The UKF-NB-4 cell line, established from bone marrow metastases of recurrent high-risk neuroblastoma, was a gift of Professor J. Cinatl Jr (J.W. Goethe University, Frankfurt, Germany). Valproic acid sodium salt (VPA), valpromide (VPM), trichostatin A (TSA) and etoposide (ETO) were purchased from Sigma Chemical Co. (St. Louis, MO, USA). Vincristine (VCR) sulfate was from Teva Pharmaceuticals, Prague, Czech Republic and cisplatin was Platadiam 50 from PLIVA-Lachema, Brno, Czech Republic. All other chemicals used in experiments were of analytical purity or better. Cells were grown at 37°C and 5% CO₂, cultivated in Iscove’s modified Dulbecco’s medium (IMDM) with 10% fetal bovine serum (both Life Technologies, Carlsbad, CA, USA). The cells were cultivated for at least 48 h with tested drugs, because this time period essentially corresponds to the time for two cycles of cell division. Moreover, such a time period is sufficient for the drugs used in this study to enter the tested cells, affect cell cycle and trigger apoptosis (17–20,33,34).

For detection of apoptosis, Annexin V-FITC Apoptosis Detection kit (Biovision, Milpitas, CA, USA) was used according to the manufacturer’s instructions and samples were analyzed using flow cytometry (LSR II, BD, Franklin Lakes, CA, USA). Briefly, 0.8×10⁶ UKF-NB-4 cells were plated in 60-mm dishes and treated with individual drugs (VPA dissolved in an IMDM medium), TSA and VPM (dissolved in dimethyl sulfoxide, DMSO), water solution of cisplatin prepared according to the manufacturer’s instructions (PLIVA-Lachema, Brno, Czech Republic), etoposide (dissolved in a small volume of DMSO; final volume of DMSO did not exceed 0.5%), vincristine sulfate (Teva Czech Industries, Prague, Czech Republic) or their combinations. After exposure to these compounds for 48 h, cells were washed with cold PBS, trypsinized and collected by centrifugation; cells were further re-suspended in 100 μl of Annexin binding buffer containing 5 μl of FITC Annexin V and 5 μl of PI. Then cells were gently vortexed and incubated for 20 min at room temperature in the dark. Binding buffer (400 μl) was added to each tube and centrifuged. Pellets of cells re-suspended in this buffer were then collected and measured using an LSR II flow cytometer (BD) and analyzed with FlowLogic software.

To detect cells with active caspase-3, 0.8×10⁶ neuroblastoma UKF-NB-4 cells were plated in 60-mm dishes and treated with individual drugs or their combinations for 48 h. Percentage of active caspase-3-positive cells was detected using the CaspGLOW™ Flourescein Active Caspase-3 Staining kit (eBioscience, San Diego, CA, USA). The procedure used was as described in the manufacturer’s instructions and samples were analyzed using flow cytometry (LSR II, BD, Franklin Lakes, CA, USA). Briefly, after exposure to the compounds, control or treated cells were washed with cold PBS, trypsinized and collected by centrifugation. Cell pellets were washed with PBS and after spinning re-suspended in 300 μl of a complete medium containing 1 μl of FITC labeled DEVD-FMK. Then the cells were incubated for 30 min at 37°C and 5% CO₂. Cells were collected by spinning and pellet was washed with wash buffer (according to the manufacturer’s instructions). Re-suspended washed cells were measured using the LSR II flow cytometer (BD, Franklin Lakes, CA, USA) and analyzed with FlowLogic software.

The xCELLigence RTCA DP Instrument (ACEA Bioscience Inc., San Diego, CA, USA) placed in a humidified incubator at 37°C and 5% CO₂ was used for real-time label-free monitoring of cell-viability (37). UKF-NB-4 cell line (20,000 cells) were seeded into 16-well plates for impedance-based detection. Each condition (control, 1 mM VPA, 4 μM cisplatin and combination of 1 mM VPA with 4 μM cisplatin) was tested in duplicate. Cell index (CI) was monitored every 30 min for 140 h and data recorded by the supplied RTCA software.

To estimate an acetylation status of histones H3 and H4, 4×10⁶ cells were plated in 100-mm dishes and cultivated and treated with tested drugs. Namely, cells were treated with 1 mM VPA, 40 or 150 nM TSA, or 4 mM VPM for 8 h or with 20 μM cisplatin or 8 μM etoposide for 24 h. Cells were also treated with cisplatin and etoposide in a combination with 1 mM VPA in two different regimens: i) cells were incubated with cisplatin or etoposide for 24 h prior to incubation with VPA for 8 h and ii) cells were incubated with VPA for 8 h prior to incubation with cisplatin or etoposide for 24 h. Cells were harvested and histones were isolated from the cell pellets. The acid extraction followed by precipitation of histones using trichloracetic acid (TCA) method described by Shechter et al was used (35). The concentration of proteins was measured using a Lowry method (36) with DC Protein Assay (Bio-Rad, Hercules, CA, USA). Histones (5 μg) were electrophoretically separated using 4–20% TGX precast gels (100 mA). After migration, histones were transferred to a nitrocellulose membrane and incubated with 5% non-fat milk to block non-specific binding. The membranes were then exposed to specific rabbit polyclonal anti-acetyl-histone H3 (1:4,000) and anti-acetyl-histone H4 (1:1,000) antibodies (both from Upstate Biotechnology Inc., Lake Placid, NY, USA) overnight at 4°C. Membranes were washed and exposed to peroxidase-conjugated anti-IgG secondary antibodies (1:2,000) and the antigen-antibody complex was visualized by enhanced chemiluminescence detection system according to the manufacturer’s instructions (Immun-Star HRP Substrate, Bio-Rad), using X-ray film MEDIX XBU (Foma, Hradec Kralove, Czech Republic). The anti-histone H3 antibody (1:10,000; Millipore, San Diego, CA, USA) was used as a loading control.

To determine phosphorylation of histone H2AX, 0.8×10⁶ cells were plated in 60-mm dishes and treated with individual drugs or their combinations. After exposure to the compounds, control or treated cells were washed with cold PBS, trypsinized and collected by centrifugation. Cell pellets were washed with PBS and after spinning the cells were fixed in 2% formaldehyde in PBS for 10 min. After PBS washing, the cells were re-suspended in ice-cold 90% methanol in PBS and incubated for 60 min at −20°C. Then the cells were washed three times with wash buffer (PBS containing 0.5% BSA and 0.2% Triton X-100). Cells were incubated in 50 μl of wash buffer containing 5 μl of pH2AX antibody [Alexa Fluor^® 647 anti-H2A.X-Phosphorylated (Ser139), Biolegend, San Diego, CA, USA] for 60 min at 4°C. Cells were washed and measured using the LSR II (BD, Franklin Lakes, CA, USA) and analyzed with FlowLogic software.

Data are expressed as mean ± SD. Student’s t-test was used when comparing two conditions. P-value <0.05 was considered as statistically significant. GraphPad Prism6 software was used for this statistical and graphical processing of data. Two-one-sided-test (TOST) was used to test equivalence, XLSTAT software was used.

The first step of this study was to examine the apoptosis induced in UKF-NB-4 cells treated with individual drugs. As translocation of phosphatidylserine to the external membrane leaflet is one of the earliest features of apoptosis, early apoptotic cells can thus be identified by the Annexin V-FITC/PI double staining assay. Besides, double staining with PI allows differentiation of early apoptotic cells with intact membranes (Annexin V⁺/PI⁻) from late apoptotic/necrotic cells with leaky membranes (Annexin V⁺/PI⁺) and normal cells (Annexin V⁻/PI⁻). As a result, the different groups of these stained cells can be distinguished and quantified by flow cytometry.

As shown in Fig. 1, treatment of UKF-NB-4 neuroblastoma cells with a non-toxic concentration of VPA, 1 mM VPA, for 48 h does not induce apoptosis in these cells (the amount of Annexin V⁻/PI⁻ cells is almost the same as in the control sample, Fig. 1A and B). However, their treatment with cisplatin alone and mainly with this drug simultaneously with VPA resulted in induction of apoptosis (Figs. 1C and D and 2). The Annexin V-FITC/PI double staining assay showed that treatment of UKF-NB-4 cells with 20 μM cisplatin for 48 h, decreased the percentage of viable cells to 61.3% (shown in a lower-left panel in Fig. 1C) with a concomitant increase in percentage of early apoptotic cells to 15.5% (shown in a lower-right panel in Fig. 1C), and late apoptotic cells to 22.2% (shown in an upper-right panel in Fig. 1C). These results confirmed that cisplatin plays a critical role in triggering apoptotic cell death in cisplatin-treated UKF-NB-4 cells.

When cells were co-treated with cisplatin and VPA, a higher decrease in the percentage of viable cells than by treatment of cells with cisplatin alone was produced, ≤43.8% (Fig. 1D). This decrease was concomitant with an increase in amounts of early apoptotic cells, to 20.7% and mainly late apoptotic cells, to 34.2% (Fig. 1D). These data indicate that VPA elevates a potency of DNA-damaging agent cisplatin to induce apoptosis in UKF-NB-4 cells. Similar trend in induction of apoptosis in UKF-NB-4 cells was also seen after their treatment with 4 μM cisplatin (the dose exhibiting a low toxicity to the tested cells, Fig. 2) and by this drug combined with VPA (Fig. 2 for a decrease in the percentage of viable UKF-NB-4 cells caused by cell treatment with cisplatin and VPA).

Caspases, a family of cysteine-aspartic proteases, are known to be crucial mediators in the apoptotic-signaling pathways (38). Since caspase-3 is the major executioner caspase essential for activation of one of the primary apoptotic signaling pathways (39) and its activation ultimately leads to cell death (40), it is thus suited as a read-out in an apoptosis assay. Therefore, in further experiments we evaluated whether apoptosis induced by the above-mentioned treatment regimens is associated with changes in caspase-3 activation. A trend similar to that found using Annexin V, the induction of apoptosis caused by a combined effect of VPA and cisplatin was detected by measuring the percentage of cells with active caspase-3 (Fig. 3). Exposure of UKF-NB-4 cells to 1 mM VPA simultaneously with 4 or 20 μM cisplatin increased levels of cells with active caspase-3, by up to 2-times as compared with cells treated with cisplatin alone. These results indicate that apoptosis induced in the UKF-NB-4 neuroblastoma cell line by cisplatin was triggered by the activation of caspase-3 that is potentiated by co-treatment of the tested cell line with VPA.

An increase in a cytotoxic potency of 4 μM cisplatin by 1 mM VPA was also proved by analyzing cell growth with the xCELLigence system (Fig. 4). In this method the relative changes in electrical impedance (expressed as cell index) due to coverage of bottom well by cells, corresponds to cell viability (41). As shown in Fig. 4, the tested cells cultivated with VPA grow slowly up till ~ 90 h of cultivation, while their cell index was not increaed after this time period. UKF-NB-4 cells treated with cisplatin grow exponentially till 50 h of cultivation, but after this time period, cisplatin caused a decrease in their viability. However, after >100 h of cultivation, these cells incubated with 4 μM cisplatin started to grow again (Fig. 4). Hence, treatment of cells with 4 μM cisplatin alone is not sufficient to inhibit cell growth totally; after 90-h treatment, a portion of viable cells were able to grow during further cultivation. The most efficient cytotoxic effect on UKF-NB-4 cells was produced by their co-cultivation with VPA and cisplatin; when this combination was used, the value of cell index decreased close to zero (Fig. 4).

Computation analysis of cell survival calculated by Compusyn software (42,43) was used to estimate whether activities of these two drugs are synergistic. Namely, we calculated the value of the combined effect of VPA and cisplatin and expressed it as combination index (CoI). When the values of CoI are <0.90, activities of two drugs are synergistic (the combination index ranging from 0.70 to 0.89 corresponds to a moderate synergism of drugs). However, the values of ≥0.9 indicate that the activities of two drugs are not synergistic (42,43). The value of CoI for the simultaneous effect of 4 μM cisplatin and 1 mM VPA equals 0.70, which corresponds to a moderate synergism of this combination.

Because the effect of the combination of cytostatic drugs with HDAC inhibitors may depend on the treatment regimens, causing either an increase or decrease in cytotoxic effects (17,20,44), we also examined various treatment regimens for a combination of the tested drugs with HDAC inhibitors.

In further experiments, several combinations of treating the UKF-NB-4 cells with VPA and cisplatin were used (Table I). The apoptosis induction in cells cultivated under these treatment regimens is shown in Fig. 5. The most effective drug combination resulting in a decrease in viability of neuroblastoma cells was their exposure to cisplatin and VPA (the cells treated with 20 μM cisplatin for 24 h and then with 1 mM VPA for 48 h) (see a combination cisplatin/VPA in Fig. 5). On the contrary, the opposite sequence of drug application (VPA/cisplatin) led to essentially no changes in viability of cells as compared with cultivation of cells with cisplatin alone (see VPA/cisplatin versus 0/cisplatin regimens in Fig. 5). These results demonstrate that VPA can potentiate the toxic effects of cisplatin only if UKF-NB-4 cells are primarily affected by this DNA-damaging drug.

In contrast to VPA, no effects of TSA, another HDAC inhibitor tested in our study, at a non-toxic concentration for UKF-NB-4 (40 nM), on cytotoxicity of cisplatin to UKF-NB-4 cells was found using Annexin V-FITC/PI double staining assay or active caspase-3 assay (data not shown).

The effect of combined treatment of UKF-NB-4 neuroblastoma cells with VPA and etoposide was also investigated. Treatment of these cells with 8 μM etoposide, and predominantly with this drug together with 1 mM VPA, resulted in induction of apoptosis (Fig. 1E and F). Treatment of UKF-NB-4 cells with etoposide for 48 h induced a decrease in the percentage of viable cells to 76.2% (shown in a lower-left panel in Fig. 1E) with a concomitant increase in percentage of early apoptotic cells, to 12.0% (shown in a lower-right, panel in Fig. 1E) and late apoptotic cells, to 10.9% (shown in an upper-right panel in Fig. 1). These findings confirmed the results found in our previous study (20) that etoposide induces apoptotic cell death in UKF-NB-4 cells.

Analogously to exposure of the tested neuroblastoma cells to cisplatin combined with VPA, a 1.7-fold decrease in cell viability (expressed as percentage of Annexin V⁻/PI⁻ cells) was generated in these cells by their exposure to VPA combined with etoposide as compared to their exposure to etoposide alone (Fig. 1E and F). This decrease was parallel with an increase in amounts of early apoptotic cells, to 17.9%, and predominantly late apoptotic cells, to 35.8% (Fig. 1F). These data indicate that VPA also elevates the potency of the second tested DNA-damaging drug, etoposide, to induce apoptosis in UKF-NB-4 cells.

Using another method testing the effect of combined treatment of UKF-NB-4 cells, measuring the activation of caspase-3, an increase in the etoposide effect by VPA was also found. Exposure of UKF-NB-4 cells to VPA simultaneously with etoposide increased an amount of cells with the active caspase-3, by ≤1.7-times (Fig. 6). All these results indicate the enhanced effect of the combination of these drugs on induction of apoptosis mediated by caspase-3 activation.

Because several schedules of combined treatment (treatment regimens) of the UKF-NB-4 neuroblastoma cell line with VPA and etoposide have already been carried out in our previous study (20), such experiments were not performed in this study. It should be emphasized that the most effective treatment leading to the highest decrease in viability of neuroblastoma cells was the combined exposure of cells to etoposide followed by their treatment with VPA (cells treated with 8 μM etoposide for 24 h and then with 1 mM VPA for 48 h) (20). Hence, the regimen analogous to that found to be most efficient in treatment of cells with cisplatin combined with VPA. These results demonstrate that VPA can exhibit the potentiation effect only if UKF-NB-4 neuroblastoma cells are primarily influenced either by cisplatin or by etoposide.

Computation analysis (42,43) was again used to estimate whether activities of VPA with etoposide are synergistic. The value of CoI for the simultaneous effect of 8 μM etoposide and 1 mM VPA equals 0.52 that corresponds to a synergism of these drugs.

When TSA at its non-toxic concentration (40 nM) was used, cytotoxicity of etoposide to UKF-NB-4 cells was essentially not influenced by this HDAC inhibitor (data not shown).

In further experiments, the influence of VPA and TSA on cytotoxicity of vincristine, the anticancer drug acting by a mechanism different from that of DNA damage, was investigated. Vincristine is known to induce cell death in tumor cells including neuroblastoma cells by inhibiting the assembly of microtubule structures and disrupting mitosis in the metaphase (45,46).

As shown in Fig. 1, treatment of UKF-NB-4 neuroblastoma cells with 20 nM vincristine led to induction of apoptosis in these cells. The Annexin V-FITC/PI double staining assay indicated that treatment of UKF-NB-4 cells with vincristine for 48 h induced a decrease in the percentage of viable cells to 72.3% (shown in a lower-left panel in Fig. 1G), with a concomitant increase in percentage of early apoptotic cells to 13.6% (shown in a lower-right panel in Fig. 1G), and late apoptotic cells to 12.5% (shown in an upper-right panel in Fig. 1G). These results confirmed that vincristine is an important drug inducing apoptotic cell death in vincristine-treated UKF-NB-4 cells. However, when the cells were exposed to vincristine combined with VPA, no further changes in cell viability as compared to UKF-NB-4 cells treated with vincristine were found (Fig. 1H). Likewise, in the case of measuring the activation of caspase-3 in the tested cells treated with vincristine simultaneously with VPA, no increase in a toxic potency of vincristine was found (Fig. 7). These findings demonstrate that the sensitizing effect of VPA on the tested cells is produced only when they are affected by the DNA-damaging drugs (cisplatin and etoposide) before their treatment with this HDAC inhibitor. When TSA at a 40 nM concentration was used, cytotoxicity of vincristine to UKF-NB-4 cells was also not influenced by this HDAC inhibitor (data not shown).

In additional experiments, we investigated cytotoxicity of valpromide (VPM), a derivative of VPA which is also used as an anti-epileptic drug, but does not exhibit HDAC inhibition activity (47), on UKF-NB-4 neuroblastoma cells. We evaluated the effect of this drug on induction of apoptosis (measured by Annexin V/PI labeling) in these cells. Treatment of UKF-NB-4 neuroblastoma cells with different concentrations of VPM up to concentrations of 4 mM did not induce apoptosis in these cells (data not shown). When UKF-NB-4 cells were treated with this non-toxic concentration of VPM simultaneously with 20 μM cisplatin, no increase in induction of apoptosis caused by cisplatin was produced by VPM (data not shown).

When activation of caspase-3 in cells treated with VPM together with cisplatin was determined, no increase in the potency of cisplatin to activate caspase-3 by VPM was found (Fig. 8). Exposure of UKF-NB-4 cells to 20 μM cisplatin combined with simultaneous treatment with 4 mM VPM does not increase amounts of cells with active caspase-3. These results indicate that the sensitizing effect of VPA on cisplatin toxicity in UKF-NB-4 neuroblastoma cells is related to its HDAC inhibition activity.

Acetylation of histones is an important epigenetic phenomenon that is dictated by HDAC activities. Therefore, here we investigated the changes in acetylation of core histones H3 and H4 in UKF-NB-4 cells treated with VPA and TSA, as well as with VPM. Cultivation of UKF-NB-4 cells with VPA and TSA at their non-toxic concentrations led to the different effects on acetylation of histones H3 and H4. Whereas 1 mM VPA increases acetylation of these histones in UKF-NB-4 cells, confirming its HDAC inhibitory efficiency, a negligible effect of 40 or 50 nM TSA on acetylation of these histones was found (Fig. 9). The concentrations of TSA of 40 or 50 nM are therefore insufficient to inhibit HDAC activities in UKF-NB-4 cells. However, when these cells were cultivated in a medium containing a higher (toxic) concentration of TSA (150 nM), a pronounced increase in acetylation of histones H3 and H4 was detected. In contrast to these results, no increase in acetylation of these histones was caused by VPM (Fig. 9). This confirms the absence of HDAC inhibitory effects of this derivative of VPA.

When UKF-NB-4 neuroblastoma cells were treated with cisplatin or etoposide, a low decrease in acetylation of histone H3 and essentially no effect on acetylation of histone H4 were found (Fig. 10). In further experiments, two combination treatment regimens of the cells with VPA and cisplatin or VPA with etoposide were used to investigate the effects of combined treatment on the histones H3 and H4 acetylation status. Only the regimen, where UKF-NB-4 cells were pretreated with cisplatin or etoposide and then treated with VPA produced an increase in acetylation of histones H3 and H4 (Fig. 10). This increase in histone acetylation paralleled the potentiation of toxic effects of cisplatin (Fig. 5) or etoposide (20). Hence, only if UKF-NB-4 cells were primarily affected by the tested DNA-damaging drugs prior to cultivation with VPA, this HDAC inhibitor increases acetylation of histones (Fig. 10) and potentiated cytotoxicity of both DNA-damaging drugs on UKF-NB-4 cells (Fig. 1).

Etoposide is known to be the DNA-damaging drug that acts, beside its intercalation into DNA, as an inhibitor of topoisomerase II activity that leads to formation of double-strand breaks in DNA (26–28). Therefore, we examined the effect of VPA on this mechanism of etoposide-mediated cytotoxic action. Phosphorylation of histone H2A on serine 139, termed γH2AX (pH2AX), by kinases sensing the double-strand DNA break is a sensitive marker of this type of DNA damage (48–51). Therefore, the levels of pH2AX were determined to analyze whether treatment of cells with etoposide is influenced by VPA. After 48-h cultivation of cells in a medium containing 1 mM VPA, 4 μM etoposide or their combination, the levels of pH2AX were examined by flow cytometry. Representative histograms of pH2AX fluorescence found in these experiments are shown in Fig. 11. Whereas essentially no increase in pH2AX was induced by VPA (Fig. 11B), a pronounced enhancement in pH2AX was caused by cell exposure to etoposide (Fig. 11C). Co-cultivation of UKF-NB-4 cells with etoposide and VPA (Fig. 11D) did not change percentage of pH2AX-positive cells as compared with cultivation of UKF-NB-4 cell with etoposide alone (Fig. 11C and D).