We have previously shown that cisplatin triggers an early acid sphingomyelinase (aSMase)-dependent ceramide generation concomitantly with an increase in membrane fluidity and induces apoptosis in HT29 cells. The present study further explores the role and origin of membrane fluidification in cisplatin-induced apoptosis. The rapid increase in membrane fluidity following cisplatin treatment was inhibited by membrane-stabilizing agents such as cholesterol or monosialoganglioside-1. In HT29 cells, these compounds prevented the early aggregation of Fas death receptor and of membrane lipid rafts on cell surface and significantly inhibited cisplatin-induced apoptosis without altering drug intracellular uptake or cisplatin DNA adducts formation. Early after cisplatin treatment, Na+/H+ membrane exchanger-1 (NHE1) was inhibited leading to intracellular acidification, aSMase was activated, and ceramide was detected at the cell membrane. Treatment of HT29 cells with Staphylococcus aureus sphingomyelinase increased membrane fluidity. Moreover, pretreatment with cariporide, a specific inhibitor of NHE1, inhibited cisplatin-induced intracellular acidification, aSMase activation, ceramide membrane generation, membrane fluidification, and apoptosis. Finally, NHE1-expressing PS120 cells were more sensitive to cisplatin than NHE1-deficient PS120 cells. Altogether, these findings suggest that the apoptotic pathway triggered by cisplatin involves a very early NHE1-dependent intracellular acidification leading to aSMase activation and increase in membrane fluidity. These events are independent of cisplatin-induced DNA adducts formation. The membrane exchanger NHE1 may be another potential target of cisplatin, increasing cell sensitivity to this compound. [Cancer Res 2007;67(16):7865–74]

Cis-diaminedichloroplatinum II (CDDP or cisplatin) is an effective antineoplastic agent commonly used in the treatment of solid tumors. Although DNA was considered as the primary target of cisplatin (1, 2), the cisplatin action at the cellular level still remains unknown. In fact, cisplatin-induced DNA adducts activate several signal transduction pathways culminating in the activation of apoptosis (3). However, these specific biochemical lesions are not sufficient to explain cisplatin cytotoxicity (4, 5), and the potential contribution of other cellular targets requires further investigation (6). Particularly, cisplatin has been shown to interact with the plasma membrane and reduce the activity of certain ion channels, transport proteins (7), and various plasma membrane enzymes (8). Elucidating the mechanisms by which cisplatin kills cancer cells may lead to develop new therapeutic strategy.

The plasma membrane constitutes the first cellular barrier that encounter antineoplastic agents, and the importance of the lipid bilayer as a medium in which diffusion of drugs takes place has been far well established (9). Many anticancer drugs show membrane effects via weak hydrophobic interaction or via electrostatic binding to membrane phospholipids before entering the cytoplasm. In this way, it has been shown that cisplatin can interact with phosphatidylserine to alter the characteristics of the membrane (10).

Recent findings indicate that variations of membrane fluidity occurs in cells exposed to various cytotoxic stimuli including cisplatin (11, 12). Increase in membrane fluidity could play a role in apoptosis because membrane-stabilizing agents reduced this increase as well as cell death induction in several cell models (1214). Moreover, recent studies showed the implication of membrane lipid rafts, which result from specific interactions between various types of lipids and proteins (15, 16), in the Fas death receptor pathway (17), and in the cytotoxicity of some anticancer agents (1820).

Several apoptotic stimuli require the acid sphingomyelinase (aSMase) to trigger apoptosis and particularly some chemotherapeutics such as doxorubicin and cisplatin (for a review, see ref. 21). Two forms of aSMase coming from the same gene (aSMase gene) have been described, an endolysosomal form and a secretory form which is targeted to the plasma membrane (22, 23) and implicated in stress-associated cell death. Intracellular acidification has been detected following exposure of cells to various apoptotic stimuli (for a review, see ref. 24). Among the nine NHE family members, the Na+/H+ membrane exchanger-1 (NHE1) is ubiquitously expressed and plays a critical role in intracellular pH and cell volume homeostasis (25).

We have previously shown that cisplatin induces Fas receptor clustering and activation independently of Fas ligand (FasL; ref. 26), and the generation of ceramide and the relocalization of Fas receptor molecules into lipid rafts of HT29 cells with a transient increase in membrane fluidity (20). The present study further explores the role of increased membrane fluidity in cisplatin-induced apoptosis and identifies for the first time the molecular mechanisms involved in this process. Cisplatin induces a very early NHE1-dependent intracellular acidification that favors aSMase activation, thereby increasing membrane fluidity and aggregation of membrane lipid rafts.

Chemicals and Antibodies

CDDP was from Merck, water-soluble cholesterol (CHOL) and exogenous sphingomyelinase (from Staphylococcus aureus) were from Sigma-Aldrich, monosialoganglioside-1 (GM1) was purchased from Alexis Biochemicals, and cariporide was a kind gift from Aventis (Frankfurt, Germany). SR33557 (SR) was kindly provided by Drs. C. Bezombes and J.P. Jaffrézou (Inserm U563, Toulouse, France).

Hoechst 33342 and fluorescein-tagged cholera toxin subunit B (CTx-FITC) were from Molecular Probes. TransFectine lipid and Bradford reagents were from Bio-Rad. Mouse monoclonal immunoglobulin G1 (IgG1) anti-Fas (ZB4) was from Beckman Coulter. Mouse monoclonal IgM anti-CER (15B4) was from Alexis Biochemicals.

Cell Culture and Treatments

HT29, HCT116, and SW480 human colon carcinoma cell lines were obtained from the American Tissue Culture Collection (Biovalley) and cultured in EMEM (Eurobio) complemented with 10% FCS (Life Technologies BRL) and 2 mmol/L l-glutamine (Life Technologies BRL). For all experiments, cells were seeded onto tissue culture Petri dishes and allowed to attach for 24 h before treatment. Cells, growing in exponential phase, were treated with 25 μmol/L CDDP for different times. When indicated, cells were pretreated for 2 h with various compounds.

The NHE1-deficient PS120 cell line (PS120) is a mutant Chinese hamster fibroblast cell line that lacks NHE1 activity. This cell line was grown in DMEM containing 15 mmol/L NaHCO3 supplemented with 7.5% FCS at 37°C in a humidified atmosphere of 5% CO2–95% air. The NHE1-PS120 cells are PS120 cells stably transfected with NHE1 (ref. 27; kindly provided by Prof. Laurent Counillon, CNRS UMR 6548, Nice, France).

Cell Death Assays

See details in Supplementary Data (28).

Detection of Membrane Lipid Rafts by GM1 Staining with CTx-FITC

HT29 cells were treated with 25 μmol/L cisplatin for various times. After treatment, cells were fixed in 4% paraformaldehyde (Sigma-Aldrich) for 15 min, washed in PBS 1×, and then preincubated with PBS-bovine serum albumin (PBS-BSA) 2% for 30 min before staining with CTx-FITC for 30 min. The fluorescent images of GM1 staining were analyzed using a DMRXA Leica microscope and a COHU high-performance CCD camera using a metavue software.

Determination of Membrane Fluidity by Electron Paramagnetic Resonance Spin-Labeling Method

Bulk membranes. The membrane fluidity of cells was determined by a spin-labeling method using electron paramagnetic resonance (EPR). After treatment, cells collected in PBS were incubated with 50 μg/mL 12-DSA spin label for 15 min at 37°C and then were washed thrice with cold PBS to eliminate the free spin label. The final pellet was kept on ice to prevent any spin label reduction before analyzing the EPR spectrum at room temperature (20°C) using a Bruker ECS 106 spectrometer (9.82 GHz frequency, 20 mW microwave power, 1.771 G modulation amplitude, and 100 kHz modulation frequency; Bruker Spectrospin). The values of inner hyperfine-splitting EPR spectra, typical for 12-DSA spin label, were used to calculate the membrane order parameter S. A decrease in the membrane order parameter reflects an increase in membrane fluidity.

Lipid rafts. Lipid rafts were isolated by ultracentrifugation on sucrose gradient as previously described (20). Briefly, after ultracentrifugation, 12 fractions of 1 mL were collected from the top of the gradient. The first six fractions corresponding to membrane lipid rafts were pooled and centrifuged at 39,000 rpm for 20 h at 4°C. The pellet was resuspended in PBS, and the spin label was incorporated for 30 min at 4°C. After three washes in cold PBS, the analysis of the EPR spectrum was done as described above for bulk membranes.

Immunofluorescence Microscopy and Fluorescence-Activated Cell Sorting Analysis

After treatment, cells were fixed in 4% paraformaldehyde (Sigma-Aldrich) for 15 min, washed in PBS 1×, and then preincubated with PBS-BSA 2% for 30 min before incubation for 2 h with a mouse monoclonal IgG1 anti-Fas (ZB4, 1:150, Immunotech) or with a mouse monoclonal IgM anti-CER (15B4, 1:50, Alexis Biochemicals) or with isotype-matched controls. Cells were then washed in PBS and stained for 45 min with FITC-labeled goat anti-mouse IgG (Molecular Probes) or FITC-labeled goat anti-mouse IgM (Jackson Immunoresearch Laboratories). The fluorescent images of ceramide and Fas stainings were analyzed using a DMRXA Leica microscope and a COHU high-performance CCD camera using a metavue software. Cell fluorescence (FL-1) corresponding to ceramide staining was analyzed by flow cytometry (FACScalibur, Becton Dickinson).

Measurement of Intracellular Platinum Content

Cells were treated with 25 μmol/L cisplatin for various times. Cells were then washed twice with ice-cold PBS, scraped in ice-cold water, and sonicated for 10 s. Proteins were measured by the Bradford protein assay. Platinum content was determined by flameless atomic absorption spectroscopy using a Zeeman apparatus with a graphite furnace (Spectra A300, Varian) and normalized to cellular proteins.

Southwestern Analysis of 1,2 Pt-[GG] Adducts

Cells were treated with 25 μmol/L cisplatin for various times. Genomic DNA was isolated by the use of the Nucleospin Tissue kit (Macherey-Nagel). DNA (2 μg) was diluted in 100 μL TE buffer (Tris-EDTA), incubated 10 min at 94°C, and thereafter placed on ice. After the addition of 100 μL ice-cold 2 mol/L CH3CO2NH4, the sample was transferred to a positively charged nylon membrane (Hybond plus, Amersham) by vacuum slot blotting. Thereafter, the membrane was washed with 1 mol/L CH3CO2NH4, neutralized with 5× SSC (saline sodium citrate), and fixed by baking the membrane for 2 h at 80°C. Monoclonal antibodies specific for 1,2 Pt-[GG] adducts (kindly provided by Prof. J. Thomale, Institute of Cell Biology, University of Essen Medical School, Essen, Germany) were used at a dilution of 1:200. The additional detection was done using monoclonal anti-rat antibodies at a dilution of 1:2,000 and the enhanced chemiluminescence detection reagent from Amersham.

Acid Sphingomyelinase Assay

After treatment, HT29 cells were lysed in 0.1% (w/v) Triton X-100 and sonicated for 10 s. Proteins were measured by the Bradford protein assay. Reaction was started by adding to the lysate (100 μL) 100 μL of substrate solution [choline-methyl-14C]SM [54.5 mCi/mol, 1 × 105 dpm per assay (∼1 nmol per assay), Perkin-Elmer Life Sciences] in 0.1% (w/v) Triton X-100, 250 mmol/L sodium acetate buffer (pH 5), and 1 mmol/L EDTA. After a 1-h incubation at 37°C, the reaction was stopped by adding 300 μL of H2O and 2.5 mL of chloroform/methanol (2:1; v/v). Phases were separated by centrifugation (3,000 rpm for 10 min), and the amount of released radioactive phosphocholine was determined by subjecting 750 μL of the upper phase to scintillation counting (Beckman LS6500, Beckman Coulter). Each value (expressed in disintegration/min) was corrected for protein content, and the activity of aSMase was expressed as the % of value determined in nontreated cells.

Measurement of Intracellular pH

The intracellular pH (pHi) of HT29 cells cultured on glass coverslips was monitored using the pH-sensitive fluorescent probe carboxy-SNARF-1 (carboxy-seminaphtorhodafluor, Molecular Probes) as previously described (29). SNARF-loaded cells were placed in a continuously perfused recording chamber (at a temperature of 36 ± 1°C) mounted on the stage of an epifluorescent microscope (Nikon Diaphot). HT29 cells were then excited with light at 514 nm, and fluorescence from the trapped probe was measured at 590 and 640 nm. The emission ratio 640:590 nm (corrected for background fluorescence) detected from intracellular SNARF was calculated and converted to a linear pH scale using in situ calibration. Addition and subsequent removal of NH4+ were used to induce an acid load to activate NHE1, which is the only pHi-regulating mechanism active under our conditions.

RNA Interference

See details in Supplementary Data.

RNA Isolation and Reverse Transcription-PCR Analysis

See details in Supplementary Data.

Statistical Analysis

Statistical analyses were carried out using the unilateral Student's t test considering the variances as unequal. The significance is shown as follows: *, P ≤ 0.05; **, P ≤ 0.01; and ***, P ≤ 0.001.

Cisplatin treatment increases fluidity in bulk membranes as well as in membrane lipid rafts. By using EPR spectroscopy and 12-DSA spin-labeling technique, which allows to calculate the membrane order parameter, inversely related to membrane fluidity, an increase in membrane fluidity was measured in HT29 cells between 15 min and 4 h after cisplatin treatment (20). Figure 1A confirmed that a 1-h treatment with 25 μmol/L CDDP induced an increase in membrane fluidity in HT29 bulk membranes (i.e., plasma membrane and possibly endosome and lysosome membranes). In addition, by using the same methodologic approach to study the modifications of fluidity in isolated lipid rafts, we showed a similar decrease in the membrane order parameter of lipid rafts purified from cisplatin-treated HT29 cells (Fig. 1A , top); this thus suggested that the fluidity of both raft and nonraft regions of the membrane was increased following cisplatin treatment. Note that the membrane order parameter calculated for lipid rafts is still greater than the one calculated for bulk membranes in both untreated and cisplatin-treated cells, showing a more ordered lipid environment in lipid rafts than in bulk membranes. In the next sets of experiments, we measured the fluidity in bulk membranes.

Figure 1.

Cisplatin-induced apoptosis is dependent on increased membrane fluidity. A, cisplatin treatment increases fluidity in bulk membranes as well as in membrane lipid rafts. HT29 cells were treated or not (NT) with 25 μmol/L cisplatin (CDDP) for 1 h. Membrane lipid rafts were purified by ultracentrifugation on a discontinuous sucrose gradient. Membrane fluidity was determined on bulk membranes or isolated lipid rafts by EPR using a spin-labeling method. The EPR spectra were used to calculate the membrane order parameter S, which is conversely proportional to membrane fluidity. Values are the mean ± SE of three independent experiments. **, P ≤ 0.01, CDDP versus NT (top). Pretreatment with membrane stabilizers inhibits cisplatin-induced increase in membrane fluidity. HT29 cells were pretreated or not (CONTROL) with membrane stabilizers [30 μg/mL cholesterol (CHOL), 80 μmol/L GM1] for 2 h and then left untreated or treated with 25 μmol/L CDDP for 1 h. Analysis of membrane fluidity was done as previously described. Data are expressed as % deviation/NT (mean ± SE of three independent experiments). *, P ≤ 0.05, membrane stabilizers-CDDP versus CDDP (bottom). B, pretreatment with membrane stabilizers prevents cisplatin-induced Fas receptor aggregation at the cell membrane. HT29 cells were pretreated or not (NT) with membrane stabilizers for 2 h and then left untreated or treated with 25 μmol/L CDDP for 4 h. Fas receptor expression and aggregation were evidenced by fluorescence microscopy with an anti-Fas antibody (ZB4) staining. One representative of three independent experiments is shown. C, cisplatin treatment induces lipid raft aggregation at the cell membrane. HT29 cells were treated or not (NT) with 25 μmol/L CDDP for the indicated times. Membrane lipid rafts were evidenced by fluorescence microscopy using cholera toxin-FITC (CTx-FITC) labeling of ganglioside GM1, a major constituent of lipid rafts. One representative of three independent experiments is shown (top). Pretreatment with cholesterol prevents cisplatin-induced lipid rafts aggregation. HT29 cells were pretreated or not (NT) with 30 μg/mL cholesterol (CHOL) for 2 h and then left untreated or treated with 25 μmol/L CDDP for 1 h. Membrane lipid rafts were evidenced by fluorescence microscopy using CTx-FITC staining. One representative of three independent experiments is shown (bottom). D, pretreatment with membrane stabilizers reduces cisplatin-induced apoptosis. HT29 cells were pretreated or not (NT) with membrane stabilizers for 2 h and then left untreated or treated with 25 μmol/L CDDP for 72 h. Percentages of apoptotic cells were estimated by nuclear chromatin staining with Hoechst 33342. Data are expressed as mean ± SE of three independent experiments. **, P ≤ 0.01; ***, P ≤ 0.001, membrane stabilizers-CDDP versus CDDP (top). Pretreatment with cholesterol reduces cisplatin-induced caspase-3 activation. HT29 cells were pretreated or not (NT) with 30 μg/mL cholesterol (CHOL) for 2 h and then left untreated or treated with 25 μmol/L CDDP for 72 h. Caspase-3 activation was measured in lysates by the cleavage of the DEVD-AMC peptide substrate. Data are expressed in arbitrary units (AU) as mean ± SE of three independent experiments. ***, P ≤ 0.001, cholesterol-CDDP versus CDDP (bottom).

Figure 1.

Cisplatin-induced apoptosis is dependent on increased membrane fluidity. A, cisplatin treatment increases fluidity in bulk membranes as well as in membrane lipid rafts. HT29 cells were treated or not (NT) with 25 μmol/L cisplatin (CDDP) for 1 h. Membrane lipid rafts were purified by ultracentrifugation on a discontinuous sucrose gradient. Membrane fluidity was determined on bulk membranes or isolated lipid rafts by EPR using a spin-labeling method. The EPR spectra were used to calculate the membrane order parameter S, which is conversely proportional to membrane fluidity. Values are the mean ± SE of three independent experiments. **, P ≤ 0.01, CDDP versus NT (top). Pretreatment with membrane stabilizers inhibits cisplatin-induced increase in membrane fluidity. HT29 cells were pretreated or not (CONTROL) with membrane stabilizers [30 μg/mL cholesterol (CHOL), 80 μmol/L GM1] for 2 h and then left untreated or treated with 25 μmol/L CDDP for 1 h. Analysis of membrane fluidity was done as previously described. Data are expressed as % deviation/NT (mean ± SE of three independent experiments). *, P ≤ 0.05, membrane stabilizers-CDDP versus CDDP (bottom). B, pretreatment with membrane stabilizers prevents cisplatin-induced Fas receptor aggregation at the cell membrane. HT29 cells were pretreated or not (NT) with membrane stabilizers for 2 h and then left untreated or treated with 25 μmol/L CDDP for 4 h. Fas receptor expression and aggregation were evidenced by fluorescence microscopy with an anti-Fas antibody (ZB4) staining. One representative of three independent experiments is shown. C, cisplatin treatment induces lipid raft aggregation at the cell membrane. HT29 cells were treated or not (NT) with 25 μmol/L CDDP for the indicated times. Membrane lipid rafts were evidenced by fluorescence microscopy using cholera toxin-FITC (CTx-FITC) labeling of ganglioside GM1, a major constituent of lipid rafts. One representative of three independent experiments is shown (top). Pretreatment with cholesterol prevents cisplatin-induced lipid rafts aggregation. HT29 cells were pretreated or not (NT) with 30 μg/mL cholesterol (CHOL) for 2 h and then left untreated or treated with 25 μmol/L CDDP for 1 h. Membrane lipid rafts were evidenced by fluorescence microscopy using CTx-FITC staining. One representative of three independent experiments is shown (bottom). D, pretreatment with membrane stabilizers reduces cisplatin-induced apoptosis. HT29 cells were pretreated or not (NT) with membrane stabilizers for 2 h and then left untreated or treated with 25 μmol/L CDDP for 72 h. Percentages of apoptotic cells were estimated by nuclear chromatin staining with Hoechst 33342. Data are expressed as mean ± SE of three independent experiments. **, P ≤ 0.01; ***, P ≤ 0.001, membrane stabilizers-CDDP versus CDDP (top). Pretreatment with cholesterol reduces cisplatin-induced caspase-3 activation. HT29 cells were pretreated or not (NT) with 30 μg/mL cholesterol (CHOL) for 2 h and then left untreated or treated with 25 μmol/L CDDP for 72 h. Caspase-3 activation was measured in lysates by the cleavage of the DEVD-AMC peptide substrate. Data are expressed in arbitrary units (AU) as mean ± SE of three independent experiments. ***, P ≤ 0.001, cholesterol-CDDP versus CDDP (bottom).

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Role of increased membrane fluidity in cisplatin-induced apoptotic pathway. To find out whether the increase in membrane fluidity after cisplatin treatment might underlie the apoptotic effect of this anticancer drug, experiments were carried out using cholesterol or GM1 ganglioside, two natural lipid constituents of cell membranes, that have been shown to protect rat hepatocytes against ethanol-induced apoptosis (12). Here, 30 μg/mL cholesterol or 80 μmol/L GM1 were added 2 h before addition of 25 μmol/L cisplatin for 1 h. Although these agents did not significantly change the basal cell membrane fluidity, they fully prevented the effect of cisplatin on this parameter (Fig. 1A,, bottom). We have previously shown that cisplatin induced Fas receptor aggregation on the cell membrane of HT29 cells (26). Here, we showed that pretreatment of HT29 cells with cholesterol or GM1 completely blocked cisplatin-induced Fas receptor aggregation on the cell membrane (Fig. 1B). Moreover, formation of large lipid raft aggregates was readily detectable on HT29 cells by staining GM1 ganglioside with the fluorescein-tagged cholera toxin (CTx-FITC) between 15 min and 4 h after the beginning of cell treatment with 25 μmol/L cisplatin (Fig. 1C,, top). The cisplatin-induced lipid raft aggregation was also completely blocked by pretreatment of HT29 cells with cholesterol (Fig. 1C,, bottom). Finally, cholesterol or GM1 pretreatment significantly inhibited cisplatin-induced apoptosis evidenced by the decreased percentage of fragmented and condensed nuclei (Fig. 1D,, top) or by the decreased caspase-3 activity (Fig. 1D , bottom).

The inhibition of cisplatin-induced apoptosis by cholesterol or GM1 could have been related to a decreased accumulation of platinum in HT29 cells. To test this hypothesis, we first studied the intracellular platinum uptake by using graphite furnace atomic absorption spectroscopy. We showed that the accumulation of platinum in HT29 cells increased with time and did not yet reach a plateau at a 24-h cisplatin treatment (Fig. 2A). Moreover, cholesterol or GM1 pretreatment did not modify the uptake of platinum in HT29 cells (Fig. 2B). Finally, we measured the time-course formation of intrastrand G-G DNA adducts (cis-Pt(NH3)2d(pGpG)–abbreviated 1,2 Pt-[GG]) in tumor cell DNA by Southwestern blot analysis using the specific monoclonal antibody R-C18 (30). The formation of 1,2 Pt-[GG] was time dependent and increased from 1 to 12 h after the beginning of 25 μmol/L cisplatin treatment and decreased after 48 h (Fig. 2C,, left). As the platinum uptake, the formation of 1,2 Pt-[GG] was not modified by pretreatment of HT29 cells with cholesterol (Fig. 2C , right).

Figure 2.

Platinum accumulation and formation of 1,2 Pt-[GG] adducts in HT29 cells treated with cisplatin. A, platinum content increases with time exposure to cisplatin. HT29 cells were treated with 25 μmol/L CDDP for the indicated times. Platinum content was determined by flameless atomic absorption spectroscopy. Data are expressed as mean ± SE of three independent experiments. B, pretreatment with membrane stabilizers does not prevent platinum uptake by HT29 cells. HT29 cells were pretreated or not with membrane stabilizers [30 μg/mL cholesterol (CHOL), 80 μmol/L GM1] for 2 h and then treated with CDDP 25 μmol/L for 1 h. Platinum content was determined as described above. Data are expressed as mean ± SE of three independent experiments. C, formation of 1,2 Pt-[GG] adducts increases with exposure time to cisplatin and is not inhibited by cholesterol pretreatment. HT29 cells were pretreated or not with cholesterol and then treated with 25 μmol/L CDDP for the indicated times. 1,2 Pt-[GG] formation was detected by Southwestern blot analysis as described in Materials and Methods. One representative of three independent experiments is shown.

Figure 2.

Platinum accumulation and formation of 1,2 Pt-[GG] adducts in HT29 cells treated with cisplatin. A, platinum content increases with time exposure to cisplatin. HT29 cells were treated with 25 μmol/L CDDP for the indicated times. Platinum content was determined by flameless atomic absorption spectroscopy. Data are expressed as mean ± SE of three independent experiments. B, pretreatment with membrane stabilizers does not prevent platinum uptake by HT29 cells. HT29 cells were pretreated or not with membrane stabilizers [30 μg/mL cholesterol (CHOL), 80 μmol/L GM1] for 2 h and then treated with CDDP 25 μmol/L for 1 h. Platinum content was determined as described above. Data are expressed as mean ± SE of three independent experiments. C, formation of 1,2 Pt-[GG] adducts increases with exposure time to cisplatin and is not inhibited by cholesterol pretreatment. HT29 cells were pretreated or not with cholesterol and then treated with 25 μmol/L CDDP for the indicated times. 1,2 Pt-[GG] formation was detected by Southwestern blot analysis as described in Materials and Methods. One representative of three independent experiments is shown.

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These data show for the first time that the early increase in membrane fluidity participates in cisplatin-induced apoptosis and facilitates the aggregation of Fas receptor and of lipid rafts on HT29 cell membrane. This alteration of membrane fluidity occurs independently of cisplatin DNA adducts formation because the increase in membrane fluidity was detected as soon as 15 min after cisplatin treatment (20), whereas the formation of 1,2 Pt-[GG] was detectable only after a 1-h treatment with cisplatin (Fig. 2C).

Mechanism of cisplatin-induced increase in membrane fluidity. We have previously shown that cisplatin-induced apoptosis is inhibited by imipramine, an inhibitor of aSMase, and involves a rapid ceramide generation in HT29 cells (20); this suggested the role of aSMase in cisplatin-induced cell death pathway. To confirm such a role, we did RNA interference with aSMase small interfering RNAs (siRNA) that have been shown to decrease aSMase RNA expression in HT29 cells (31). We observed that transient transfection with aSMase siRNAs (si aSMase) decreased aSMase RNA expression level and cisplatin-induced apoptosis (Fig. 3A). On the contrary, transfection with nonspecific siRNAs (si GFP) neither affected cell death induction nor aSMase RNA expression (Fig. 3A). Following these results, we studied the kinetics of aSMase activation after treatment with 25 μmol/L cisplatin and showed an ∼25% increase in aSMase activity, which peaked within 5 to 10 min after the beginning of cisplatin treatment (Fig. 3B,, left). To look for the specificity of this enzymatic assay, HT29 cells were preincubated for 1 h with 30 μmol/L SR, a potent aSMase inhibitor (32) and then treated with cisplatin for 10 min. As expected, SR completely abrogated cisplatin-induced aSMase activity (Fig. 3B,, right). Because the sphingomyelin (SM) is predominantly present in the outer leaflet of the cell membrane, we then studied ceramide generation on the cell surface of HT29 cells by using an anti-ceramide monoclonal antibody (15B4; ref. 20). By fluorescence microscopy, ceramide was detected and seemed to be aggregated on the cell membrane of HT29 cells after a 30-min cisplatin treatment (Fig. 3C,, left). Moreover, by using a flow cytometry analysis, the anti-ceramide labeling (FL-1) increased on the cell surface of HT29 cells after treatment with cisplatin as well as after cell treatment with an exogenous SMase from S. aureus (0.01 unit/mL for 30 min), which was used as a positive control (Fig. 3C,, right). We then studied the effect of S. aureus SMase on the membrane fluidity measured by EPR in HT29 cells. The exposure of HT29 cells to 0.01 unit/mL SMase resulted in a marked decrease of the membrane order parameter S in a time-dependent fashion, showing an increase in membrane fluidity (Fig. 3D,, left). Finally, pretreatment of HT29 cells with SR, an inhibitor of aSMase, blocked cisplatin-induced increase in membrane fluidity (Fig. 3D , right). These results suggested that the early activation of aSMase following cisplatin treatment of HT29 cells leads to an early increase in membrane fluidity.

Figure 3.

Cisplatin treatment rapidly activates acid sphingomyelinase and generates membrane ceramide. A, interfering with aSMase mRNA expression reduces cisplatin-induced apoptosis. HT29 cells were transfected with siRNA aSMase (si aSMase) or siRNA green fluorescent protein (si GFP used as a negative control). For reverse transcription-PCR analysis (right), cells were harvested 48 h after transfection. For cell death analysis, following 24 h of cell transfection, HT29 cells were treated or not with 25 μmol/L CDDP for 72 h. One representative of three independent experiments is shown. B, cisplatin induces early activation of aSMase. HT29 cells were treated with 25 μmol/L CDDP for the indicated times, and aSMase activity was determined as described in Materials and Methods. Data are expressed as % of control (0; mean ± SE of three independent experiments). *, P ≤ 0.05; ***, P ≤ 0.001, CDDP versus control (0; left). Pretreatment with SR inhibits cisplatin-induced aSMase activation. HT29 cells were pretreated or not (NT) with 30 μmol/L SR for 1 h and then left untreated or treated with 25 μmol/L CDDP for 10 min, and aSMase activity was determined as in the left panel. Data are expressed as % of control (NT; mean ± SE of three independent experiments). *, P ≤ 0.05, SR-CDDP versus CDDP; ***, P ≤ 0.001, CDDP versus NT (right). C, cisplatin induces ceramide generation on cell membrane. HT29 cells were treated or not (NT) with 25 μmol/L CDDP or 0.01 units/mL S. aureus SMase for 30 min, and ceramide expression was studied by fluorescence microscopy (left) or by flow cytometry (right) using a mouse monoclonal anti-CER (15B4; open peak) and an isotype-matched control (full peak). One representative of three independent experiments is shown. D, aSMase is involved in cisplatin-induced increase in membrane fluidity. Treatment of HT29 cells with exogenous S. aureus SMase increases membrane fluidity. HT29 cells were treated or not (NT) with 0.01 units/mL S. aureus SMase for the indicated times, and membrane fluidity was measured by EPR as described in Fig. 1. Data are expressed as % deviation/NT (mean ± SE of three independent experiments). *, P ≤ 0.05; **, P ≤ 0.01, treated versus nontreated (NT; left). Pretreatment with SR inhibits cisplatin-induced increase in membrane fluidity. HT29 cells were pretreated or not (NT) with 30 μmol/L SR for 1 h and then left untreated or treated with 25 μmol/L CDDP for 1 h. Membrane fluidity was analyzed as described above. Data are expressed as % deviation/NT (mean ± SE of three independent experiments). *, P ≤ 0.05, CDDP versus NT; **, P ≤ 0.01, SR-CDDP versus CDDP (right).

Figure 3.

Cisplatin treatment rapidly activates acid sphingomyelinase and generates membrane ceramide. A, interfering with aSMase mRNA expression reduces cisplatin-induced apoptosis. HT29 cells were transfected with siRNA aSMase (si aSMase) or siRNA green fluorescent protein (si GFP used as a negative control). For reverse transcription-PCR analysis (right), cells were harvested 48 h after transfection. For cell death analysis, following 24 h of cell transfection, HT29 cells were treated or not with 25 μmol/L CDDP for 72 h. One representative of three independent experiments is shown. B, cisplatin induces early activation of aSMase. HT29 cells were treated with 25 μmol/L CDDP for the indicated times, and aSMase activity was determined as described in Materials and Methods. Data are expressed as % of control (0; mean ± SE of three independent experiments). *, P ≤ 0.05; ***, P ≤ 0.001, CDDP versus control (0; left). Pretreatment with SR inhibits cisplatin-induced aSMase activation. HT29 cells were pretreated or not (NT) with 30 μmol/L SR for 1 h and then left untreated or treated with 25 μmol/L CDDP for 10 min, and aSMase activity was determined as in the left panel. Data are expressed as % of control (NT; mean ± SE of three independent experiments). *, P ≤ 0.05, SR-CDDP versus CDDP; ***, P ≤ 0.001, CDDP versus NT (right). C, cisplatin induces ceramide generation on cell membrane. HT29 cells were treated or not (NT) with 25 μmol/L CDDP or 0.01 units/mL S. aureus SMase for 30 min, and ceramide expression was studied by fluorescence microscopy (left) or by flow cytometry (right) using a mouse monoclonal anti-CER (15B4; open peak) and an isotype-matched control (full peak). One representative of three independent experiments is shown. D, aSMase is involved in cisplatin-induced increase in membrane fluidity. Treatment of HT29 cells with exogenous S. aureus SMase increases membrane fluidity. HT29 cells were treated or not (NT) with 0.01 units/mL S. aureus SMase for the indicated times, and membrane fluidity was measured by EPR as described in Fig. 1. Data are expressed as % deviation/NT (mean ± SE of three independent experiments). *, P ≤ 0.05; **, P ≤ 0.01, treated versus nontreated (NT; left). Pretreatment with SR inhibits cisplatin-induced increase in membrane fluidity. HT29 cells were pretreated or not (NT) with 30 μmol/L SR for 1 h and then left untreated or treated with 25 μmol/L CDDP for 1 h. Membrane fluidity was analyzed as described above. Data are expressed as % deviation/NT (mean ± SE of three independent experiments). *, P ≤ 0.05, CDDP versus NT; **, P ≤ 0.01, SR-CDDP versus CDDP (right).

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Role of NHE1 in cisplatin-induced aSMase activation and in cisplatin-induced apoptosis. Because the activity of aSMase may be regulated by acidic pH, we measured pHi by microspectrofluorimetry using the pH-sensitive fluorescent probe, carboxy-SNARF-1, and showed a very rapid decrease of about 0.05 to 0.1 pH unit at 5 and 15 min after cisplatin treatment in HT29 cells (Fig. 4A,, left). The cisplatin-induced acidification was prevented by a 2-h pretreatment with cariporide, suggesting that the cytosolic acidification was due to the inhibition of plasma membrane NHE1 by cisplatin (Fig. 4A,, left). To confirm this hypothesis, we next measured the rate of pHi recovery following an acid load induced by an NH4+ prepulse in HT29 cells treated or not with 25 μmol/L CDDP. As expected, the rate of pHi recovery was markedly decreased after cisplatin exposure due to NHE1 inhibition and was completely inhibited by 30 μmol/L cariporide (Fig. 4A , right).

Figure 4.

Cisplatin induces a NHE1-dependent intracellular acidification, which is involved in cisplatin-induced aSMase activation, membrane fluidification, and apoptosis. A, pretreatment with cariporide inhibits cisplatin-induced intracellular acidification. HT29 cells were pretreated or not (NT) with 30 μmol/L cariporide (CAR) for 2 h and then left untreated or treated with 25 μmol/L CDDP for the indicated times. Intracellular pH (pHi) measurements were done using the pH-sensitive fluorescent probe, carboxy-SNARF-1 as described in Materials and Methods. Data are expressed as ΔpHi (treated − control; mean ± SE of nine experiments). **, P ≤ 0.01; ***, P ≤ 0.001; CAR-CDDP versus CDDP (left). Cisplatin inhibits NHE1 exchanger. The time course of pHi recovery due to NHE1 activity, following an acid load induced by an NH4+ prepulse, was estimated from HT29 cells treated or not (control) with 25 μmol/L CDDP or 30 μmol/L cariporide (CAR). One representative of eight experiments is shown (right). B, pretreatment with cariporide inhibits cisplatin-induced aSMase activation. HT29 cells were pretreated or not (NT) with 30 μmol/L cariporide (CAR) for 2 h and then left untreated or treated with 25 μmol/L CDDP for 10 min. aSMase activity was done as described in Materials and Methods. Data are expressed as % of control (NT; mean ± SE of three independent experiments). *, P ≤ 0.05, CAR-CDDP versus CDDP; **, P ≤ 0.01, CDDP versus NT (left). Pretreatment with cariporide inhibits cisplatin-induced ceramide generation at the cell membrane. HT29 cells were pretreated or not (NT) with 30 μmol/L cariporide (CAR) for 2 h and then left untreated or treated with 25 μmol/L CDDP for 30 min. Ceramide membrane expression was studied by flow cytometry using a mouse monoclonal anti-CER (15B4; open peak) and an isotype-matched control (filled peak). One representative of three independent experiments is shown (right). C, pretreatment with cariporide inhibits cisplatin-induced increase in membrane fluidity. HT29 cells were pretreated or not (NT) with 30 μmol/L cariporide (CAR) for 2 h and then left untreated or treated with 25 μmol/L CDDP for 1 h. Membrane fluidity was measured as previously described in Fig. 1. Data are expressed as % deviation/NT (mean ± SE of three independent experiments).*, P ≤ 0.05, CDDP versus NT, CAR-CDDP versus CDDP. D, pretreatment with cariporide significantly inhibits cisplatin-induced apoptosis. HT29 cells were pretreated or not (NT) with 30 μmol/L cariporide (CAR) for 2 h and then left untreated or treated with 25 μmol/L CDDP for 72 h. Percentage of apoptosis was determined as in Fig. 1D. Data are expressed as mean ± SE of three independent experiments. *, P ≤ 0.05, CAR-CDDP versus CDDP.

Figure 4.

Cisplatin induces a NHE1-dependent intracellular acidification, which is involved in cisplatin-induced aSMase activation, membrane fluidification, and apoptosis. A, pretreatment with cariporide inhibits cisplatin-induced intracellular acidification. HT29 cells were pretreated or not (NT) with 30 μmol/L cariporide (CAR) for 2 h and then left untreated or treated with 25 μmol/L CDDP for the indicated times. Intracellular pH (pHi) measurements were done using the pH-sensitive fluorescent probe, carboxy-SNARF-1 as described in Materials and Methods. Data are expressed as ΔpHi (treated − control; mean ± SE of nine experiments). **, P ≤ 0.01; ***, P ≤ 0.001; CAR-CDDP versus CDDP (left). Cisplatin inhibits NHE1 exchanger. The time course of pHi recovery due to NHE1 activity, following an acid load induced by an NH4+ prepulse, was estimated from HT29 cells treated or not (control) with 25 μmol/L CDDP or 30 μmol/L cariporide (CAR). One representative of eight experiments is shown (right). B, pretreatment with cariporide inhibits cisplatin-induced aSMase activation. HT29 cells were pretreated or not (NT) with 30 μmol/L cariporide (CAR) for 2 h and then left untreated or treated with 25 μmol/L CDDP for 10 min. aSMase activity was done as described in Materials and Methods. Data are expressed as % of control (NT; mean ± SE of three independent experiments). *, P ≤ 0.05, CAR-CDDP versus CDDP; **, P ≤ 0.01, CDDP versus NT (left). Pretreatment with cariporide inhibits cisplatin-induced ceramide generation at the cell membrane. HT29 cells were pretreated or not (NT) with 30 μmol/L cariporide (CAR) for 2 h and then left untreated or treated with 25 μmol/L CDDP for 30 min. Ceramide membrane expression was studied by flow cytometry using a mouse monoclonal anti-CER (15B4; open peak) and an isotype-matched control (filled peak). One representative of three independent experiments is shown (right). C, pretreatment with cariporide inhibits cisplatin-induced increase in membrane fluidity. HT29 cells were pretreated or not (NT) with 30 μmol/L cariporide (CAR) for 2 h and then left untreated or treated with 25 μmol/L CDDP for 1 h. Membrane fluidity was measured as previously described in Fig. 1. Data are expressed as % deviation/NT (mean ± SE of three independent experiments).*, P ≤ 0.05, CDDP versus NT, CAR-CDDP versus CDDP. D, pretreatment with cariporide significantly inhibits cisplatin-induced apoptosis. HT29 cells were pretreated or not (NT) with 30 μmol/L cariporide (CAR) for 2 h and then left untreated or treated with 25 μmol/L CDDP for 72 h. Percentage of apoptosis was determined as in Fig. 1D. Data are expressed as mean ± SE of three independent experiments. *, P ≤ 0.05, CAR-CDDP versus CDDP.

Close modal

Moreover, cariporide inhibited cisplatin-induced aSMase activity (Fig. 4B,, left), ceramide generation on the cell membrane (Fig. 4B,, right), as well as increase in membrane fluidity (Fig. 4C), and significantly decreased cisplatin-induced apoptosis (Fig. 4D). Taken together, these results pointed to NHE1 as a potential target of cisplatin in HT29 cells, responsible for early intracellular acidification with consequences on aSMase activity and, hence, on membrane fluidity.

PS120 Chinese hamster fibroblast cell lines expressing or not an active form of NHE1 were used to confirm the role of NHE1 as a potential target of cisplatin. Firstly, we showed that NHE1-PS120 cells were more sensitive than PS120 cells to cisplatin-induced apoptosis (Fig. 5A). As expected, an increase in membrane fluidity was only detected in NHE1-PS120 cells following cisplatin treatment (Fig. 5B). Moreover, pretreatment of NHE1-PS120 cells with SR prevented the increase in fluidity induced by cisplatin treatment (Fig. 5C). Finally, cisplatin-induced apoptosis in NHE1-PS120 cells was reduced by the pretreatment of cells with SR or cariporide (Fig. 5D). Altogether, these data strongly supported that cisplatin induces a very early NHE1-dependent intracellular acidification leading to aSMase activity and increase in membrane fluidity.

Figure 5.

An active NHE1 expression increases sensitivity to cisplatin in PS120 fibroblast cell line. A, NHE1-PS120 cells are more sensitive than NHE1-deficient PS120 cells (PS120) to cisplatin. PS120 or NHE1-PS120 cells were treated or not (0) with increased concentrations of CDDP for 48 h. Percentage of apoptosis was determined as in Fig. 1D. Data are expressed as mean ± SE of three independent experiments. **, P ≤ 0.01; ***, P ≤ 0.001, NHE1-PS120 versus PS120 (left). Caspase-3 activity was enhanced in CDDP-treated NHE1-PS120 cells compared with CDDP-treated PS120 cells. PS120 or NHE1-PS120 cells were treated or not (NT) with 30 μmol/L CDDP for 48 h. Caspase-3 activity was measured as in Fig. 1D. Data are expressed in arbitrary units (AU) as mean ± SE of three independent experiments. *, P ≤ 0.05, NHE1-PS120 versus PS120 (right). B, cisplatin induces an increase in membrane fluidity only in NHE1-PS120 cells. PS120 or NHE1-PS120 cells were treated with 30 μmol/L CDDP for 1 h. Analysis of membrane fluidity was done as described in Fig. 1. Data are expressed as % deviation/NT (mean ± SE of three independent experiments). **, P ≤ 0.01 CDDP versus NT. C, pretreatment with SR inhibits cisplatin-induced increase in membrane fluidity in NHE1-PS120 cells. NHE1-PS120 cells were pretreated or not (NT) with 30 μmol/L SR for 1 h and then left untreated or treated with 30 μmol/L CDDP for 1 h. Analysis of membrane fluidity was done as described in Fig. 1. Data are expressed as % deviation/NT (mean ± SE of three independent experiments). *, P ≤0.05, SR-CDDP versus CDDP; **, P ≤ 0.01, CDDP versus NT. D, pretreatment with SR or cariporide significantly inhibits cisplatin-induced apoptosis in NHE1-PS120 cells. NHE1-PS120 cells were pretreated or not with 30 μmol/L SR or 30 μmol/L cariporide (CAR) for 2 h and then left untreated or treated with 30 μmol/L CDDP for 48 h. Data are expressed as mean ± SE of three independent experiments. ***, P ≤ 0.001, SR-CDDP versus CDDP, CAR-CDDP versus CDDP.

Figure 5.

An active NHE1 expression increases sensitivity to cisplatin in PS120 fibroblast cell line. A, NHE1-PS120 cells are more sensitive than NHE1-deficient PS120 cells (PS120) to cisplatin. PS120 or NHE1-PS120 cells were treated or not (0) with increased concentrations of CDDP for 48 h. Percentage of apoptosis was determined as in Fig. 1D. Data are expressed as mean ± SE of three independent experiments. **, P ≤ 0.01; ***, P ≤ 0.001, NHE1-PS120 versus PS120 (left). Caspase-3 activity was enhanced in CDDP-treated NHE1-PS120 cells compared with CDDP-treated PS120 cells. PS120 or NHE1-PS120 cells were treated or not (NT) with 30 μmol/L CDDP for 48 h. Caspase-3 activity was measured as in Fig. 1D. Data are expressed in arbitrary units (AU) as mean ± SE of three independent experiments. *, P ≤ 0.05, NHE1-PS120 versus PS120 (right). B, cisplatin induces an increase in membrane fluidity only in NHE1-PS120 cells. PS120 or NHE1-PS120 cells were treated with 30 μmol/L CDDP for 1 h. Analysis of membrane fluidity was done as described in Fig. 1. Data are expressed as % deviation/NT (mean ± SE of three independent experiments). **, P ≤ 0.01 CDDP versus NT. C, pretreatment with SR inhibits cisplatin-induced increase in membrane fluidity in NHE1-PS120 cells. NHE1-PS120 cells were pretreated or not (NT) with 30 μmol/L SR for 1 h and then left untreated or treated with 30 μmol/L CDDP for 1 h. Analysis of membrane fluidity was done as described in Fig. 1. Data are expressed as % deviation/NT (mean ± SE of three independent experiments). *, P ≤0.05, SR-CDDP versus CDDP; **, P ≤ 0.01, CDDP versus NT. D, pretreatment with SR or cariporide significantly inhibits cisplatin-induced apoptosis in NHE1-PS120 cells. NHE1-PS120 cells were pretreated or not with 30 μmol/L SR or 30 μmol/L cariporide (CAR) for 2 h and then left untreated or treated with 30 μmol/L CDDP for 48 h. Data are expressed as mean ± SE of three independent experiments. ***, P ≤ 0.001, SR-CDDP versus CDDP, CAR-CDDP versus CDDP.

Close modal

Cariporide and cholesterol also inhibited cisplatin-induced apoptosis in two other human colon cancer cell lines HCT116 and SW480. HCT116 and SW480 were pretreated with cariporide or cholesterol for 2 h before cisplatin treatment. In agreement with results obtained in HT29 cells, cariporide and cholesterol pretreatment also reduced cisplatin-induced apoptosis in these cell lines (Fig. 6), thus suggesting that inhibition of NHE1 or membrane fluidification might be common early plasma membrane events involved in cisplatin-induced apoptosis in human colon cancer cells.

Figure 6.

Pretreatment with cariporide or cholesterol significantly inhibits cisplatin-induced apoptosis in HCT116 and SW480 human colon cancer cells. HCT116 and SW480 cells were pretreated or not (NT) with 30 μmol/L cariporide (CAR) or 30 μg/mL cholesterol (CHOL) for 2 h and then left untreated or treated with 25 μmol/L CDDP for 48 h. Percentage of apoptosis was determined as in Fig. 1D. Data are expressed as mean ± SE of three independent experiments. *, P ≤ 0.05, CAR-CDDP versus CDDP or cholesterol-CDDP versus CDDP.

Figure 6.

Pretreatment with cariporide or cholesterol significantly inhibits cisplatin-induced apoptosis in HCT116 and SW480 human colon cancer cells. HCT116 and SW480 cells were pretreated or not (NT) with 30 μmol/L cariporide (CAR) or 30 μg/mL cholesterol (CHOL) for 2 h and then left untreated or treated with 25 μmol/L CDDP for 48 h. Percentage of apoptosis was determined as in Fig. 1D. Data are expressed as mean ± SE of three independent experiments. *, P ≤ 0.05, CAR-CDDP versus CDDP or cholesterol-CDDP versus CDDP.

Close modal

In this study, we showed for the first time that, in vitro, cisplatin treatment induced a rapid increase in fluidity measured by EPR spectroscopy in bulk membranes as well as in isolated lipid rafts in HT29 human colon cancer cells. Pretreatment with membrane stabilizers (cholesterol or GM1) totally inhibited the increase in membrane fluidity induced by cisplatin, but also the aggregation of lipid rafts and of Fas receptor on cell membrane; thus, this suggested for the first time a role of increased membrane fluidity in the reorganization of certain plasma membrane components in HT29 cells following cisplatin treatment. Until now, such variations of membrane fluidity were supposed to induce a mobility of the membrane components and a reorganization of membrane proteins and/or lipid rafts likely leading to the modulation of many cellular functions (33). Our data support these hypotheses and, although increased membrane fluidity is associated with a more fluid environment which could disorganize the membrane, we here show that the membrane order parameter S measured in lipid rafts was higher than that measured in bulk membranes of both untreated and cisplatin-treated HT29 cells. This suggested that the membrane of cisplatin-treated cells may still be organized in less fluid-state domains similar to lipid rafts evidenced by CTx-FITC staining of GM1, a constituent of membrane microdomains. Finally, pretreatment with cholesterol or GM1 significantly reduced cisplatin-induced apoptosis, suggesting that the increase in membrane fluidity was involved in cisplatin-induced apoptosis.

Membrane fluidity is probably the most important physicochemical property of cell membranes involved in many cellular processes, including response to cancer therapy (34). However, the molecular mechanisms involved in alterations of membrane fluidity upon drug exposure require definition; such alterations could involve membrane-drug interaction, modification of membrane lipid composition, membrane lipid peroxidation, cytoskeleton alteration, or activation of sphingomyelinases such as the acid sphingomyelinase (aSMase; refs. 12, 35). Our present study confirmed that aSMase was involved in cisplatin-induced apoptosis because decreased expression of aSMase by RNA interference (31) significantly reduced cisplatin-induced apoptosis in HT29 cells. Moreover, cisplatin induced an early aSMase activation in the first 15 min following the onset of cisplatin treatment and, subsequently, the generation of ceramide at the plasma membrane. Both these events were totally prevented by pretreatment of HT29 cells with SR, a potent inhibitor of aSMase (32). These results corroborate our previous data and other data showing a rapid hydrolysis of sphingomyelin into ceramide and the formation of ceramide-enriched membrane domains upon cisplatin, irradiation, or UV treatment (ref. 20; for a review, see ref. 21).

Recently, it has been reported that SMase activity had several effects on membrane architecture (increased membrane permeability, membrane aggregation, and fusion; refs. 36, 37). Treatment with exogenous sphingomyelinase that decreased cholesterol content in crude plasma membrane has been shown to increase membrane fluidity in preadipocytes (38) and a decrease in cholesterol in detergent-resistant membrane fractions has been described in rat astrocytes treated with SMase (39). Finally, ceramide displaces cholesterol from lipid rafts in both model membranes (40) and cells (41). These data led us to investigate the relationship between aSMase activation and the increase in membrane fluidity following cisplatin treatment in HT29 cells. Firstly, we observed that the treatment of HT29 cells with an exogenous S. aureus sphingomyelinase induced an increase in membrane fluidity. Moreover, pretreatment with SR also prevented the increase in membrane fluidity induced by cisplatin. Altogether, these data suggest for the first time that cisplatin-induced increase in membrane fluidity was dependent on aSMase activation. It is noteworthy that staurosporine-induced apoptosis is independent of aSMase activation in several cell types, including HT29 (20, 42, 43), and was not related to change in membrane fluidity in HT29 cells (data not shown).

At present, little is known about the molecular mechanisms involved in cisplatin-induced aSMase activation. Although the enzyme activity reaches an optimum at acidic pH, the full activity of the enzyme at the cell surface is achieved even at neutral pH values (44). Here, we show that in HT29 cells, cisplatin induced a rapid intracellular acidification due to an inhibition of membrane NHE1 protein. Cariporide, an inhibitor of NHE1, completely prevented cisplatin-induced intracellular acidification as well as the related aSMase activation, ceramide generation, and membrane fluidification, thus pointing to a role of intracellular acidification in aSMase activation.

In tumors, the transmembrane pH gradient is reversed, leading to a more acidic extracellular environment and a more alkaline intracellular medium (45). Besides, it has been shown that cancer cells had a constitutively higher pHi than normal cells as a result of increased activity of NHE1. Because the inhibition of NHE1 can induce apoptosis specifically in cancer cells, NHE1 inhibitors have been considered as potential antitumor agents (46). Accordingly, we observed that transient transfection with specific siRNA NHE1 induced apoptosis in HT29 cells (data not shown); however, transient inhibition of NHE1 by cariporide for 2 h did not induce apoptosis in human colon cancer cells, but significantly reduced cisplatin-induced apoptosis, suggesting a role of this membrane exchanger in this cell death pathway. Employing Chinese hamster fibroblast cell lines NHE1-PS120 and PS120 that expressed or lacked, respectively, an active NHE1, we firmly confirm a role of NHE1 in cell sensitivity to cisplatin by showing that cisplatin-induced apoptosis was increased in NHE1-PS120 cells and was dependent on both NHE1 and aSMase. Moreover, cisplatin elicited a SR-sensitive increase in membrane fluidity only in NHE1-PS120 cells. Altogether, these data suggest that NHE1 may be another potential target of cisplatin.

The molecular mechanism involved in NHE1 inhibition by cisplatin is yet unknown, but seems to be independent of cisplatin-induced DNA adduct formation because this inhibition occurred very rapidly (5 min after the onset of cisplatin treatment) and before formation of 1,2 Pt-[GG] adducts (detectable at 1 h of treatment with cisplatin). Indeed, NHE1 inhibition by cisplatin might stem from a direct interaction of the drug with the exchanger because cisplatin could bind to nucleophilic amino acid residues in proteins, including cysteine, methionine, and histidine. Additionally, cisplatin-induced NHE1 inhibition might result from the inhibition of the Na+, K+ ATPase activity by binding this enzyme through lipid carbonyl group (47) or by preventing the formation of a phospho-intermediate of the enzyme (48). A key to the understanding the molecular mechanisms involved in this inhibition might be the observation that NHE-1 inhibition by cisplatin elicits completely different effects than NHE-1 inhibition by a competitive inhibitor, cariporide, or than the absence of this transporter in PS120 cells. This might constitute an area worthy of investigation for future studies. Nevertheless, in the field of cell response to cisplatin, our data suggest that NHE1 might be another potential cellular target of cisplatin in human colon cancer cells, and that cisplatin treatment may be more potent in tumors having a high basal NHE1 activity (49).

In conclusion, it has been shown that cisplatin-induced apoptosis depends on cisplatin interactions with DNA (3) and also, more recently, on interactions with mitochondrial DNA and voltage-dependent anion channel protein in the mitochondrial membrane (50). Our data now suggest that the inhibition of the exchanger NHE1 at the plasma membrane level may also play a role in the cytotoxicity of this agent, notably by activating acid sphingomyelinase and increasing membrane fluidity. Altogether, these findings may serve to define new therapeutic strategies based on cisplatin therapy, taking into account the activity status of NHE1 in human cancers.

Note: Supplementary data for this article are available at Cancer Research Online (http://cancerres.aacrjournals.org/).

Grant support: Ligue Nationale Contre le Cancer (the Côte d'Armor, Ille et Vilaine and Loire-Atlantique Comittees), Rennes Métropole, and the Région Bretagne.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

We thank Prof. Thierry Levade and Drs Jean-Pierre Jaffrézou, Christine Bezombes, and Nathalie Andrieu-Abbadie for helpful discussions on acid sphingomyelinase assay. We also thank Stéphanie Dutertre and the platform of fluorescence microscopy (IFR 140 GFAS, Rennes).

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Supplementary data