MCF-7 human breast cancer cells from the American Type Culture Collection (ATCC, Manassas, VA, USA) were maintained in RPMI-1640 (Gibco, Grand Island, NY, USA) supplemented with 10% fetal calf serum (HyClone, Logan, UT, USA), 100 U/ml penicillin, and 100 mg/ml streptomycin at 37°C in a humidified atmosphere containing 5% CO₂. The cells were passaged every two to three days to maintain exponential growth.

Cantharidin, PD98059, SP600125, GF109203X and Ro 31-8220 were purchased from Enzo Life Sciences International Inc., (Plymouth Meeting, PA, USA). Norcantharidin was purchased from Sigma (St. Louis, MO, USA).

Cellular growth was evaluated by the MTT (3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyltetrazolium bromide) assay (10). The cells were seeded into 24-well tissue culture plates at 5×10⁴ cells/well. Following treatment, MTT (Sigma) was added to each well at a final concentration of 0.5 mg/ml, followed by incubation at 37°C for 4 h. The medium was then removed and 800 μl dimethyl sulfoxide was added to each well. The absorbance of the mixture was measured at 490 nm using a microplate ELISA reader (Bio-Rad Laboratories, Hercules, CA, USA). The inhibition rate was calculated as follows: inhibition rate = [(mean control absorbance − mean experimental absorbance)/mean control absorbance] × 100%. To evaluate the effect of cantharidin and norcantharidin on cellular growth, the concentration that caused 50% growth inhibition (IC₅₀) was calculated, as previously described (7).

The cells were seeded at a density of 200 cells/well in 24-well plates and treated 12 h later. After 15 days, the cells were stained with 1% methylrosanilinium chloride and the numbers of visible colonies were counted. The relative clone formation ability was calculated as relative clone formation ability = (mean experimental clone number/mean control clone number) × 100%.

Apoptosis was evaluated using the Annexin V-FITC/PI Apoptosis Detection kit (Biouniquer Technology, Nanjing, China) according to the manufacturer’s instructions. The cells were resuspended in binding buffer, and Annexin V-FITC and propidium iodide (PI) were added to the buffer and incubated at room temperature for 15 min in the dark, followed by flow cytometry using a Beckman Coulter FC500 dual-laser five-color flow cytometer (Beckman Coulter, Fullerton, CA, USA).

The cells were resuspended in complete medium and seeded in 24-well plates at a concentration of 1×10⁴ cells/ml. After a 5-h incubation, the unattached cells were removed to another well. The attached cells and unattached cells were evaluated using the MTT assay. The adhesion rate was calculated as: [absorbance of attached cells/(absorbance of attached cells + absorbance of unattached cells)] × 100%.

The cells were seeded in 96-well plates at a density of 1×10⁴ cells/well and grown to confluence. The monolayer culture was then artificially scrape-wounded with a sterile micropipette tip to create a denuded zone (gap) of constant width. Each well was washed with phosphate-buffered saline (PBS) twice to remove the detached cells before treatment. Cells that had migrated to the wounded region were observed using an XDS-1B inverted microscope (Chongqing Optical & Electrical Instrument Co., Ltd., Chongqing, China) and photographed (x40 magnification). Images were captured every 4 h to monitor the wound healing process. The wound areas were measured using ImageJ (National Institutes of Health, Bethesda, MA, USA).

Fresh blood obtained from healthy volunteers was anticoagulated with a 1/7 volume of acid-citrate dextrose (85 mM trisodium citrate, 110 mM dextrose and 78 mM citric acid) as previously described (11). The Ethics Committee of the First Affiliated Hospital of Soochow University, Suzhou, China, approved the study. Platelet-rich plasma was collected after centrifugation at 1,300 rpm for 13 min. Then, CFDA-SE (carboxyfluorescein diacetate succinimidyl ester; Beyotime, Shanghai, China) was added at a final concentration of 5 μM and incubated at room temperature for 10 min. The fluorescence-labeled platelets were then washed twice with CGS buffer (0.12 M sodium chloride, 0.0129 M trisodium citrate and 0.03 M D-glucose, pH 6.5), resuspended in freshly prepared Tyrode’s buffer (11), and allowed to rest for at least 1 h at 37°C prior to use.

The traditional platelet adhesion assay was performed as previously described (12). In brief, 5×10⁴ cells/ml MCF-7 cells were seeded in a 96-well plate and grown until confluent. Washed platelets preloaded with CFDA-SE were allowed to adhere to the cells for 30 min at 37°C. The non-adherent platelets were discarded. Platelet adhesion to the MCF-7 cells was imaged by Olympus IX51 inverted fluorescence microscopy (Olympus, Tokyo, Japan). The flow cytometry-based platelet adhesion assay was performed as follows: the cells were harvested and resuspended in RPMI-1640 containing 1 mM CaCl₂, 1 mM MgCl₂ and 0.1% bovine serum albumin. One hundred microliters of cells (5×10⁶ cells/ml) were incubated with 400 μl CFDA-SE-labeled platelets for 30 min. Then, 4% paraformaldehyde (250 μl) was added to a final concentration of 2%. The cells were fixed for 1 h at 4°C, followed by flow cytometry analysis.

Total RNA was extracted using TRIzol reagent (Invitrogen, Carlsbad, CA, USA) according to the manufacturer’s protocol. After spectrophotometric quantification, 1 μg total RNA was used for reverse transcription in a final volume of 20 μl with AMV reverse transcriptase (Promega, Madison, WI, USA) according to the manufacturer’s instructions. Aliquots cDNA of corresponding to equal amounts of RNA were used for the quantification of mRNA by real-time PCR using the TL988-II Real-time Quantitative PCR Detection system (TianLong Science And Technology, Co., Ltd., Shaanxi, China). The reaction system (25 μl) contained the corresponding cDNA, forward and reverse primers, and SYBR Green PCR Master Mix (Roche Diagnostics, Indianapolis, IN, USA). All data were analyzed using β-actin gene expression as an internal standard. The specific primers were: i) integrin α2, forward, 5′-CTGGAGTGGCTTTCCTGAG-3′ and reverse, 5′-ACTG ATTCCCACATTGCTG-3′, product, 224 bp; ii) integrin β1, forward, 5′-GAGATGGGAAACTTGGTGG-3′ and reverse, 5′-GACAAGGTGAGCAATAGAAGGAT-3′, product, 113 bp; iii) p65, forward, 5′-GAGCCGCACAGCATTCAGG-3′ and reverse, 5′-CGCTGCATCCACAGTTTCCA-3′, product, 156 bp; and iv) β-actin, forward, 5′-TCATGAAGTGTGA CGTGGACAT-3′ and reverse, 5′-CTCAGGAGGAGCAA TGATCTTG-3′, product, 158 bp.

The abundance of α2 integrin on the surface of the MCF-7 cells was measured by flow cytometry. Following treatment, the cells were incubated with saturating concentrations (10 μg/ml) of anti-α2 integrin antibody clone P1E6 for 30 min at room temperature. Subsequently, the cells were incubated with 10 μg/ml fluorescein-isothiocyanate-conjugated anti-mouse antibody for 30 min at room temperature and were analyzed using a Beckman Coulter FC500 dual-laser five-color flow cytometer (Beckman Coulter).

Target-specific small interfering RNAs (siRNAs) were designed and synthesized by GenePharma (Shanghai, China). The specific sequences were as: i) control-siRNA, sense, 5′-UUCUCCGAACGUG UCACGUdTdT-3′ and antisense, 5′-ACGUGACACGUUCG GAGAAdTdT-3′; ii) p65-siRNA-1, sense, 5′-CGGAUUGAG GAGAAACGUAdTdT-3′ and antisense, 5′-UACGUUUCUC CUCAAU CCGdTdT-3′; iii) p65-siRNA-2, sense, 5′-GGAGU ACCCUGAGGCUAUAdTdT-3′ and antisense, 5′-UAUAGC CUCAGGGUACUCCdTdT-3′. The transfections were performed with the siRNA-Mate Transfection reagent (GenePharma) according to the manufacturer’s instructions.

Each experiment was performed in triplicate, at minimum. The results are expressed as the means ± standard deviation. Statistical analysis was performed using unpaired Student’s t-tests; P<0.05 was considered to indicate a statistically significant different.

First, we investigated the effects of cantharidin and norcantharidin on the growth, adhesion and migration of the MCF-7 cells. The inhibitory effects of cantharidin and norcantharidin on MCF-7 cell growth were first evaluated using the MTT assay. As presented in Fig. 1A, cantharidin or norcantharidin treatment inhibited MCF-7 cell growth in a dose- and time-dependent manner, and the IC₅₀ values at 72 h after treatment were 11.96 and 105.34 μM, respectively. The dose-dependent inhibitory effect of cantharidin and norcantharidin on the growth of MCF-7 cells was also confirmed by clone-formation assays (Fig. 1B).

We performed apoptosis assays to further investigate the mechanism involved in the growth inhibition effect. The percentages of cell populations at various stages of apoptosis are shown in Fig. 1C. After cantharidin or norcantharidin treatment, the number of cells that underwent early apoptosis (Annexin V+/PI−) increased significantly in a dose-dependent manner. These data suggested that the growth-inhibition effect of cantharidin and norcantharidin could be due to induction of apoptosis.

The effects of cantharidin and norcantharidin on the adhesion and migration ability of MCF-7 cells were also evaluated. Cantharidin or norcantharidin dose-dependently repressed the adhesion ability of MCF-7 cells (Fig. 1D) and repressed cell migration in a dose- and time-dependent manner (Fig. 1E).

Thus, the above data indicate inhibition of multiple biological behaviors of MCF-7 cells following cantharidin and norcantharidin treatment.

We then focused on the effect of cantharidin and norcantharidin on the adhesion of MCF-7 cells to platelets. First, we observed the appearance of the cells following 24-h cantharidin and norcantharidin treatment. As shown in Fig. 2A, the untreated cells were confluent, whereas several gaps were observed in the cantharidin or norcantharidin-treated groups. This could have been due to the decreased cell number induced by growth inhibition, as well as the detachment triggered by the decreased adhesion ability and apoptosis as presented in Fig. 1. Thus, when the traditional platelet adhesion assay (12) was performed, the platelets adhered directly to the plate through the gaps (Fig. 2A). Therefore, the fluorescent intensity did not accurately reflect the actual ability of the MCF-7 cells to adhere the platelets.

Consequently, we developed a platelet adhesion assay based on the flow cytometry. As shown in Fig. 2B, the fluorescent positive rate was increased when the dose of platelets was increased. When incubated with 10×10⁶/ml platelets, few cells were covered by the platelets, while the adhesion became saturated when the platelet concentration was increased to 100×10⁶ cells/ml. Thus, a moderate concentration of 30×10⁶ cells/ml was used for further investigation.

As shown in Fig. 2C, the fluorescent rate was repressed when the cells were treated with cantharidin or norcantharidin, suggesting that cantharidin and norcantharidin could inhibit the adhesion between the MCF-7 cells and platelets. Thus, anticancer therapy using cantharidin and norcantharidin might be able to repress the platelet-mediated hematogenous metastasis potential of breast cancer cells.

α2β1 integrin, a heterodimeric transmembrane receptor composed of the α2 and β1 subunits, is expressed by some tumor cells to facilitate binding to the extracellular matrix. Recent studies have suggested that this adhesion molecule could be involved in the interaction between cancer cells and platelets (13). Therefore, we investigated whether the regulation of α2β1 integrin expression could facilitate the inhibition of cancer cell-platelet adhesion induced by cantharidin and norcantharidin treatment.

As presented in Fig. 3A, the mRNA expression of α2 integrin was downregulated following treatment with cantharidin or norcantharidin, while the expression of β1 integrin was not affected. The abundance of α2 integrin on the cell surface was further evaluated using flow cytometry. As shown in Fig. 3B, the distribution of α2 integrin on the cell surface was decreased upon cantharidin or norcantharidin treatment.

As cantharidin and norcantharidin treatment repressed the expression of α2 integrin, we subsequently investigated whether the inhibition of MCF-7 cell adhesion to platelets was executed due to downregulation of α2 integrin. As presented in Fig. 3C, P1E6, the function-blocking antibody to α2 integrin, repressed the adhesion between the MCF-7 cells and platelets, suggesting that α2 integrin mediated the interaction between the MCF-7 cells and the platelets. Moreover, pretreatment with P1E6 repressed the decline of the cantharidin and norcantharidin-treated MCF-7 cell adhesion to platelets, suggesting that cantharidin and norcantharidin repress the adhesion of MCF-7 cells to platelets by downregulating α2 integrin.

Mechanistically, cantharidin and norcantharidin have been shown to be potent and selective inhibitors of protein phosphatase 2A (PP2A) (14), a multimeric serine/threonine phosphatase that can dephosphorylate several kinases (15,16). In our previous studies (7–9), we found that extracellular signal-regulated kinase (ERK), c-Jun N-terminal kinase (JNK), IκB kinase (IKK) and protein kinase C (PKC) were activated following treatment with PP2A inhibitors. To investigate whether these pathways were involved in the downregulation of α2 integrin, we tested the expression of α2 integrin following cantharidin and norcantharidin treatment when these pathways were blocked (Fig. 4A).

Pretreatment with PD98059, an ERK pathway inhibitor, or SP600125, a JNK pathway inhibitor, did not affect the downregulation of α2 integrin in cantharidin or norcantharidin-treated cells (Fig. 4B and C). Blocking the nuclear factor κB (NF-κB) by transfection of a siRNA-targeting p65 (Fig. 4D) did not affect the downregulation of α2 integrin either (Fig. 4E). However, pretreatment with the PKC inhibitors GF109203X or Ro 31-8220 attenuated the downregulation of α2 integrin (Fig. 4F). Ro 31-8220 even reversed the effects of cantharidin and norcantharidin on α2 integrin expression, indicating that cantharidin and norcantharidin repress α2 integrin expression through the PKC pathway.