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Carcinogenesis Advance Access originally published online on March 28, 2006
Carcinogenesis 2006 27(9):1787-1796; doi:10.1093/carcin/bgl021
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© The Author 2006. Published by Oxford University Press. All rights reserved. For Permissions, please email: journals.permissions@oxfordjournals.org

H1 histamine receptor antagonists induce genotoxic and caspase-2-dependent apoptosis in human melanoma cells

Shawkat-Muhialdin Jangi1, José Luís Díaz-Pérez2, Borja Ochoa-Lizarralde, Itziar Martín-Ruiz, Aintzane Asumendi, Gorka Pérez-Yarza, Jesús Gardeazabal2, José Luis Díaz-Ramón2 and María Dolores Boyano*

Department of Cell Biology and Histology, Faculty of Medicine and Dentistry, University of the Basque Country Leioa E-48940, Vizcaya, Spain
1 Department of Medicine and Pharmacology, Faculty of Medicine, University of Salahaddin Kurdistan-Iraq
2 Department of Dermatology, Cruces Hospital Baracaldo E-48903, Vizcaya, Spain

*To whom correspondence should be addressed. Tel: +34 94 601 5689; Fax: +34 94 601 3266; Email: lola.boyano{at}ehu.es


   Abstract
Top
 Abstract
Introduction
 Material and methods
 Results
 Discussion
 References
 
Previously, we found that the H1 histamine receptor antagonist diphenhydramine induces apoptosis in human acute T-lymphocytic leukemia cells. Since histamine has been shown to act as a growth factor in malignant melanoma cells, we decided to evaluate the in vitro effect of diphenhydramine and other H1 histamine receptor antagonists, such as terfenadine, astemizol and triprolidine on four malignant human melanoma cell lines. These antagonists were found to induce apoptotic cell death in all four melanoma cell lines. Apoptosis was determined by assessment of phosphatidylserine exposure on the surface of the cells and nuclear fragmentation. Importantly, H1 antagonist treatments did not adversely affect the viability of human melanocytes and murine fibroblasts at the same doses and duration of exposure. Treatment of melanoma cells with terfenadine induced DNA damage and caspases 2, 3, 6, 8 and 9 activation. Furthermore, the general caspase inhibitor (z-VAD-FMK) and a selective inhibitor of caspase-2 (z-VDVAD-FMK) protected melanoma cells from terfenadine-induced apoptosis. In contrast, the caspase-8 inhibitor (z-IETD-FMK) was ineffective. In addition, we found that mitochondria are involved in TEF-induced apoptosis, characterized by the dissipation of the mitochondrial transmembrane potential, the release of cytochrome c into the cytosolic compartment and caspase-9 activation. On the basis of these results we conclude that H1 histamine receptor antagonists induce apoptosis in human melanoma cells but not in normal melanocytes and embryonic murine fibroblasts; this apoptosis appears to be caspase-2-dependent and involves the mitochondrial pathway. The present results may contribute to the elaboration of novel therapeutic strategies for the treatment of malignant human melanoma.

Abbreviations: AST, astemizol; Cyt c, cytochrome c; DMEM, Dulbecco's Modified Eagle's Medium; DMSO, dimethyl sulfoxide; DPH, diphenhydramine; FBS, fetal bovine serum; PBS, phosphate-buffered saline; PI, propidium iodide; TEF, terfenadine; TRI, triprolidine


   Introduction
Top
 Abstract
 Introduction
Material and methods
 Results
 Discussion
 References
 
Malignant melanoma is a rapidly spreading skin tumor with a very high invasive capacity and growing incidence (1). The management of disseminated melanoma represents one of the most challenging problems in clinical oncology. Standard treatment consists of a monotherapy regimen with dacarbazine, but the total response is ~15–20% and the median survival period is only ~6–7 months (2). Therefore, the search for new agents capable of selectively killing melanoma cells constitutes an urgent priority.

Histamine is a well-known biogenic amine that has been found to act as a growth modulator in various normal and malignant cells (3,4). Histamine acts in a paracrine and autocrine manner through a group of receptors on the cell membrane known as histamine H1, H2, H3 and H4 receptors through which it regulates several physiological and pathological processes including cell proliferation (5). High levels of histamine have been reported in malignant melanoma cells (6). Furthermore, it has been found that melanoma cells express high levels of both histidine decarboxylase, the unique enzyme responsible for the synthesis of histamine, and H1 histamine receptors (H1 receptors) on their surface (4,7,8). H1 histamine receptor antagonists are classified as specific antagonists such as terfenadine (TEF) and astemizol (AST), and non-specific antagonists such as diphenhydramine (DPH) and triprolidine (TRI). TEF appears to be the first antihistaminic that acts as a highly potent H1 histamine receptor antagonist (9).

Apoptosis or programmed cell death is an active process by which cells appear to commit suicide. It is characterized by cell shrinkage and rounding, membrane blebbing, chromatin condensation and margination, breakdown of chromosomal DNA into internucleosome fragments and caspase activation (10,11). This process enables organisms to control cell number and to eliminate cells that threaten survival. A deregulation of the apoptotic process can lead to insufficient ‘physiological’ cell death, thus contributing to the pathogenesis of various human diseases such as cancer (12). Apoptosis is widely considered to be one of the most important mechanisms by which anticancer agents kill tumor cells. Therefore, any deficiency in the apoptotic cascade could lead to a drug-resistant phenotype (13).

Caspases are aspartate-specific cysteine proteases, which can be classified in general terms as initiator caspases and executive caspases (14,15). Depending on the initiator caspase involved in the apoptotic process, three major pathways have been identified: the death receptor pathway (16), the mitochondrial pathway (17) and the endoplasmic reticulum stress pathway (18). Caspase-2 is one of the initiator caspases that shares sequence homology with caspases 1 and 9. It was the second caspase reported, and it is one of the most conserved caspases among species (19). The precise mechanism of caspase-2 activation is still unknown, although recent findings indicate that it can be activated rapidly by an autoproteolytic mechanism in response to diverse apoptotic signals (20), such as in response to DNA-damaging by antineoplastic agents (19,21). Although its involvement in the mitochondrial apoptotic pathway has been reported previously (22), the complete details of the mechanism by which caspase-2 executes apoptosis are still being worked out.

In this study we found that TEF and other H1 receptor antagonists induced apoptotic cell death in malignant melanoma cells, but not in normal melanocytes and fibroblasts and that the corresponding apoptotic process involves DNA damage, caspase-2 activation and the mitochondrial pathway. These findings may have important therapeutic applications in the treatment of this class of lethal cancer.


   Material and methods
Top
 Abstract
 Introduction
 Material and methods
Results
 Discussion
 References
 
Chemicals and reagents
DPH, TEF, AST, TRI, Dulbecco's Modified Eagle's Medium (DMEM), L-glutamine and streptomycin/penicillin antibiotic solution were all obtained from The Sigma Chemical Co. (St Louis, MO). Propidium iodide (PI), 3,3 dihexyloxacarbo-cyanine iodide [DiOC6(3)], dimethyl sulfoxide (DMSO), bicinchoninic acid solution, poly-L-lysine, paraformaldehyde, picric acid and electrophoresis reagents were from Sigma-Aldrich (Quimica, SA, Madrid, Spain). Culture flasks were from Falcon Plastics, Becton-Dickinson Laboratories (Orangeburg, NY) and fetal bovine serum (FBS) was from Biochrom AG (Berlin, Germany). An XTT viability assay kit was purchased from Roche Molecular Biochemicals and the Annexin V FITC assay kit was from Oncogene (San Diego, CA). An anti-cytochrome c monoclonal antibody was purchased from BD Pharmingen (San Diego, CA), whereas the anti-actin antibody was from Chemicon International (Temecula, CA). Goat anti-mouse IgGs and goat anti-rabbit IgGs were from Zymed Laboratories (South San Francisco, CA.). Polyclonal rabbit anti-cleaved caspase-3 antibody was obtained from Cell Signalling Technology (Danvers, MA) and ALEXA Fluor goat anti-rabbit secondary antibody from Molecular Probes (Europe BV, Leiden, The Nertherlands). The apotarget caspase colorimetric protease assay kit was from Biosource (Europe SA, Nivelles, Belgium). The general caspase inhibitor (Z-VAD-FMK) was purchased from BD pharmingen (San Diego, CA), whereas caspases 2 and 8 inhibitors were from Calbiochem, Germany. Acrylamide and bis-acrylamide solutions and precision Plus Protein Standards were obtained from Bio-Rad Laboratories (Hercules, CA).

Cells and culture conditions
Human melanoma cell lines, A375, HT144, HSs294T (obtained from American Type Culture Collection, ATCC, Rockville, MD) and MJOI (established in our laboratory); the HEM normal melanocyte cell line (obtained from Gentaur Molecular Products, Brussels, Belgium) and mouse embryonic fibroblasts (obtained in our laboratory) were routinely cultured in 175 cm2 flasks and grown in a monolayer in DMEM, supplemented with 10% FBS, 2 mM L-glutamine and antibiotic solution (100 µg/ml streptomycin and 100 IU/ml penicillin) at 37°C in a 5% CO2–95% air–water saturated atmosphere. The density of cultured cells was maintained at ~105 cells/ml. Trypsinization was performed at harvesting. Isolated cells were then cultured in 96-well culture plates for viability and cytotoxicity studies. The cell lines were examined during culture with an inverted microscope (Olympus CK-2, Olympus, Melville, NY).

Cells were incubated for various periods of time with different concentrations of the H1 receptor antagonists, DPH, TRI, AST and TEF. For AST and TEF, stock solutions (50 mM) were prepared by dissolving the compounds in DMSO and then in DMEM to a final concentration of 1–10 µM. The final treatment concentration of DMSO was not >0.05% (v/v) DMSO/culture medium. DPH and TRI were diluted directly in DMEM. Control groups were either untreated or treated with the corresponding doses of DMSO alone.

Cell viability assay
Cells were seeded into flat-bottomed 96-well microtiter plates at a density of 104 cells/well in 100 µl culture medium and allowed to attach to the wells overnight. Subsequently, cells were treated for 6–24 h with different concentrations (0.1–1 mM) of DPH and TRI, and (1–10 µM) of AST and TEF separately, in a serum-free culture medium. Controls without treatment were also included in each experiment. Cell viability was determined by means of a colorimetric XTT viability assay in accordance with the instructions of the supplier. Absorbance at 490 nm was measured using a microtiter (enzyme-linked immunosorbent assay; ELISA) plate reader (ELX 800 Bio-Tek Instruments, Winooski, VT) and cell viability was calculated in relation to untreated control cells as follows: (experimental absorbance/untreated control absorbance) x 100. Six replicate wells were analyzed for each test and each assay was repeated 3–4 times. The product concentration required to inhibit viability by 50% (IC50) was determined using GraphPad Prism software.

Clonogenic assay
Clonogenic assays were used to determine the effect of TEF on cellular colony formation capability. A375 human melanoma cells were trypsinized in the exponential growth phase and single-cell suspension was prepared. The cells were plated overnight to be adhered in a 60 mm plastic dish at a rate of 1000 cells in 2 ml DMEM supplemented with 10% FBS. The cells were treated with 1–10 µM TEF for 10 days at 37°C in a 5% CO2–95% air–water saturated atmosphere. After that, the cells were fixed in ice-cold methanol for 30 min and the colonies were stained with trypan blue. The number of colonies containing >50 cells was calculated per dish using Leitz Labovert FS inverted phase contrast microscope with high optical resolution. All experiments were performed in triplicate and the data presented are average of three independent experiments.

Morphological assay for apoptosis
The morphological features of melanoma cells treated with or without TEF or AST were determined in parallel with viability studies. Superficial changes associated with apoptosis were observed by phase contrast microscopy. Untreated and treated cells were visualized after various incubation periods using a Leitz Labovert FS inverted phase contrast microscope and photographed using a Leica digital camera.

Nuclear staining and fluorescent microscopy
Morphological evaluation of the nuclei of untreated and treated cells was determined by nuclear staining with PI. Briefly, melanoma cells were seeded overnight on sterile glass coverslips in 24-well plates before treating them with various concentrations of the tested products for 2–8 h. Treated and untreated cells on the coverslips were washed twice with phosphate-buffered saline (PBS) and fixed in 1% formaldehyde in PBS for 20 min at room temperature, rinsed three times with PBS and permeabilized with methanol at –20°C for 10 min. Fixed cells were stained with 20 µg/ml PI, and subsequently examined using a Leitz DMRB fluorescence microscope; digital images were taken using a Leica digital camera.

Flow cytometry
Evaluation of exposed membrane phosphatidylserine was performed using an annexin V FITC assay kit. Treated and untreated cells were harvested, stained with Annexin V-FITC and PI and processed according to the manufacturer's protocol.

In order to study the distribution of the cells through the cell-cycle phases, exponentially growing melanoma cells were treated with 10 µM of AST and TEF for 8 h. Untreated and treated cells were harvested by trypsinization, fixed and stained with PI as described previously (23). The intensity of fluorescence emitted by bound PI is directly proportional to the amount of DNA in each sample. Results were expressed as a percentage of cell nuclei with hypodiploid DNA residing in the sub-G0/G1 phase and in G0/G1, S and G2M phases of the cell cycle.

To study the mitochondrial membrane potential ({Delta}{Psi}m), cells were cultured overnight and then treated with 10 µM TEF for various time periods. During the last 30 min of treatment, 40 nM DiOC6(3) was added to the culture. Floating cells in the culture medium and trypsinized adherent cells were collected and washed twice with PBS before being analyzed by flow cytometer. Untreated cells were used as controls. Flow cytometric analysis was performed on at least 10 000 cells using a Coulter EPICS ELITE ESP (EPICS Division Coulter Corp) and results were analyzed using the WinMDI 2.8 program.

Caspase activity assays
The activity of caspases 2, 3, 6, 8 and 9 were measured using a commercially available colorimetric protease Apotag assay kit. Briefly, samples of untreated and treated cells were washed with cold PBS and lysed on ice in 50 µl of cold lyses buffer. Cell lysates were centrifuged at 10 000 g for 1 min to precipitate cellular debris. The protein content of each sample was quantified by means of the Bradford method. Hundred micrograms of protein extracts were incubated with 50 µl of kit products, and assays were performed in triplicate in 96-well plates on the basis of the manufacturer's protocol using a microtiter plate reader at 405 nm. Comparison of the absorbance from apoptotic samples with that of an untreated control allows determination of fold increase in caspase activity.

Caspase inhibitors have been widely used to confirm the dependency of apoptotic processes on caspase activation. Briefly, cells were pre-treated for 1 hour with z-VDVAD-FMK and z-IETD-FMK (specific inhibitors of caspases 2 and 8, respectively) and with a z-VAD-FMK (general caspase inhibitor). Cell viability was subsequently measured using an XTT cell viability kit. Furthermore, caspase-3 activation was detected in A375 melanoma cells after incubation with a specific inhibitor of caspase-2, followed by treatment with 10 µM TEF using immunofluorescence technique. In brief, A375 human melanoma cells were grown over poly-L-lysine-coated glass coverslips in a 24-well plate. Untreated and treated cells with 100 µM caspase-2 inhibitor for 1 h were incubated with 10 µM TEF for 1, 2, 4 and 6 h time points. The trypsinized adherent cells and the floating cells in the supernatant were collected and transferred onto glass slides by cytospin. The cells were fixed with 4% paraformaldehyde containing 0.9% picric acid for 30 min and then permeabilized with SDS 0.1% in PBS for 10 min. The cells were blocked with 10% FBS in PBS for 30 min and then incubated with polyclonal rabbit anticleaved caspase-3 antibody diluted at 1:100 in PBS supplemented with bovine serum albumin (BSA) 1 mg/ml for 2 h at room temperature. The cells were rinsed with PBS and incubated with appropriate ALEXA goat anti-rabbit secondary antibody diluted at 1:200 in PBS/BSA 1 mg/ml for 1 h at room temperature. After three rinses with PBS, the cells were incubated with 4,6-diamidino-2-phenylindole (DAPI) for 10 min at room temperature to stain the nucleus. Lastly, the cells were viewed and analyzed with a confocal microscope (Olimpus FV 500). Images were obtained by sequential acquisition to avoid overlapping. The cells of 10 microscopic fields were counted and the percentage of caspase-3 positive cells was calculated.

Comet assay
Treated and untreated A375 melanoma cells were harvested by trypsinization, centrifuged and resuspended into 0.5 ml of 1x PBS. The comet assay was basically performed as described by Hajji et al. (24). Cells were mixed with 85 µl of 0.7% low melting agarose (LMA, Sigma) in 1x PBS at 37°C. The mixture was rapidly seeded on the standard slides pre-coated with normal melting agarose (1%) and kept at 4°C for ~10 min to solidify the LMA. Then, another layer of LMA was added in the same manner. The slides were immersed in a chilled lysis solution and kept at 4°C in the dark for at least 1 h to lyse the cells and to allow DNA unfolding. The slides were then drained and left in horizontal gel electrophoresis unit filled with chilled fresh alkaline solution at 4°C and pH 12.8, for 20 min pre-electrophoresis to allow the unwinding of DNA. Then, electrophoresis was carried out at low temperature (4°C) for 20 min at 1.6 V/cm and 300 mA. In order to prevent additional DNA damage, all the steps described above were conducted under yellow light or in the dark. After electrophoresis, slides were gently washed in a neutralization buffer to remove alkali and detergent, stained with PI (2 µg/ml), covered with coverslips and visualized using a Leitz DMRB fluorescence microscope; digital images were taken using a Leica digital camera.

SDS–PAGE and immunoblotting
Melanoma cell cultures treated with and without 10 µM TEF for 2–6 h were harvested by trypsinization. Cytosol and mitochondrial extracts were obtained by digitonin fractionation as previously described by Ramsby et al. (25). To detect cytochrome c (Cyt c) release into the cytosol during TEF-induced apoptosis, samples of cytosolic and mitochondrial extracts containing 40 µg of protein were subjected to electrophoresis using 15% polyacrylamide gel, transferred and developed as described by Asumendi et al. (26). An anti-cytochrome c monoclonal antibody was used at a concentration of 2 µg/ml and an anti-actin antibody at a 1:200 dilution was used as a control.

Statistical analysis
The level of statistical significance between sample means was determined using the Student t-test; P < 0.05 was considered to be statistically significant. Non-linear regression analysis was performed using GraphPad Prism version 3 for Windows (GraphPad Software, San Diego, CA) to calculate the IC50 values for assayed reagents.


   Results
Top
 Abstract
 Introduction
 Material and methods
 Results
Discussion
 References
 
H1 receptor antagonists induce time- and dose-dependent apoptotic cell death in four human melanoma cell lines
The effect of the non-specific H1 receptor antagonists DPH and TRI was characterized on the survival of four human melanoma cell lines. We found that both DPH and TRI at concentrations of 0.2–1 mM were cytotoxic for A375, Hs294T and HT144 melanoma cell lines as determined by the XTT viability assay. However, MJOI cells were more resistant to these treatments. Figure 1A and B represents the effect of DPH and TRI (both at 1 mM) following 8, 12 and 24 h exposure. Since DPH and TRI are non-specific H1 receptor antagonists, we tested the effect of AST and TEF, two specific H1 receptor antagonists, at 1–10 µM. We found that both antihistaminics also induced time- and dose-dependent cytotoxicity in the four melanoma cell lines; once again, the MJOI cell line was more resistant to this treatment (Figure 1C and D). Figure 1E shows the dose-dependent cytotoxicity induced by AST and TEF on A375 and Hs294T melanoma cell lines. The half-maximal effective concentration (IC50) values for AST in the A375 and Hs294T cell lines after 8 h of treatment were 8.2 and 7.4 µM, respectively. The IC50 values for TEF were 6.8 and 6 µM, respectively, indicating that TEF is more effective than AST in both of these cell lines. Finally, we analyzed the effect of TEF on normal human melanocytes and embryonic murine fibroblasts and, interestingly, no cytotoxic effect was observed at any tested concentration following 8 h of treatment (Figure 1F).


Figure 1
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  		Fig. 1 Effects of H1 receptor antagonists on melanoma cell viability. Cells were treated with (A) DPH, (B) TRI, (C) AST and (D) TEF for the indicated times, and cell viability was assessed by means of an XTT viability assay. (E) The dose-dependent effect of AST and TEF on A375 and Hs294T melanoma cells. (F) The effect of TEF on the viability of normal human melanocytes and murine embryonic fibroblasts. Data represent mean ± SD of six determinations of three separate experiments for each cell line and for each product. All of the tested H1 receptor antagonists significantly reduced cell viability in a time-dependent fashion in A375, Hs294T and HT144 melanoma cell lines (P < 0.01). This reduction was somewhat less significant (P < 0.05) in the MJOI melanoma cell line. No statistically significant reduction in cell viability was observed in normal melanocytes and murine fibroblasts.

 
In order to test the effectiveness of TEF in inhibiting melanoma cell proliferation, colony assays were carried out for 10 days in the presence of 10% FBS. Figure 2 shows the data on colony-forming ability of both A375 and HT144 melanoma cells. In agreement with the XTT data, TEF has shown a dose-dependent negative effect on the clonogenic activity of both cell lines.


Figure 2
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  		Fig. 2 Colony-forming ability in A375 and HT144 human melanoma cells. The cells were treated with 2–10 µM TEF; after 10 days of culture in the presence of 10% FBS, the number of colonies with >50 cells was counted. The histograms represented the percentage of colonies with respect to the untreated control. Data represent mean ± SD of three separate experiments for each cell line.

 
At the level of light microscopy, AST- and TEF-treated cells exhibited morphological changes that are characteristic of apoptosis in the form of cell shrinkage and membrane blebbing (Figure 3B and C). Furthermore, PI staining showed that AST and TEF induce nuclear condensation and fragmentation characteristic of apoptosis. In accordance with the viability assay, TEF appeared to induce a more intense apoptotic effect than AST (Figure 3E and F). Since AST and TEF are specific H1 receptor antagonists, they were chosen to complete our study on A375 human melanoma cells.


Figure 3
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  		Fig. 3 Morphological changes characteristic of apoptosis induced by 10 µM AST and TEF during 8 h in A375 melanoma cells. (A–C) Morphological characteristics observed by phase contrast microscopy. (D–F) Nuclear changes observed by staining the nucleus with PI. (A and D) Untreated cells; (B and E) AST-treated cells; (C and F) TEF-treated cells. Images show the typical cellular changes associated with apoptosis: cell shrinkage and rounding, blebbing formation and nuclear condensation and fragmentation in response to the treatment. All fields were photographed at a magnification of x100.

 
The apoptosis induced by AST and TEF in human melanoma cells was further confirmed and quantified by flow cytometry. Cells treated with 10 µM AST or TEF for 2–8 h were tested for externalization of phosphatidylserine head groups to the cell surface, an event that occurs during the early stages of apoptosis (27). As can be seen in Figure 4, an 8 h treatment with 10 µM AST induced apoptosis in ~48% of cells. Of these, 25% were only annexin V positive indicating early apoptosis. Similar treatment with TEF induced apoptosis in ~74% of the cell population; of these apoptotic cells, 28% were at the early apoptosis stage. These results were further corroborated by detection of hypodiploid cells, characteristic of apoptosis, as cells with low DNA content located in the sub-G0/G1 phase of the cell cycle. TEF and AST induced time-dependent increase of the percentage of hypodiploid cells in A375 melanoma cells (Table I). Furthermore, the cell-cycle analysis indicated a transient arrest of the cells at S- and G2-M-phase of the cell cycle at the first 2 h of the treatment.


Figure 4
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  		Fig. 4 Flow cytometric analysis showing that AST and TEF induce apoptosis in A375 melanoma cells. Annexin V-FITC assay. Untreated and 10 µM AST- and TEF-treated cells were harvested following 8 h exposure and stained with annexin V-FITC and PI. The percentage of early apoptotic cells in the lower right quadrant (annexin V-FITC positive/PI negative cells), as well as late apoptotic cells located in the upper right quadrant (annexin V-FITC positive/PI negative cells) increased very significantly in treated cultures. Treatment with TEF induced more apoptosis (74%) than AST (48%) in A375 melanoma cells, in keeping with its lower IC50 value.

 

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  		Table I Cell-cycle distribution of AST- and TEF-treated A375 melanoma cells

 
Apoptosis induced by TEF and AST in melanoma cells is caspase-2-dependent
To study if caspases were involved in TEF-induced apoptosis, we measured their activity in A375 cells treated with 10 µM TEF. The time course of activation of the initiator caspases 2, 8 and 9, and the executive caspases 3 and 6, induced by TEF in A375 cells is shown in Figure 5A. TEF was found to activate caspases 2 and 3 after 2 h of treatment. In contrast, there was no apparent increase in the activation of caspases 8 and 9 at that time point. Time course analysis of caspase family activity revealed that caspase-2 was activated first, and after 4 h of TEF treatment, the activity of caspases 2, 3 and 6 increased dramatically. However, low levels of activated caspases 8 and 9 were detected. The activation levels of the studied initiator and executive caspases reached a maximal level at 6 h.


Figure 5
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  		Fig. 5 TEF induces caspase family activation. (A) Induction of apoptosis in A375 melanoma cells by TEF was accompanied by activation of caspases 2, 3, 6, 8 and 9. Cells were treated with 10 µM TEF for various periods of time. Total cell lysates were prepared according to the manufacturer's instruction. Assays were performed in triplicate in 96-well plates on the basis of the manufacturer's protocol using a microtiter plate reader at 405 nm. The activity of caspases was calculated as fold increase relative to controls (without treatments). Cell lysates at 2 h showed increased activity of caspases 2 and 3, as measured using specific substrates; after 4 and 6 h, peak activities of caspases 2, 3, 6, 8 and 9 were detected. At 8 h, the activity of all the studied caspases started to decline. Data represent mean ± SD of three determinations from three separate experiments. (B) Effect of a general caspase inhibitor (z-VAD-FMK), a caspase-2 inhibitor (z-VDVAD-FMK) and a caspase-8 inhibitor (z-IETD-FMK) on cell death induced by TEF. Cells were pretreated with or without caspase inhibitors for 1 h and then challenged with 10 µM TEF for 8 h, after which cell viability was assessed using the XTT viability assay. Data represent mean ± SD of three determinations from three separate experiments. *P-values < 0.05 (statistically significant); **P-values < 0.01 (statistically very significant).

 
To confirm the role of individual caspases in TEF-induced apoptosis in A375 cells and to verify whether this process is caspase-dependent or not, we performed a series of blocking experiments. We observed that pre-treatment of A375 cells with 10–100 µM of z-VAD-FMK (a pan-caspase inhibitor) and z-VDVAD-FMK (caspase-2 inhibitor) for 1 hour before addition of TEF at 10 µM for 8 h inhibited cell death in a dose-dependent manner. Figure 5B represents the effect of caspase inhibitors (100 µM) on apoptosis induced by TEF. We observed that both the caspase-2 inhibitor and the pan-caspase inhibitor blocked the effects of TEF almost completely. Furthermore, we have found that the specific caspase-2 inhibitor was able to abolish caspase-3 activation (Figure 6A). We have found ~80% caspase-3 positive cells in TEF-treated cells compared with 4% caspase-3 positive cells when cells were pre-incubated with caspase-2 inhibitor before TEF treatment. Similar results were obtained when A375 cells were treated with 10 µM AST and with the Hs294T melanoma cell line (data not shown). These results indicate that the H1 receptor antagonist AST and TEF kill human melanoma cells by caspase activation signaling, mainly involving caspase-2.


Figure 6
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  		Fig. 6 Involvement of caspase-2 and DNA damage in the apoptosis induced by TEF in A375 melanoma cells. (A) Specific caspase-2 inhibitor abolished caspase-3 activation by TEF treatment. Untreated and treated cells with 100 µM caspase-2 inhibitor for 1 h were incubated with 10 µM TEF. The cells were collected, and after fixation and permeabilization, they were incubated with polyclonal rabbit anticleaved caspase-3 antibody. The cells were rinsed and incubated with ALEXA goat anti-rabbit secondary antibody. After three rinses the cells were incubated with DAPI for 10 min to stain the nucleus. Lastly, the cells were viewed and analyzed with a confocal microscope. Bar = 20 µm. (B) TEF induces DNA damage in A375 human melanoma cells. Comet assay or single-cell gel electrophoresis. Images show time-dependent DNA damage in single A375 cells treated with 10 µM TEF. The DNA damage induced by TEF occurs upstream to caspase activation. All fields were photographed at a magnification of x50.

 
TEF induces DNA damage in A375 human melanoma cells
The induction of DNA damage by TEF was investigated using the alkaline single-cell gel electrophoresis (SCGE) or ‘comet assay’, which provides a measure of both single- and double-strand breaks in DNA. Following treatment with 10 µM TEF, DNA damage was observed in A375 human melanoma cell line. The significant DNA damage was observed after 1 h of treatment and it was time-dependent. Furthermore, we have found that DNA damage occurs upstream to the caspase cascade activation since we have observed that pre-treatment of the cells with pan-caspase and specific caspase-2 inhibitors could not prevent TEF-induced DNA damage (Figure 6B)

TEF modifies mitochondrial function
The involvement of caspases 2 and 9 in the TEF-induced apoptotic death of melanoma cells suggested that TEF treatment may act by disrupting the mitochondrial membrane. We therefore employed DiOC6(3), a fluorescent dye that localizes to intact mitochondria and which has been widely used in recent years for studies of mitochondrial function (28), to evaluate the effect of TEF on A375 cells. A 2 h exposure to TEF (10 µM) was found to induce hyperpolarization of the mitochondrial membrane reflected by an increase in fluorescence intensity (Figure 7A). However, after 3 and 4 h of treatment, fluorescence intensity decreased significantly, suggesting that TEF treatment resulted in dissipation of {Delta}{Psi}m, pointing to loss of mitochondrial membrane integrity.


Figure 7
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  		Fig. 7 TEF induces mitochondrial membrane damage as indicated by mitochondrial membrane potential collapse and release of Cyt c to the cytosol. (A) Cells were incubated with 10 µM TEF for the indicated times and then stained with DiOC6(3) followed by flow cytometry evaluation. (B) Cyt c release from the mitochondria into the cytosol in A375 melanoma cells induced by TEF. Untreated and 10 µM TEF-treated cells for the indicated times were harvested and cytosolic and mitochondrial fractions were extracted. Forty micrograms of proteins per lane were separated by SDS–PAGE and analyzed by immunoblotting. A significant increase in the levels of cytosolic Cyt c can be seen in cells in response to TEF, accompanied by a parallel decrease in the levels of mitochondrial Cyt c.

 
We next measured mitochondrial release of Cyt c to the cytosol in response to TEF treatment. Analysis by western blotting revealed an increase in Cyt c in the cytosolic protein fraction and a concomitant decrease in the mitochondrial fraction 2 h after treatment with TEF (Figure 7B). This dual effect increased up to 6 h of treatment. Taken together, these data indicate that mitochondria are involved in TEF-induced apoptosis in melanoma cells.


   Discussion
Top
 Abstract
 Introduction
 Material and methods
 Results
 Discussion
References
 
Almost all available anticancer treatments induce cytotoxic effects by means of apoptosis. Since most of the currently available agents affect all rapidly proliferating cells, whether they are normal or malignant, they may have non-specific adverse side-effects (29). Hence, new effective anticancer agents with less non-specific toxic effects are currently being intensively searched for. In the present study, we show that H1 histamine receptor antagonists, which are frequently used in current clinical practice for the treatment of allergic diseases, are selectively cytotoxic for a variety of human malignant melanoma cell lines, and do not affect the viability of normal human melanocytes or embryonic murine fibroblasts.

Numerous in vitro and in vivo studies have been performed to explore the effects of histamine and its receptor antagonists on cancer cell proliferation, but a large deal of contradictory findings have been reported previously (3,4,30–34). Recently, histamine has been demonstrated to play an important role in the proliferation of human melanoma cells and represents a newly recognized member of the growth factors regulating the growth of melanoma cells by acting on H1 plasma membrane receptors (6,34). However, the cytotoxic effect of histamine and antihistaminics and their relation to apoptosis in cancer cells have not been clarified, particularly in melanoma cells. In a previous report, we demonstrated that DPH induces apoptosis in human T-cell acute lymphocytic leukemia cells (23). Other groups have demonstrated that TEF triggers apoptosis in cultures of thymocytes (35), human hepatoma cells, human colon adenocarcinoma cells and human myeloid leukemia cells (36,37). Here, we demonstrate for the first time that H1 receptor antagonists are cytotoxic for various human malignant melanoma cells through the induction of apoptosis.

The apoptotic effect of H1 receptor antagonists was demonstrated by studying the morphological and biochemical changes typical of apoptosis (9,10,38,39). Importantly, we have found that H1 receptor antagonists did not affect the growth and survival of normal melanocytes and embryonic murine fibroblasts at the same doses and duration of exposure. The lack of effect on normal cells may be due to the absence of high levels of histamine and histidine decarboxylase in normal melanocytes; such elevated levels have been reported in melanoma cells (5–7). In addition, normal melanocytes do not predominantly express the H1 receptor (6,40). Furthermore, H1 receptor antagonists may mainly affect rapidly proliferating cells, since we found that all of the studied agents induce very weak effect on MJOI human melanoma cells, which we observed to exhibit a very slow growth rate in comparison with the other tested melanoma cells.

Apoptosis induced by different stimuli leads to the activation of various members of the caspase family (13,14). In this study, we demonstrated that caspases 2, 3, 6, 8 and 9 were activated during apoptosis induced by various concentrations of TEF. Apoptosis seems to be caspase-dependent, since a general caspase inhibitor protected the cells nearly completely from the cytotoxic effects of TEF. Furthermore, caspase-2 appears to be a key factor in the cascade of caspase activation and in TEF-induced apoptosis, since it was the first to be activated. In addition, a specific caspase-2 inhibitor was able to protect cells almost completely from the cytotoxic effects of TEF and also to inhibit caspase-3 activation. Although the exact mechanism of caspase-2 activation is not clear yet, it has recently been found that caspase-2 can be activated in different cells by various stimuli causing DNA damage (19,20). Interestingly, in accordance with these reports, in our model, we have found for the first time that TEF induces time-dependent DNA damage before activation of caspase-2. Furthermore, we have found that pre-treatment of the cells with pan-caspase and specific caspase-2 inhibitors were not able to prevent DNA damage, indicating that caspase cascade activation occurs downstream to the TEF-induced DNA damage. In agreement with our results, other groups have reported that methapyrilene, another H1 histamine antagonist, induces genotoxic effect and DNA damage in hepatic cells (41). It has been demonstrated that DNA damage may occur as a consequence of a modulation of the intracellular calcium level (42). In addition, histamine has been reported to play an important role in intracellular calcium homeostasis through the activation of H1 histamine receptors (43). We think that, in our study, TEF could induce DNA damage via modulation of calcium level in the cells.

On the other hand, we have found a transient accumulation of the treated cells at S-phase and G2-M-phase of the cell cycle before passing to apoptosis. Previous studies have shown that cell-cycle arrest may occur as a result of DNA damage that makes the cells to be arrested at G2 checkpoint in order to allow DNA repair and to prevent entry into mitosis in the presence of damaged DNA (44–46). p53 protein induces apoptosis in response to various kinds of DNA damage in normal cells, while it is not necessary in tumor cells. Arita et al. (44) have demonstrated in colon cancer cell lines that apoptotic induction by DNA damage is not necessarily related to p53 status, and that induction of p53-independent apoptosis following DNA damage may correlate with G2-M arrest in the cell cycle.

In contrast to the important role of caspase-2 in TEF-induced apoptosis, caspase-8 appears to play a minor role. In our study, activation of caspase-8 was followed the activation of caspase-3 and 6, corroborating the findings of other studies (47,48). In addition, a specific caspase-8 inhibitor did not exert any significant effect on TEF-induced apoptosis. Furthermore, we did not find any changes in Fas expression or in Fas ligand detection in TEF-treated A375 cells compared with untreated control cells; indeed a Fas antibody failed to induce apoptosis in these cells (data not shown). Other groups have also demonstrated that melanoma cells are resistant to Fas-induced apoptosis (49). In keeping with our results, various recent studies have proposed that caspase-8 can be activated independently of death receptors (50,51).

One of the important denominators of apoptosis in many systems is mitochondrial dysfunction, characterized by loss of mitochondrial transmembrane potential and opening of mitochondrial permeability transition pores (52). Many apoptotic stimuli can lead to mitochondrial dysfunction either directly or indirectly (53). Caspase-2 activation is one of the mechanisms by which apoptotic stimuli may affect mitochondrial function (54). Once mitochondria are affected, they release various important proteins to the cytosol to terminate the apoptotic process. One of the most studied of such molecules is Cyt c, which is responsible for the activation of caspase-9. The finding that caspases 2 and 9 are activated in TEF-treated A375 melanoma cells is indicative of the involvement of mitochondrial pathway. Thus, we studied the effect of TEF on mitochondrial function in A375 melanoma cells by treating them with DiOC6 and analyzing the mitochondrial membrane potential. Interestingly, we found that TEF first caused hyperpolarization of the mitochondrial membrane, which is recorded to be one of the earlier apoptotic events (55), followed by collapse of the mitochondrial membrane potential. Furthermore, we found a release of Cyt c from the mitochondria to the cytosol. These results indicate the involvement of the mitochondrial pathway in TEF-induced apoptosis in melanoma cells. It has been demonstrated that the release of Cyt c from the mitochondria to the cytosol is well controlled by Bcl-2 family proteins (56). Indeed, in previous studies we have demonstrated that apoptosis induced by DPH, an H1 histamine antagonist, in leukemic cells can be abolished by Bcl-2 overexpression in the cells (23).

Taken together, the present results indicate that caspase-2 may regulate TEF-induced apoptosis by altering mitochondrial membrane integrity, the release of Cyt c and caspase-9 activation. This latter finding is consistent with a recent report that nuclear pro-caspase-2 is cleaved to produce active caspase-2 in advance of Cyt c release from mitochondria (21). Thus, caspase-2 activation may represent another major class of apoptotic pathways distinct from that mediated by caspases 8 and 9.

In summary, we conclude that H1 receptor antagonists induce apoptotic cell death in various human melanoma cell lines but not in normal human melanocytes or mouse embryonic fibroblasts. The corresponding process of apoptosis is related with DNA damage and caspase-2 activation and proceeds via the mitochondrial pathway. These findings may have important therapeutic implications in the future treatment of human melanoma.


   Acknowledgments
 
The authors are grateful to B. Zhivotovsky for critical reading of the manuscript and would like to express their thanks to I. Bernales-Pujana and A. Alvarez for invaluable assistance with cytometric analysis and to the General Service of Analytical and High Resolution Microscopy of the University of the Basque Country. This study was supported by grants from the Health Department of the Government of the Basque Country, the FIS of the Spanish Government (C03/10, nodo 11) and the University of the Basque Country (UPV 00075.327-14466/2002). S.-M.J. received a research fellowship from the Basque Government and B.O. was supported by a research fellowship from the Gangoiti Foundation.

Conflict of Interest Statement: None declared.


   References
Top
 Abstract
 Introduction
 Material and methods
 Results
 Discussion
 References
 

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Received October 12, 2005; revised March 11, 2006; accepted March 17, 2006.


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