Killing tumor cells without causing too much harm to healthy cells is a key issue in chemotherapy (Reedijk, 2003). Cisplatin (cis-diamminedichloroplatinum II) is one of the most effective chemotherapeutic agents for cancer treatment (Loehrer and Einhorn, 1984). However, acquired resistance to cisplatin frequently cause treatment failure (Wozniak and Blasiak, 2002).
Although resistance to cisplatin has been widely studied, it is still difficult to unravel its molecular changes. Several mechanisms are thought to be involved in drug resistance, including decreased intracellular drug accumulation, increased drug detoxification, increased DNA repair activity, and reduced ability to undergo apoptosis; however, no particular chemoresistance system has been identified (Wernyj and Morin, 2004; Wang and Lippard, 2005; Piulats et al., 2009; Brozovic et al., 2010).
Cell apoptosis is usually accompanied with apoptotic volume decrease. Apoptotic volume decrease is induced by ionic effluxes resulting mainly from increased K+ and Cl− conductances via volume regulatory anion channel (Shimizu et al., 2004). The volume regulatory anion channel plays a cell-rescuing role in the necrotic insults. Normotonic activation of the anion channel plays a cell-killing role in the apoptotic process by triggering apoptotic volume decrease following stimulation with apoptosis inducers (Okada et al., 2004). Therefore, we hypothesize that the blocking of the Cl− channels might prevent cisplatin-induced cell apoptosis.
In this study, we found that NPPB, a chloride channel blocker, could induce resistance to cisplatin in human erythroleukemia K562 cells. To further determine whether cisplatin resistance is associated with ATP-binding cassette transporters that cause drug efflux from cancer cells, the human erythroleukemia cell line RK562, which expresses P-glycoprotein and multidrug resistance protein, is also studied here. We also found that NPPB treatment increased the viabilities of K562 and RK562 cells treated with cisplatin. The acridine orange assay showed that acidification of the intracellular compartment is correlated with cisplatin resistance.
MATERIALS AND METHODS
The human erythroleukemia K562 and RK562 cells (a gift from Tianjin Institute of Hematology, China) were cultured in Iscove's modified Dulbecco's medium (IMDM; Life Technologies, Gaithersburg, MD), supplemented with 10% fetal bovine serum (Gibco BRL), and with 100 U/mL penicillin and 100 U/mL streptomycin at 37°C, 5% CO2 with high humidity. In addition, RK562 cells were maintained in IMDM with 10% fetal bovine serum medium containing 1 μg/mL doxorubicin (Sigma) to maintain resistance. RK562 cells were incubated in doxorubicin-free medium for 2 weeks and then used for experiments. Both cell lines were divided into four groups: non-treated cells, cells treated with 10 μg/mL cisplatin (Sigma), cells treated with 50 μM NPPB (Sigma), and cells treated with 10 μg/mL cisplatin combined with 50 μM NPPB, respectively.
Cell viability was determined by MTT assay. The MTT assay is a colorimetric assay that relies on the ability of viable cells to convert a soluble tetrazolium salt, 3-(4, 5-dimethyl-2-tetrazolyl)-2, 5-diphenyl-2H tetrazolium bromide (MTT), into a formazan precipitate, causing a yellow-to-purple color change (Shimizu et al., 2004). Exponentially growing K562 cells and RK562 cells at a density of 2 × 104 cells in 100 μL were seeded into 96-well culture plates and incubated for 24 hr. Then, the indicated concentration of cisplatin and/or NPPB were added into four parallel wells of different groups' cells and inoculated for another 24 hr, additionally K562 cells were treated with 5 μg/mL doxorubicin and/or NPPB and 2 mM hydrogen peroxide and/or NPPB and 20 μM vitamin K3 and/or NPPB for 24 hr. Twenty microliters of MTT solution (5 mg/mL in PBS) was added into each well and incubated for 4 hr. Then, 150 μL of dimethyl sulfoxide (Beijing chemical industry limited company, China) was added into each well. After shaking for 10 min, the absorbance of each well was measured at 570 nm using a Microplate Reader (Bio-Tek Instruments). Cell viability was calculated as follows:
RNA Extraction and RT-PCR Analysis
Total RNA was extracted from cells using Trizol reagents (Invitrogen) according to the manufacturer's direction. RT-PCR was performed to analyze the mRNA levels of Bax, Bcl-2, and CIC-3. GAPDH was used as the internal standard. The cDNA was amplified using Takara RNA PCR kit. The PCR primers used for different factors were as follows: The GAPDH primers: (sense strand: 5′-GGGTGATGCTGGTGCTGAGTATGT-3′, antisense strand: 5′-AAGAATGGGAGTTGCTGTTGAAGTC-3′); The Bax primers: (sense strand: 5′-AGGGTTTCATCCAGGATCGAGC-3′, antisense strand: 5′-AGGCGGTGAGGACTCCAGCC-3′); The Bcl-2 primers: (sense strand: 5′-AACACCAGAATCAAGTGTTCG-3′, antisense strand: 5′-TCAGGTGGACCACAGGTGGC-3′); The ClC-3 primers: (sense strand: 5′-CCTCTTTCCAAAGTATAGCAC-3′, antisense strand: 5′-TTACTGGCATTCATGTCATTTC-3′). The cyclin D1 primer: (sense strand: 5′-TGGATGCTGGAGGTCTGCGAG-3′, antisense strand: 5′-GGCTTCGATCTGCTCCTGGC-3′). The PCR products were measured by 1% agarose gels. Each mRNA transcript was then quantified and normalized against that of GAPDH. Autoradiographic films of RT-PCR assays were subjected to densitometric analyses (Tanon GI-2010, China). The ratio of the density of each group was compared with the GAPDH group.
Proteins were extracted from K562 and RK562 cells using cell lysis buffer (50 mM Tris-HCl, pH 7.5, 150 mM NaCl, 1 mM Na2EDTA, 1 mM EDTA, 1% Triton, 2.5 mM sodium pyrophosphate, 1 mM β-glycerophosphate, 1 mM Na3VO4, 1 mM NaF, 1 μg/mL leupeptin, and 1 mM PMSF; Okada et al., 2004). Protein concentrations were measured using BIO-RAD kit (Pierce). Anti-Bax, anti-Bcl-2, anti-cytochrome C, anti-caspase-3, anti-P-gp, and anti-β-actin antibody (1:200 dilution), horseradish peroxidase-conjugated secondary antibody (1:2000 dilution) were obtained from Santa Cruz Biotechnology. The protein bands against anti-Bax, anti-Bcl-2, anti-cytochrome C, anti-caspase-3, anti-P-gp, and anti-β-actin were visualized by chemiluminescence reagents (Santa Cruz).
Caspase-3 Activity Assay
Caspase-3 activity was used as a quantitative method for measuring apoptosis. Briefly, cells were lysed and the supernatant was used for measurement of caspase-3 activity using the caspase-3 activity kit (Beyotime Institute of Biotechnology, Haimen, China) according to the instructions provided by the manufacturer. To evaluate the activity of caspase-3, cells were homogenized in 100 mL reaction buffer [1% NP-40, 20 mM Tris-HCl (pH 7.5), 137 mM Nad, and 10% glycerol] containing 10 mL substrate for caspase-3 (Ac-DEVD-pNA; 2 mM) after all treatments. Lysates were incubated at 37°C for 2 hr. Samples were measured with an ELISA reader (Eppendorf, Germany) at an absorbance of 405 nm.
Analysis of the Apoptosis Rate by Annexin-V and FITC/Propidium Iodide
The apoptosis rate was measured by flow cytometry (BD Biosystems) according to the instructions supplied with the Annexin V-FITC Apoptosis Detection Kit (KeyGEN, China). The percentage of apoptotic cells in a population of 10,000 cells was determined. All the experiments reported in this study were performed in triplicate.
Cell Organelle Acidification Assay
Cells in exponential growth phase were incubated at 5 × 104/mL density in IMDM containing AO (Sigma) (6 mM in medium from a 10 mM stock in water) at 37°C for 15 min. Cells were washed free of AO and seeded in Petri dishes containing IMDM without phenol red. Scanning of cells was performed under laser-scanning confocal microscopy (FV-500, Olympus, Japan). To characterize the profile of AO emission spectra, the red band contribution (R%) within the whole emission spectrum has been calculated as follows by Fluoview Version 4.2 (Olympus, Japan):
where I530 and I655 are the green (520–540-nm) and the red (645–665-nm) integrated emission intensities, respectively (Millot et al., 1997).
Data are representative of three independent experiments each in triplicate determination. Statistical analysis of the data was performed using one-way ANOVA. The Tukey post hoc test was used to determine the significance for all pairwise comparisons of interest. P Values of <0.05 were considered to represent a statistically significant difference.
On the basis of our dose-dependent assay and previous studies, 10 μg/mL cispaltin could inhibit RK562 cells obviously, so we treated both cell lines with 10 μg/mL cisplatin. Although MTT assays indicated that NPPB alone had no significant effect on cell viability, NPPB treatment decreased the cytotoxic effect of cisplatin (Fig. 1A). At the same time, under the phase contrast microscope, we found that NPPB treatment could protect K562 and RK562 cells treated with cisplatin (Fig. 1B). NPPB treatment did not affect cytotoxicity of doxorubicin, hydrogen peroxide or vitamin K3 in K562 cells (Fig. 1C).
The Different Expressions of MDR1, MRP1 mRNA, and P-Glycoprotein
Using RT-PCR, we found that RK562 cells expressed MDR1 and MRP1 mRNA but K562 cells did not (Fig. 2A). By Western Blot analysis, we demonstrated that RK562 cells expressed the 170KD multi-drug resistance protein P-glycoprotein but K562 did not (Fig. 2B).
The Expressions of Bax/Bcl-2, Cyclin D1, and ClC-3 mRNA
Bax/Bcl-2 was upregulated in both cell lines. The expressions of Cyclin D1 and ClC-3 were decreased in both cells following the treatment of cisplatin. Compared with cells treated only with cispaltin, Bax/Bcl-2 mRNA decreased, Cyclin D1 and ClC-3 mRNA increased in both cell lines treated with cispaltin combined with NPPB (Fig. 3A,B).
The Effect of NPPB Treatment on the Expressions of Bax/Bcl-2, Cytochrome C, and Caspase-3
Using Western Blot analysis, we found that compared with cells treated with cisplatin alone, the expressions of Bax/Bcl-2, cytochrome C and caspase-3 significantly decreased in both cell lines treated with cisplatin combined with NPPB (Fig. 4A,B).
Effects of NPPB on Caspase-3 Activity in Cisplatin-Treated K562 and RK562 Cells
After treated with cisplatin or cisplatin combined with NPPB for 24 hr, cells were harvested, and caspase-3 activity was measured. Compared with control group, a rise in caspase-3 activity was found in cisplatin-treated cells, while cells treated cisplatin combined with NPPB showed a significant decrease in the caspase-3 activity (Fig. 4C).
Apoptosis Analysis of K562 and RK562 Cells with NPPB Treatment
After treated with cisplatin or cisplatin combined with NPPB for 24 hr, cells were then double stained with annexin-V and FITC/PI and analyzed by flow cytometry. Compared with cells treated with cisplatin alone, the apoptotic rates decreased in cells treated with cisplatin combined with NPPB (Fig. 5A,B). These results indicate that NPPB protect K562 and RK562 cells from cisplatin-induced apoptosis.
Effects of NPPB on Cellular Acidification
Our previous results indicated that NPPB could regulate the expression of ClC-3. In addition, ClC-3 was reported to be associated with intracellular compartment acidification (Weylandt et al., 2007). We used acridine orange (AO) to analyze the effect of NPPB on the organelle acidification of K562 and RK562 cells (Millot et al., 1997). It is a weak basic fluorescent probe that emits green at low concentrations and red at high concentration. AO accumulates in acidic compartments, producing fluorescence (Fig. 6A,B). Compared with cells treated with cisplatin alone, the rate of acidification of intracellular compartment increased in cells treated with cisplatin combined with NPPB (Fig. 6C). These results indicate that NPPB could increase the organelle acidification in K562 and RK562 cells.
Chemotherapeutic drug resistance is a major problem in cancer chemotherapy (Chang, 2010). Cis-diamminedichloroplatinum II (cisplatin), as a single agent or in combination with other drugs, is effective in the treatment of a variety of tumors (Muggia, 2009). It functions to crosslink with DNA and to form intra- and inter-strand adducts (Chaney et al., 2005; Basu and Krishnamurthy, 2010). Like most chemotherapeutic agents, the anticancer activity of cisplatin is known to be attained by initiating apoptosis (Rebillard et al., 2008; Konstantakou et al., 2009; Suzuki et al., 2009; Huang et al., 2010). However, the exact mechanisms are not clear.
Bcl-2 family members are the key regulators of cispaltin-induced apoptosis. The antiapoptotic protein Bcl-2 helps to inhibit apoptosis by preventing the release of mitochondrial apoptogenic factors, presumably via interaction with the mitochondrial porin channel (Shimizu et al., 1999; Shimizu et al., 2000). Bax interacts with Bcl-2 and inhibits the antiapoptotic role of Bcl-2. Upregulating expression of Bax/Bcl-2 indicates the mitochondrial apoptotic pathway activation. The expression of Bax/Bcl-2 regulates the expressions of cytochrome C and caspase-3, which are mitochondrial apoptogenic factors inducing apoptosis, and also affects cyclin D1 which is a cell cycle regulating protein controlling cell proliferation (Cho et al., 2006). In this study, we found that cisplatin treatment upregulated the expressions of Bax/Bcl-2, cytochrome C, and caspase-3, and downregulated the expression of cyclin D1, and showed a rise in caspase-3 activity. On the contrary, compared with cells treated only with cisplatin, NPPB treatment downregulated the expressions of Bax/Bcl-2, cytochrome C, and caspase-3, and upregulated the expression of cyclin D1 in cells treated with cisplatin combined with NPPB, and showed a significant decrease in the caspase-3 activity, which decreased cisplatin-induced apoptosis.
Clinically, significant levels of resistance to cisplatin represent a major obstacle to effective cancer therapy (Olszewski and Hamilton, 2010). Acquired cisplatin resistance is associated with defects in the apoptotic program, decreased cisplatin accumulation, and increased drug inactivation (Basu and Krishnamurthy, 2010). ATP-binding cassette transporters that enhance drug efflux from cancer cells could lead to an increase in drug resistance (Panczyk et al., 2007). One of the ATP-binding cassette transporters P-glycoprotein, the product of MDR1 gene, is an important drug efflux protein, and acts as a drug pump in multidrug-resistant cancer cells (Borst and Schinkel, 1997; Allen et al., 2000). It is generally believed that reduced cisplatin accumulation in cisplatin resistant cells is due to decrease in uptake of cisplatin rather than an increase in drug efflux (Basu and Krishnamurthy, 2010).
In our studies, we clearly found that RK562 cell lines, which express ATP-binding MDR1 and MRP1, were partially resistant to apoptosis induced by cisplatin. These results suggested that ATP-binding cassette transporters did not play a key role in resistance to cisplatin.
Membrane ion channels are known to be of importance in the development of cancer (Kunzelmann, 2005). The apoptotic volume decrease, which precedes caspase activation, DNA laddering and cell fragmentation into apoptotic bodies, is known to be a prerequisite to apoptosis and to be induced by activation of volume-regulatory K+ and Cl− channels (Lang et al., 2005, 2006). The apoptotic volume decrease inducing Cl− channel has been functionally identified as the volume regulatory anion channel for both mitochondrion-mediated and death receptor-mediated apoptosis (Shimizu et al., 2004). In our study, we detected the role of NPPB, a chloride channel blocker in K562 and RK562 cell lines with cisplatin treatment. We found that NPPB could protect both cell lines from apoptosis induced by cisplatin, but did not affect cytotoxicity of vitamin K3, hydrogen peroxide or doxorubicin. The results showed that Cl− channel is involved in cisplatin resistance. Some studies reported that NPPB can rival the apoptotic capacity of some anticancer drugs to a certain extent (Okada et al., 2001; Tafani et al., 2002; Ise et al., 2005; Heimlich and Cidlowski, 2006), which is consistent with our results. In contrast, one report suggested that 100 μmol/L NPPB can inhibit cell proliferation (Wondergem et al., 2001). The difference between these results may be due to the different conditions and different cell lines.
Several studies have implied a relationship between endosomal and lysosomal acidity and chemotherapy resistance, hypothesizing that weakly basic drug, such as cisplatin can be sequestered to acidic intracellular membrane compartments (De Milito and Fais, 2005; Safaei et al., 2005). Weylandt et al. (2007) found that as a chloride channel/transporter of the CLC protein family, ClC-3 expression enhances etoposide resistance by increasing acidification of the late endocytic compartment. The results presented here also showed that while the resistance to cisplatin in both cell lines increased, the ClC-3 mRNA was increased. The acidification of the late endocytic compartment was increased as well. This is in accordance with the results of Weylandt et al. (2007).
In summary, we found that the chloride channel blocker NPPB upregulated ClC-3, which made K562 and RK562 cells effectively avoid mitochondrion-mediated apoptosis. This evidence indicates the involvement of Cl− channels in drug resistance by allowing cells to avoid cisplatin-induced apoptosis. The Cl− channels could serve as therapeutic targets to improve the efficacy of cisplatin. However, the exact cellular and molecular mechanism by which NPPB protects cells from cisplatin-induced apoptosis still needs to be clarified.
The authors appreciate the assistance of Professor William Orr from the University of Manitoba Canada for his help in revising the manuscript.