease mitochondrial membrane potential

To investigate the cellular mechanism underlying Pt14–Pt16 induced intrinsic apoptosis in breast cancer cells, we assessed the alterations of mitochondrial transmembrane potential (ΔΨ_m) by using flow cytometry analysis. We found that Pt14–Pt16 decreased the levels of ΔΨ_m in breast cancer cells at 24 h (Figure 8). Mitochondria play an essential role in the propagation of apoptosis15,16. It is well established that, at an early stage, apoptotic stimuli alter the mitochondrial transmembrane potential (ΔΨ_m). Mitochondrial transmembrane potential was monitored by the fluorescence of the dye JC-1. In untreated cells (high ΔΨ_m), JC-1 displays a red fluorescence (590 nm). This is caused by spontaneous and local formation of aggregates that are associated with a large shift in emission. In contrast, when the mitochondrial membrane is depolarized (low ΔΨ_m), JC-1 forms monomers that emit at 530 nm. As shown in Figure 8, Pt14–Pt16 in MCF-7 and MDA-MB-231 cells induced a concentration-dependent increase in the proportion of cells with depolarized mitochondria. These results are in accord with those obtained in the annexin-V and PI assay.

Figure 8. Fluorescence of MCF-7 and MDA-MB-231 breast cancer cells treated for 24 h with 20 μM and 50 μM of Pt14–Pt16 and cisplatin incubated with mitochondrial membrane potential probe JC-1. X- and y-axes are green and red fluorescence, respectively. Pt14–Pt16-treated cells show loss of red fluorescence (loss of membrane potential-dependent accumulation in mitochondria) indicative of mitochondrial membrane depolarization. Mean percentage values from three independent experiments (n = 3) done in duplicate are presented.

Pt14–Pt16 induce caspase activation and caspase cascade

Caspases, which are proteolytic enzymes, are the central executioners of apoptosis, and their activation is mediated by various inducers17. Synthesized as proenzymes, caspases are themselves activated by specific proteolytic cleavage reactions. Caspase-8, caspase-9 and caspase-10 are termed apical caspases and are usually the first to be activated in the apoptotic process. Following their activation, they in turn activate effector caspases, in particular caspase-318. Following the treatment of MCF-7 and MDA-MB-231 cells with compounds Pt14–Pt16 (20 μM and 50 μM), we observed activation of caspase-8 at 24 h (Figure 9). Caspase-8 protein levels were increased Pt14–Pt16-treated breast cancer cells, as compared to untreated control (Figure 9). The highest concentration of caspase-8 was observed after Pt15 treatment at dose 50 μM. According to these results, we hypothesized that the death receptor-mediated apoptotic pathway may be preferred in breast cancer cells. We observed the statistically significant increase in caspase-9 concentration above the control value after 24 h of incubation with Pt14–Pt16 and cisplatin (Figure 10). The highest concentration of caspase-9 was detected after Pt15 treatment. We also observed activation of caspase-3 at 24 h (Figure 11). These results are in good agreement with the mitochondrial depolarization described earlier, indicating that Pt14–Pt16 induces the apoptotic intrinsic (mitochondrial) apoptosis pathway.

Figure 9. Flow cytometric analysis of populations MCF-7 and MDA-MB-231 breast cancer cells treated for 24 h with 20 μM and 50 μM of Pt14–Pt16 and cisplatin for active caspase-8. Mean percentage values from three independent experiments (n = 3) done in duplicate are presented.

Figure 10. Flow cytometric analysis of populations MCF-7 and MDA-MB-231 breast cancer cells treated for 24 h with 20 μM and 50 μM of Pt14–Pt16 and cisplatin for active caspase-9. Mean percentage values from three independent experiments (n = 3) done in duplicate are presented.

Figure 11. Flow cytometric analysis of populations MDA-MB-231 breast cancer cells treated for 24 h with 20 μM and 50 μM of Pt14–Pt16 and cisplatin for active caspase-3. Mean percentage values from three independent experiments (n = 3) done in duplicate are presented.

Caspase-3 is also activated by the action of caspase-8 and caspase-10. At the same time, increase in the expression of caspase-8 and casepase-10 suggests the fact that it has been activated external apoptotic pathway19. After 24 h of treatment with Pt14–Pt16, activation of caspase-8 and caspase-10 was observed (Figures 9 and 12).

Figure 12. Flow cytometric analysis of populations MCF-7 and MDA-MB-231 breast cancer cells treated for 24 h with 20 μM and 50 μM of Pt14–Pt16 and cisplatin for active caspase-10. Mean percentage values from three independent experiments (n = 3) done in duplicate are presented.

Detection of apoptosis by DNA fragmentation

DNA fragmentation is a key feature of the apoptosis process, largely due to the activation of endogenous endonucleases (such as caspases) which subsequently induce cleavage of chromatin DNA into internucleosomal fragments of roughly 180 base pairs20. Monitoring the pattern of DNA fragmentation by TUNEL assay can be a qualitative way to assess apoptosis. Under normal growth conditions, any measurable effects on TUNEL staining were not observed (Figure 13). After treatment with Pt14–Pt16 (20 μM and 50 μM) for 24 h, the number of TUNEL-positive cells dose dependently increased, suggesting that Pt14–Pt16 treatment increased fragmentation DNA in MCF-7 and MDA-MB-231 cells, which is a hallmark of apoptosis. To determine whether Pt14–Pt16 can induce apoptosis in breast cells, cells were treated with different concentrations of Pt14–Pt16 for 24 h and then percentage of DNA fragmentation was quantified. Pt14–Pt16 significantly induced DNA fragmentation in both breast cancer cells. As shown in Figure 13, DNA fragmentation in breast cancer cells in the presence of Pt15 is 2.5 times higher in comparison with cisplatin. Even though, the Pt14–Pt16-treated fibroblast cells underwent apoptosis, percentage of DNA fragmentation was lower compared to all of the breast cancer cells (data not shown).

Figure 13. Flow cytometric analysis of DNA fragmentation of MCF-7 and MDA-MB-231 breast cancer cells after 24 h of incubation with Pt14–Pt16 and cisplatin (20 μM and 50 μM) using TUNEL assay. Histograms present TUNEL negative and TUNEL positive cells. Mean percentage values from three independent experiments (n = 3) done in duplicate are presented.

Discussion

Human breast cancer cell lines provide an excellent platform for in vitro breast cancer research studies and widely used as experimental models. Our studies have shown that the compounds of Pt14–16 have a higher cytotoxicity in comparison with cisplatin in relation to MCF-7 and MDA-MB-231 breast cancer cells. The highest cytotoxicity showed Pt₂(3,4-dimethylpyridine)₄(berenil)₂ (Pt15). In case of human skin fibroblast cell, cytotoxicity assays demonstrated that these compounds are less toxic to normal cells than cisplatin. The results are consistent with previous observations that berenil–platinum complexes have a high cytotoxicity against tumor cells and a moderate compared to normal cells21,22. Cytotoxic interaction of the test compounds may be the result of impaired biosynthesis of DNA. Previous studies on berenil complexes of platinum showed the ability of these compounds to inhibit the biosynthesis of DNA in tumor cells21–24. This study showed that Pt14–Pt16 inhibited more DNA biosynthesis than cisplatin in MCF-7 and MDA-MB-231 breast cancer cells. After 24 h of incubation breast cancer cells with test compounds, the most active compound was Pt15. Tests on human skin fibroblasts showed that the compounds of Pt14–Pt16 substantially less ability to inhibit DNA in normal cells in compare to tumor cell lines. In summary, we conclude that Pt14–Pt16 may be compounds with improved therapeutic properties relative to the tumor cell and less toxicity to normal cells as compared to cisplatin.

IC₅₀ values of all three platinum complexes were very similar and all complexes showed significantly higher cytotoxic effect on breast cancer cells than cisplatin. Comparison of IC₅₀ values of compounds Pt14-Pt16 with those of their previously obtained analogs reveals that when the amine ligand is varied the level of cytotoxicity increases in the order: 3-cyanopyridine < 3-hydroxymethylpyridine < pyridine < piperidine < aliphatic amines < monoalkylpyridine ≈ dimethylpyridine. The antitumor evaluation of the berenil–platinum complexes on cancer cells indicates that the insertion of the alkyl substituents on the pyridine ring increases the cytotoxic potency in almost all the evaluated cell lines. In terms of substitution patterns in the pyridine complexes, the following general observations were established, albeit qualitatively. For the mono alkyl groups, potency varied 3-Me > 3-Bu > 4-(t-Bu) > 4-Me > 3-Et > 4-Et > 2-Me21–24. The reason for this correlation is unclear and appears to not be related to either the DNA binding strength nor the DNA adduct formed. While some of the cytotoxicity is probably due to DNA binding, it is possible that the compounds their activity either from a different target or through other means. It is also possible that the different conformations of the ligands drive factors such as cellular uptake or drug transport within the cell. For instance, human serum albumin has been shown to play a role in the transport of other platinum drugs. The structure–activity relationships determined in this study are, however, only preliminary since they are based on only two cell lines, sensitive to cisplatin and therefore testing in further cell lines, both sensitive and resistant to cisplatin, need to be completed before the structure–activity relationships are confirmed. Taking into account the high activity of Pt14-Pt16 it cannot be excluded that the formation of active metabolites plays an important role in the mechanism of action.

Polynuclear platinum complexes constitute a novel class of prospective anticancer agents that have shown some peculiar activities as compared with mononuclear platinum compounds25–28. Farrell et al. synthesized a series of multinuclear platinum(II) complexes, which form DNA adducts that differ markedly in structure, sequence specificity and formation kinetics from those generated by cisplatin and its mononuclear analogs. More importantly, they display high antitumor activity in vitro and in vivo against cisplatin-sensitive and cisplatin-resistant tumor cell lines. The increased cytotoxicity is explained with the ability for precovalent association and the formation of long-range adducts with DNA. However, clinical trials show that the trinuclear complex BBR3464, and other multinuclear platinum drugs, did not yield results substantially different from cisplatin, possibly due to their binding and degradation by human plasma proteins. Therefore, research efforts have been directed to reduce the high irreversible plasma protein binding and improve the chemical and metabolic stability, as well as to decrease the toxic effects that led to failure in clinical trials. The dinuclear berenil–platinum complexes with amine ligands are cationic in nature and show excellent solubility in water. The adducts formed by these dinuclear platinum complexes are vastly different from the adducts formed by cisplatin29. It has been suggested that the distortions induced by these complexes are only weakly recognized by DNA repair proteins. Berenil (1,3-bis(4′-aminophenyl)triazene) forming part of these complexes is a compound belonging to the class of aromatic bisamidine. Bisamidines have a wide spectrum of biological activity, which covers properties antiparasitic, antiviral and anticancer properties30,31. In analyzing the structure–activity relationship of the dinuclear complexes, it was seen that berenil provided H-bonding and an electrostatic preassociation with duplex DNA in the minor groove. Through an increased local concentration at particular sites on DNA, the precovalent binding association can be used to control the site of platination. The berenil complexes of platinum to interact with nucleic acids penetrating deeply into the small groove of DNA to form hydrogen bonds between the amidine groups of berenil, and thymine and/or adenine nucleic acid. Berenil–platinum complexes have a lower affinity to the sequence of guanine-cytosine due to the presence of the amino group of guanine, which sterically hindered the formation of such calls32,33. The berenil–platinum complexes differentiate this from other alkylating agents, which primarily relate to the major groove of DNA. Structurally novel platinum complexes that bind to DNA differently than cisplatin may have distinct cytotoxicity and side effect profiles26,28. While it has been clearly established that the dinuclear platinum complexes can form an array of different DNA adducts to cisplatin, the challenge now is to determine which of the particular adducts lead to the enhanced cytotoxicity. In addition, there is considerable emphasis on developing new dinuclear platinum complexes that target-specific biomolecules and cellular pathways.

A flow cytometric analysis of breast cancer cells was carried out to further explore the ability of the tested agents to induce apoptosis. Apoptotic pathways play an important role in the pathogenesis and progression of breast cancer. Two major pathways either in alone or in crosstalk are involved in driving the cells to apoptosis. First one is death receptor (extrinsic) pathway including Fas/FasL, tumor necrosis factor (TNF)/TNF receptors, and death receptors 4 (DR4) and 5 (DR5) and their downstream molecules such as caspases-8. The other one is the mitochondria-mediated (intrinsic) pathway characterized by the loss of mitochondrial membrane potential (MMP), release of cytochrome c, Smac/DIABLO, Htra/Omi, Apaf-1, and activation of procaspase-9 and effector caspases. Both breast cancer cells underwent apoptosis in presence of Pt14–Pt16.

Several studies demonstrated that cisplatin-induced cytotoxicity culminates in the activation of apoptosis in cancer cells34–36. The activation of apoptosis occurs through a mitochondrial membrane potential change and caspase-3 activation37,38. Our results support these findings showing that Pt14–Pt16 induces apoptosis in MCF-7 and MDA-MB-231 cells through changing mitochondrial membrane potential. The implication of the decrease mitochondrial membrane potential and increased permeability of the mitochondrial membrane is the release of cytochrome c39. Then, it activates caspase-9 and then caspase-3 and caspase-7, leading to the activation of caspase cascade and apoptosis40,41. According to Kojima and coauthors, cisplatin may cause the release of mitochondrial cytochrome c leading to activation of caspase-9 and caspase-342. It was also found that the compounds of Pt14–Pt16 greater than cisplatin affect the growth of the cell population with active caspase-9. These results are consistent with the results of testing the expression of caspase-3. No caspase-3 in MCF-7 breast cancer cells indicates that apoptosis in these cells is activated by other transmitters in the suppressor protein p53, NF-kB or TNF-α43–45.

In recent years, it has been found that cisplatin also can induce apoptosis by activates the extrinsic pathway activated by members of the tumor necrosis factor (TNF) super family such as FAS receptor46. FAS can trigger apoptosis through its FAS ligand (FASL), upon engagement of the FAS-associated death domain protein (FADD), and recruitment of procaspase-8 and procaspase-10 to the intracellular death domain of FAS receptor form the death-inducing signaling complex (DISC) that either directly initiates a downstream caspase cascade47,48. As a result of our research, we showed that the Pt14–Pt16 give rise to expression of active caspase-8 and active caspase-10 in MCF-7 and MDA-MB 231 breast cancer cells. This may suggest that Pt14–Pt16 complexes also induce programed cell death by caspase-dependent pathway of caspase-8 and caspase-10. In addition, these observations can be combined with the result of a study illustrating that the Pt14–Pt16 affect the increase in the expression of active caspase-3. It plays a key role in apoptosis, participating in the creation of chromatin fragmentation and DNA damage49. We also observed that Pt14–Pt16 induced apoptosis in the MCF-7 and MDA-MB-231 cells, which was characterized by marked DNA fragmentation. Investigation of molecular mechanisms mediating the anticancer effect of Pt14–Pt16 will provide valuable information for the further development of novel strategies for cancer treatment. Apoptosis induction, possibly enhanced by a contribution of targets other than DNA, seems to be an important factor in the mechanism of action of Pt14–Pt16. The cytotoxic effects of Pt14–Pt16 cause mitochondrial dysfunction and activation of the caspase cascade. They might induce apoptosis by extrinsic and intrinsic pathways. In order to design better dinuclear berenil–platinum complexes, the mechanism of action of our drugs needs to be better explored. To this end, our group is conducting further biochemical studies involving cellular uptake, DNA binding experiments, further cytotoxicity assays with cell lines with differing modes of resistance to cisplatin, and we hope to report the results in the near future.

Conclusion

In this study, we tested the effect of new dimethylpyridine–platinum complexes (Pt14–Pt16) in MCF-7 and MDA-MB-231 breast cancer cells. Evaluation of the cytotoxicity of Pt14–Pt16 in both MDA-MB-231 and MCF-7 breast cancer cells demonstrated that these compounds were more potent antiproliferative agents than cisplatin. Moreover, the novel compounds displayed less pronounced inhibitory activity against cultured human skin fibroblasts, as compared to cisplatin. Further analyses found that Pt14–Pt16 induced caspase-dependent apoptosis by activating the mitochondria-associated intrinsic apoptosis pathway. Our data demonstrated that Pt14–Pt16 also induced apoptosis through extrinsic pathway with caspase activation. These findings indicate that Pt14–Pt16 are endowed with high antiproliferative activity and should be further investigated to determine their therapeutic potential.

Declaration of interest

This research was supported by Medical University of Bialystok (Grant N/ST/ZB/15/002/2217). This study was conducted with the use of equipment purchased by Medical University of Bialystok as part of the OP DEP 2007–2013, Priority Axis I(0).3, contract No. POPW.01.03.00–20-022/09. The authors report no conflicts of interest. The authors alone are responsible for the content and writing of this article.