Cantharidin (CTD) is an active compound isolated from the traditional Chinese medicine blister beetle and displayed anticancer properties against various types of cancer cells. However, little is known about its effect on human chronic myeloid leukemia (CML) cells, including imatinib-resistant CML cells. The objective of this study was to investigate whether CTD could overcome imatinib resistance in imatinib-resistant CML cells and to explore the possible underlying mechanisms associated with the effect. Our results showed that CTD strongly inhibited the growth of both imatinib-sensitive and imatinib-resistant CML cells. CTD induced cell cycle arrest at mitotic phase and triggered DNA damage in CML cells. The ATM/ATR inhibitor CGK733 abrogated CTD-induced mitotic arrest but promoted the cytotoxic effects of CTD. In addition, we demonstrated that CTD downregulated the expression of the BCR-ABL protein and suppressed its downstream signal transduction. Real-time quantitative PCR revealed that CTD inhibited BCR-ABL at transcriptional level. Knockdown of BCR-ABL increased the cell-killing effects of CTD in K562 cells. These findings indicated that CTD overcomes imatinib resistance through depletion of BCR-ABL. Taken together, CTD is an important new candidate agent for CML therapy.
Chronic myeloid leukemia (CML) is a myeloproliferative disorder caused by the constitutively active tyrosine kinase BCR-ABL, the result of the reciprocal chromosomal translocation (9:22) (q34;q11) (Deininger et al., 2000; Pane et al., 1996). BCR-ABL abnormally activates multiple signal pathways, including JAK-STAT, MAPK-ERK, and PI3K pathways, leading to deregulated proliferation and malignant transformation of CML cells (Danial and Rothman, 2000; Gesbert et al., 2000; Lugo et al., 1990). Targeting BCR-ABL with tyrosine kinase inhibitor (TKI), imatinib, has revolutionized the treatment of CML patients (Baselga and Arribas, 2004; Deininger et al., 2005). Unfortunately, about 20–30% of chronic phase CML patients are unresponsive to imatinib, owing to innate or acquired resistance, which has been a great challenge for CML therapy (Hochhaus et al., 2009). Although the second and third generations of tyrosine kinase inhibitors have been developed, their efficacy and safety in clinical use still need to be further investigated (Bixby and Talpaz, 2010). Therefore, exploring alternative treatment strategies is an urgent need.
Cantharidin (CTD) is a type of terpenoid extracted from the traditional Chinese medicine mylabris (
The detailed mechanisms underlying the cell cycle arrest by CTD remain unclear. Furthermore, the therapeutic potential of CTD in imatinib-resistant CML cells has not been evaluated until now. Therefore, the aim of this study was to investigate whether CTD could overcome imatinib resistance in human CML cells and to explore the mechanisms underlying this activity.
CTD (98% or higher purity) was purchased from Shanghai Shifeng Biological Technology Company (China). CTD was dissolved in DMSO as a 40 mM stock solution and stored at − 20°C. Primary antibodies against pCdc2, Cdc2, Cdc25c, cyclin B1, cyclin D1, pSTAT5, STAT5, pAKT, AKT, pERK1/2, ERK1/2, PARP1, cPARP, and pH3 (Ser10) were purchased from Cell Signal Technology, Inc. (USA). Antibodies against GAPDH and BCR were purchased from Santa Cruz Biotechnology (USA). Antibody against γH2AX was from EMD Millipore (USA). Imatinib and CGK733 were purchased from Selleck (USA). Caffeine was purchased from Amquar Biology (AMQUAR Bio., USA). RPMI 1640 medium, fetal bovine serum (FBS), penicillin G and streptomycin were provided by Gibco (USA). All the other reagents used in this study were of analytical grade.
Human CML cell line K562 was purchased from Cell Bank of Shanghai Institute of Biochemistry and Cell Biology. K562 cells were cultured in RPMI 1640 medium supplemented with 10% FBS. The imatinib-resistant cell line, K562R, was kindly provided by Guangbiao Zhou (Institute of Zoology, Chinese Academy of Sciences, China). K562R cells were cultured in RPMI 1640 medium supplemented with 10% FBS, and 1 μM imatinib, which was removed before experiments with a wash-out period of 2 to 3 days.
Normal blood PBMCs were isolated from healthy donors by density gradient centrifugation by Ficoll paque plus (GE Healthcare Life Sciences, Marlborough, USA) density sedimentation, followed by two washes in 1 × phosphate buffered saline. Cells were then cultured in liquid culture (RPMI1640, supplemented with 20% FBS). Use of the PBMC samples was approved by the Institutional Review Board of Committee of Jiangsu Province Academy of Traditional Chinese Medicine.
Cells were seeded into a 96-well plate at a density of 1 × 104 cells/well, pre-cultured for 24 h, and then treated with CTD at various concentrations (0, 5, 10, 20, 40, or 80 μM) for 24 or 48 h. Cell Counting Kit-8 (CCK-8) (Dojindo, Japan) was used to evaluate cell proliferation. Briefly, the medium from each well was removed after CTD treatment and 100 μl of fresh serum-free medium with 10 μl of CCK-8 was added. The absorbance was measured at 450 nm after further incubation for 2 h at 37°C. Cell death was assessed by trypan blue dye exclusion test. After CTD treatment, cells were incubated with 0.4% trypan blue solution diluted with PBS. Stained cells and unstained cells were counted in a Neubauer chamber under microscope.
Cells were collected and lysed with RIPA buffer containing protease inhibitor cocktail. The lysates were centrifuged and the supernatant was collected. Total proteins in the cells were quantitated by BCA protein assay, separated by sodium dodecyl sulfate polyacrylamide gel electrophoresis, and subsequently transferred onto a PVDF membrane. The membrane was blocked with 5% skimmed milk in TBS-T for 1 h at room temperature, then incubated with primary antibodies at 4°C overnight, followed by incubation with IRDye conjugated secondary antibody. All the antibodies were diluted in 5% skimmed milk with TBS-T. The primary antibodies were diluted 1:500 – 1:1000 and the secondary antibodies were diluted 1:5000. Odyssey infrared fluorescent scanner (LI-COR) was used for detecting the relevant proteins.
The cells were exposed to CTD at indicated concentrations for 24 h and plated on glass slides by centrifuging using a cytospin. The Hoechst 33258 staining was done using the Hoechst staining kit (Beyotime, China). Briefly, cells were fixed with fixing buffer for 20 min and stained with Hoechst 33258 staining buffer for 15 min. The cells were then observed under a confocal laser scanning microscope, Fluoview FV10i (Olympus) and analyzed using FV10-ASW4.0 software.
The cell cycle was analyzed using FlowCellect™ bivariate cell cycle kit (EMD Millipore). CTD-treated cells were harvested, fixed, and permeabilized according to the instructions of the kit. The permeabilized cells were stained with anti-p-Histone H3-AlexaFluor® 488 antibody and propidium iodide/RNase solution. Fluorescence was analyzed using FACScan laser flow cytometry (Guava easyCyte HT, Millipore).
Cells were plated on glass slides by centrifugation, fixed with 4% paraformaldehyde for 20 min and washed thrice with PBS. After permeabilizing with 0.3% Triton X-100 for 15 min, the cells were blocked with 5% bovine serum albumin and incubated with antibody against γH2AX (diluted 1:1000) overnight at 4°C, followed by incubation with Alexa Fluor 488 conjugated secondary antibody (diluted 1:1000). Nuclei were stained with DAPI. Fluorescence was observed with confocal laser scanning microscope Fluoview FV10i (Olympus) and analyzed using FV10-ASW4.0 software.
BCR-ABL specific siRNA and scramble RNA duplexes were transfected into K562 cells using lipofectamine 2000 (Invitrogen). The siRNA sequence for BCR-ABL was as follows: 5′-GCAGAGUUCAAAAGCCCTT-3′. The scramble siRNA (negative control, NC) sequence was 5′-UUCUCCGAACGUGUCAC GUTT-3′. The knockdown efficiency was assessed by western blotting. Cell viability of BCR-ABL knockdown cells after treating with CTD was measured using CCK-8 assay.
Total RNA was extracted from K562 and K562R cells using TRIzol reagent (Invitrogen). Five micrograms of total RNA was reverse transcribed using RNA PCR first-strand SuperScript Kit (Invitrogen). Fifty nanograms of total cDNA was used for qRT-PCR with the SYBR Premix Ex Taq Kit (TaKaRa). Each PCR reaction was carried out in a total volume of 20 μl on a 96-well optical reaction plate in Applied Bioscience 7500. The specific primers for qRT-PCR were BCR-ABL-sense: 5′-CATTCCGCT GACCATCAATAAG-3′, BCR-ABL-antisense: 5′-GATGCTACT GGCCGCTGAAG-3′, 18S-sense: 5′-AAACGGCTACCACATC CAAG-3′, and 18S-antisense: 5′-CCTCCAATGGATCCTCGT TA-3′. The relative gene expression was normalized against 18S RNA expression.
All experiments were performed at least three times and data were presented as the mean ± SD. GraphPad Prism5.0 software (GraphPad Software Inc., USA) was used for statistical analyses. One-way analysis of variance (ANOVA) followed by
In this experiment, the imatinib-resistant CML cell line K562R was used. We first characterized the resistance of K562R. Both K562 and K562R cells were treated with BCR-ABL kinase inhibitor, imatinib, at a concentration of 1 μM for 48 h. Apoptosis assay showed that K562R cells exhibited strong resistance against imatinib-induced apoptosis compared with K562 cells (
To investigate the anticancer potential of CTD against CML, the cytotoxicity of CTD toward normal PBMCs, imatinib-sensitive CML cell line, K562, and imatinib-resistant cell line, K562R, was tested using CCK-8 assay. The results demonstrated that CTD suppressed the viability of both CML cell types (Figs. 1A and 1B) with little effect on normal blood cells (Fig. 1C). The IC50 value of CTD for PBMCs (>100 μM) was significantly higher than that for K562 and K562R cells (28.23 and 54.42 μM, respectively) at 24 h. The IC50 values for PBMCs, K562, and K562R cells at 48 h were 102.69, 27.63 and 31.34 μM, respectively. Trypan blue exclusion assay showed that treatment of CTD induced cell death in K562 and K562R cells at the concentration of 5 to 80 μM (Figs. 1D and 1E).
Morphologic changes of the cells were examined under phase contrast microscope. The normal spherical shape of K562 and K562R cells changed into unusual ellipsoid or spindle shape, with significant enlargement, after exposure to CTD (5–20 μM) for 24 h (Fig. 2A). This result suggests that CTD treatment may result in a failure of cytokinesis in CML cells. The cell cycle can be divided into two distinct stages: the interphase stage and mitotic stage. In the second stage, or M-phase, chromatin condenses and cell division takes place. Previous studies have shown that Histone H3 phosphorylated (pH3) at Ser10 could be a reliable and specific mitotic marker (Crosio et al., 2002). To examine whether CTD could trigger mitotic arrest in CML cells, we analyzed CTD-treated cells by flow cytometry after anti-p-Histone H3/propidium iodide double staining. The results showed that CTD-treatment induced a significant increase in mitotic phase inK562 and K562R cells (Fig. 2B). As shown in Fig. 2C, after 24 h of CTD treatment, 19.2 to 24.5% of K562 cells were in mitotic phase, compared to only 1.6% of the control cells in mitotic phase; and 10.8 to 13.0% of K562R cells were in mitotic phase, compared to 3.11% of the control cells in mitotic phase. These results indicate that CTD induced mitotic failure in CML cells.
To further verify that CTD induced mitotic perturbation, we studied the changes in nuclear morphology after exposure to CTD. The cells underwent pronounced changes in nuclear morphology, including chromatin condensation (Fig. 3A). K562 cells with the abnormal mitotic nuclei accounted for about 1.05%, 17%, 24% and 36% after treatment with CTD at the concentration of 0 μM, 5 μM, 10 μM, and 20 μM, respectively (Fig. 3B).
We next investigated the mechanism of CTD triggered mitotic arrest. Activation of cyclin B1/Cdc2 complex, a heterodimer called mitosis-promoting factor, is essential for the transition of G2 to M phase. Upregulation of cyclin B1 is a typical marker of mitotic abnormality (Sánchez and Dynlacht, 2005; Wolanin et al., 2006). Therefore, we examined the expression levels of cyclin B1 by immunoblotting. As shown in Figs. 3C and 3D, CTD treatment caused a marked increase in the expression of cyclin B1 in K562 and K562R cells, respectively. Since dephosphorylation at Thr14 and Tyr15 of Cdc2 is crucial for its activation (Norbury et al., 1991), we next tested the phosphorylation status of Tyr 15 of Cdc2 in CTD-treated CML cells. The results showed that CTD reduced Tyr15-phosphorylated Cdc2 with no effect on the total protein level. Phosphatase, Cdc25c, is an upstream activator of Cdc2 through dephosphorylation at both Thr14 and Tyr15 sites (Gautier et al., 1991). We, therefore, examined the expression of Cdc25c and found a band shift of Cdc25c in a dose dependent manner. We also assessed the expression of cyclin D1, which drives the G1 to S phase transition and gets degraded in G2/M phase. The results showed the CTD-induced cyclin D1 reduction in both K562 (Fig. 3E) and K562R (Fig. 3F) cells. Taken together, these results demonstrated that CTD caused changes in mitotic signaling pathway.
As DNA damage was shown to be linked with cell cycle arrest, experiments to detect the occurrence of DNA damage were conducted. As shown in Fig. 4A, an increase in γH2AX foci was observed in K562 cells treated with 20 μM CTD for 24 h. Subsequently, we assessed the expression levels of γH2AX by immunoblotting. As shown in Fig. 4B, CTD triggered an increase in γH2AX in a dose-dependent manner in both K562 and K562R cells. K562 and K562R cells were treated with 10 μM CTD for 0–24 h, the expression of γH2AX increased in a time-dependent manner (Fig. 4C).
Previous studies have demonstrated that several compounds, such as curcumin (Blakemore et al., 2013) and diallyl trisulfide (Xiao et al., 2009), induced mitotic arrest involving DNA damage, and that inhibiting DNA damage signaling pathway abrogated the mitotic arrest. Therefore, we were interested in studying whether DNA damage signaling pathway was associated with CTD-induced mitotic arrest and cell death. K562 or K562R cells were treated with CTD in the presence or absence of caffeine (a pan-inhibitor of DNA damage network signaling kinases, ATM, ATR, DNA-PK and Chk1), and cell cycle status was evaluated. As shown in Figs. 5A and 5B, caffeine reduced CTD-induced mitotic arrest, although the reduction was not statistically significant. One explanation may be that we used caffeine at the concentration of 2.5 mM, which is much lower than the 10 mM used in previous studies (Blakemore et al., 2013). In our experiment, 10 mM caffeine caused significant cell death in K562 cells, and hence, we reduced the concentration to 2.5 mM. To further verify the role of DNA damage signaling kinases in CTD-mediated mitotic arrest, K562 or K562R cells were treated with CTD in the presence or absence of ATM/ATR inhibitor, CGK733, and the cell cycle arrest, cell viability and the levels of cleaved PARP, Mcl-1, and pH3 proteins were evaluated. We found that CGK733 significantly abrogated the CTD-mediated mitotic arrest (Figs. 5A and 5B), but enhanced cell death (Figs. 5C and 5D) as evidenced by increased level of cleaved PARP and decreased level of Mcl-1 (Fig. 5E). These results confirmed that inhibition of CTD-induced mitotic arrest leads to greater cell death in K562 and K562R cells.
The presence of oncoprotein BCR-ABL is the main characteristic of CML (Deininger et al., 2000; Pane et al., 1996). Increased expression of BCR-ABL and mutations in the tyrosine kinase domain of BCR-ABL are the major mechanisms underlying imatinib resistance (Bixby and Talpaz, 2010). Therefore, we examined whether CTD has any impact on BCR-ABL. CTD treatment significantly decreased BCR-ABL in a dose-dependent manner (Fig. 6A). CTD also inhibited the expression of the downstream target proteins of BCR-ABL. The phosphorylation of STAT5, AKT, and ERK1/2 was significantly decreased, whereas there was no dramatic change in the amount of total proteins. The downstream events of decreased BCR-ABL also included PARP cleavage. To study the mechanism of CTD-induced reduction of BCR-ABL protein, we measured the expression of BCR-ABL at the transcription level. K562 and K562R cells were treated with CTD or vehicle for 16 h, and then RNA was extracted. qRT-PCR data revealed a clear decrease in BCR-ABL mRNA levels in both cell lines (Fig. 6B), suggesting that CTD-induced suppression of BCR-ABL transcription may be responsible for the decreased BCR-ABL protein levels.
To further investigate whether BCR-ABL is the cause of cell-killing effect of CTD, we knocked down BCR-ABL in K562 cells. We first examined the CTD-mediated reduction of BCR-ABL protein in the cells transfected with scramble siRNA and siBCR-ABL. The results showed that the degree of BCR-ABL protein reduction in siBCR-ABL-transfected cells is more notable than it is in scramble siRNA-transfected cells (Fig. 6C). In addition, CTD triggered dramatic increase in cleaved PARP level and decrease in the level of Mcl-1 in BCR-ABL knockdown cells. Subsequently, we compared the cell-killing effect of CTD on scramble siRNA and siBCR-ABL-transfected cells. Since the cell viability was altered by the knockdown of BCR-ABL, we designated the growth rate of CTD-untreated, siRNA negative control cells as 100% for comparing the cell-killing effect of CTD on the the siRNA negative control and siBCR-ABL K562 groups. The results indicated that BCR-ABL knockdown cells were more sensitive to the cytotoxic effects of CTD compared to negative control cells (Fig. 6D). These findings suggested that the cell-killing effects of CTD are at least in part dependent on BCR-ABL protein downregulation.
CTD is an active constituent of blister beetles of the genus
An earlier study showed that CTD disrupted the metaphase alignment of chromosomes and induces a prolonged mitotic arrest in A549 cells (Bonness et al., 2006). Remarkably, the dose of CTD that induced mitotic arrest did not result in apoptosis in A549 cells, because these cells can progress from metaphase into anaphase after a 10 to 15-hour delay. In this study, we showed that CTD causes accumulation of K562 and K562R cells in mitotic phase, as indicated by the morphologic changes and increased expression of pH3 (S10) (Fig. 2). Hoechst 33258 staining indicated chromosomal condensation (Fig. 3A). The ATM/ATR inhibitor, CGK733, abrogated CTD-mediated mitotic arrest but promoted cell death in CML cells (Fig. 5). Although we are not sure whether CTD-treated CML cells can later enter anaphase, CTD-induced mitotic arrest may not result in cell death.
We found that CTD-induced γH2AX accumulation and DNA damage signaling pathway regulate CTD-induced mitotic arrest in CML cells. As shown in Figs. 5A and 5B, caffeine and CGK733 can reduce the degree of CTD-induced mitotic arrest. Indeed, DNA damage signaling pathway is closely associated with mitotic arrest (Blakemore et al., 2013; Xiao et al., 2009). In Caco-2 and HCT116 cells, curcumin disrupted the chromosomal alignment in mitotic cells and arrested the cells at the prophase/prometaphase stage of mitosis. Pretreatment with caffeine abrogated curcumin-induced mitotic arrest. The authors proposed that DNA damage might contribute to mitotic arrest caused by curcumin. The compound, diallyl trisulfide, also triggers Chk1-mediated mitotic arrest and drives apoptotic DNA fragmentation in LNCaP and HCT116 cells. These reports, together with our work, suggest that DNA damage signaling pathway is closely associated with mitotic arrest.
Imatinib resistance is becoming a great challenge in the treatment of CML patients. Although new generations of TKIs have been developed to overcome the resistance to imatinib, most of them are extremely expensive and their effects need further clinical investigations (Apperley, 2015). A novel strategy to overcome imatinib resistance could possibly be downregulating the addiction oncogene, BCR-ABL, at mRNA level or promoting the degradation of BCR-ABL protein (Chen et al., 2014; Lu et al., 2010; Shi et al., 2014). In this study, we reported that CTD decreased BCR-ABL expression and contributed to cell death in CML cells (Fig. 6). BCR-ABL is a constitutively active tyrosine kinase that phosphorylates several substrates and activates multiple signal transduction pathways. CTD treatment results in dose-dependent downregulation of BCR-ABL and its downstream signaling pathways (Fig. 6A). Further study revealed that CTD depleted BCR-ABL at transcriptional level (Fig. 6B). We also showed that BCR-ABL knockdown makes the cells sensitive to CTD (Figs. 6C and 6D). Even though CTD might be affecting multiple molecules, depleting BCR-ABL is at least one of the major factors that induce cell death in CML cells.
Previous reports have demonstrated that the phosphatase activity of PP2A is suppressed by the BCR-ABL enhanced expression of PP2A inhibitor SET, and reactivating PP2A can kill both TKI-sensitive and TKI-resistant CML cells (Neviani, 2005). It is known that CTD is a selective PP2A inhibitor, which seems contradictory to previous reports. We reasoned that CTD-induced downregulation of BCR-ABL mRNA is independent of its inhibitory effects on PP2A.
Recently, it has been demonstrated that CTD (0.2 and 1.0 mg/kg) treatment led to a reduction of tumor size in A431 xenograft mouse model (Li et al., 2016). Han et al. (2013) also reported that CTD treatment (0.25, 0.5, and 1.0 mg/kg) caused significant regression of S180 cell xenograft tumors in ICR male mice. In this study, treatment of CTD induced cell death in K562 and K562R cells at the concentration of 5–80 μM (Figs. 1D and 1E). In addition, the CTD-mediated BCR-ABL depletion effect was observed at 10 and 20 μM concentrations after 24 h treatment (Fig. 6A), suggesting that CTD has an anticancer effect in CML xenograft murine models. Therefore, more investigations on the anticancer effects of CTD on CML murine models are needed in future.