In this study, we investigated the influence of platelet-activating factor (PAF) on the induction of apoptosis-regulating factors in B16F10 melanoma cells. PAF increased the expression of mRNA and the protein synthesis of antiapoptotic factors, such as Bcl-2 and Bcl-xL, but did not increase the expression of the proapoptotic factor, Bax. A selective nuclear factor-κB (NF-κB) inhibitor, parthenolide, inhibited the effects of PAF. Furthermore, PAF inhibited etoposide-induced increases in caspase-3, caspase-8, and caspase-9 activities, as well as cell death. p50/p65 heterodimer increased the mRNA expression of Bcl-2 and Bcl-xL and decreased etoposide-induced caspase activities and cell death. In an in vivo model in which Matrigel was injected s.c., PAF augmented the growth of B16F10 cells and attenuated etoposide-induced inhibition of B16F10 cells growth. These data indicate that PAF induces up-regulation of antiapoptotic factors in a NF-κB-dependent manner in a melanoma cell line, therefore suggesting that PAF may diminish the cytotoxic effect of chemotherapeutic agents. (Cancer Res 2006; 66(9): 4681-6)

Apoptosis (i.e., programmed cell death) plays an important role in a wide variety of physiologic processes. When apoptosis is dysregulated, it can contribute to the pathogenesis of many diseases, including cancer, autoimmunity, and neurodegenerative disorders (1, 2). The role of apoptosis in cancer and the possible exploitation of this role for prognostic and therapeutic purposes are extensively discussed in the literature. Numerous studies have shown a positive correlation between the apoptotic index and malignancy grade (37).

Apoptosis is controlled by regulating factors, such as p53, Bcl-2, and Bax. The Bcl-2 family, in particular, regulates apoptosis induced by a wide variety of stimuli (79). Some proteins within this family, including Bcl-2 and Bcl-xL, are apoptosis-inhibitory proteins; others, such as Bax, Bad, and Bak, are promoters of apoptosis.

Platelet-activating factor (PAF), which is produced by a variety of inflammatory cells, is a potent lipid messenger involved in cellular activation, intracellular signaling, apoptosis, and diverse inflammatory reactions (1012). In addition to this important role, PAF plays a role in apoptosis in various cell types (1316), and PAF activates signal transduction pathways similar to the antiapoptotic effects of growth factors and cytokines (17, 18). Notably, PAF has also been detected in tumor cells and its effect in tumor development has recently been investigated (1921). This information suggests the possibility that PAF may affect tumor growth via the regulation of apoptosis-regulating factors.

In this study, we have investigated the role of PAF in the expression of apoptosis-regulatory proteins in B16F10 melanoma cells. We have found that PAF increased the expression of mRNA and protein synthesis of antiapoptotic factors via nuclear factor-κB (NF-κB) activation. PAF also seemed to eliminate etoposide-induced apoptosis.

Animals. Specific pathogen-free female C57BL/6 mice were obtained from the Korean Institute of Chemistry Technology (Daejeon, Republic of Korea) and were kept in our animal facility for at least 2 weeks before use. All mice were used at 8 to 10 weeks of age.

Reagents. PAF (1-O-alkyl-2-acetyl-sn-glyceryl-3-phosphorylcholine), etoposide, and caspase substrates (Ac-DEVD-pNA, Ac-IETD-pNA, and Ac-LEHD-pNA) were purchased from Sigma Chemical Co. (St. Louis, MO). The PAF receptor antagonist, WEB 2170, was a donation from Dr. C.K. Rhee (College of Medicine, Dankook University, Republic of Korea). The NF-κB inhibitor, parthenolide, was purchased from BIOMOL Research Laboratories, Inc. (Plymouth Meeting, PA). Matrigel, an extract of murine basement membrane proteins, consisting predominantly of laminin, collagen IV, heparin sulfate proteoglycans, and nidogen/entactin, was purchased from Collaborative Research, Inc. (Bedford, MA). Antibodies against Bcl-2, Bcl-xL, Bax, and PAFR were purchased from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA). Anti-β-actin antibody was from Sigma Chemical.

Cell culture. The B16F10 mouse melanoma, which is metastatic in the lungs of C57BL/6 mice, was originally supplied by the Tumor Repository of the National Cancer Institute (Bethesda, MD), and was maintained in RPMI 1640 (Life Technologies, Grand Island, NY) supplemented with 10% fetal bovine serum (Cambrex Co., Walkersville, MD).

Real-time reverse transcription-PCR. RNA was prepared as previously described (22). Reverse transcription was done using 0.1 μg total RNA in a 10 μL reaction mixture (Promega, Madison, WI) containing oligo(dT) and avian myeloblastosis virus reverse transcriptase. PCR was done on the Rotor-Gene 3000 System (Corbett Research, Morklake, Australia) using the SYBR Green PCR Master Mix Reagent kit (Qiagen, Valencia, CA). The primers used for real-time PCR are as follows: Bcl-2, 5′-TTCGCAGACATGTCCAGTCAGCT-3′ and 5′-TGAAGAGTTCCTCCACCACCGT-3′; Bcl-xL, 5′-AATGAACTCTTTCGGGATGGAG-3′ and 5′-CCAACTTGCAATCCGAC TCA-3′; Bax, 5′-ATGCGTCCACCAAGAAGCTGA-3′ and 5′-AGCAATCATCCT CTGCAGCTCC-3′; and β-actin, 5′-CTGAAGTCACCCATTGAACATGGC-3′ and 5′-CAGAGCAGTAATCTCCTTCTGCAT-3′. The relative levels of mRNA were calculated using the standard curve generated from cDNA dilutions. The mean cycle threshold (Ct) values from quadruplicate measurements were used to calculate the gene expression, with normalization to β-actin as an internal control. Calculations of the relative level of gene expression were conducted according to the complementary computer software (Corbett Research) using a standard curve.

Immunoblotting analysis. Cell lysates were prepared in radioimmunoprecipitation assay buffer [0.1% SDS, 1% IGEPAL, 0.5% sodium deoxycholate, and 1 mmol/L phenylmethylsulfonyl fluoride (PMSF)], and equal amounts of the lysates were separated on 10% SDS polyacrylamide gel under reducing conditions. The lysates were then transferred onto Protran nitrocellulose membranes (Schleicher & Schuell, Keene, NH). Equal protein loading was verified by staining both the gel and the membrane with Coomassie brilliant blue R-250 and Ponceau S (Sigma Chemical), respectively. Membranes were blocked for 1 hour at room temperature in 5% skim milk in TBS, followed by 1 hour of incubation with primary antibodies. Blots were washed for 15 minutes with three changes of TBS-0.05% Tween 20 solution, followed by incubation for 1 hour at room temperature with the alkaline phosphatase–conjugated anti-rabbit IgG antibody (Santa Cruz Biotechnology). Alkaline phosphatase activity was detected using an nitroblue tetrazolium/5-bromo-4-chloro-3-indolyl phosphate color development system (Promega).

Electrophoretic mobility shift assay. Nuclear extracts were prepared from the cells as previously described (22). To inhibit endogenous protease activity, 1 mmol/L PMSF was added. An oligonucleotide containing the Igκ-chain-binding site (κB, 5′-CCGGTTAACAGAGGGGGCTTTCCGAG-3′) was synthesized as a probe for the gel retardation assay. The two complementary strands were annealed and labeled with [α-32P]dCTP. Labeled oligonucleotides (10,000 cpm), 12 μg nuclear extracts, and binding buffer [10 mmol/L Tris-HCl (pH 7.6), 500 mmol/L KCl, 10 mmol/L EDTA, 50% glycerol, 100 ng poly(deoxyinosinic-deoxycytidylic acid), and 1 mmol/L DTT] were incubated for 30 minutes at room temperature in a final volume of 20 μL. For the supershift/inhibition assay, 1 μg specific supershifting anti-p50, anti-p65, or control rabbit antibody (Santa Cruz Biotechnology) was incubated with the nuclear extract on ice for 1 hour before the addition of labeled oligonucleotide to the binding reaction. The reaction mixture was analyzed by electrophoresis on a 5% polyacrylamide gel in 0.5× Tris-borate buffer. Specific binding was controlled by competition with a 50-fold excess of cold κB or cyclic AMP response element (CRE) oligonucleotide.

Analysis of apoptosis. For determination of DNA fragmentation, we used the Cell Death Detection ELISA kit (Roche Diagnostics Corporation, Indianapolis, IN) according to the instructions of manufacturer, and DNA was also prepared in lysis buffer containing 10 mmol/L Tris-HCl (pH 7.4), 10 mmol/L EDTA (pH 8.0), and 0.5% Triton X-100. The DNA was then analyzed on a 2% agarose gel.

We used the specific colorimetric substrates for the determination of caspase-3, caspase-8, and caspase-9 activities. Cells were harvested in lysis buffer containing 50 mmol/L Tris-HCl (pH 7.4), 1 mmol/L EDTA, 10 mmol/L EGTA, and 10 μmol/L digitonin, and the lysates were incubated with each substrate for 2 hours at 37°C. The lysates were then read at a wavelength of 405 nm.

Transfection of cDNA encoding p65, p50, or IκBα cloned in pcDNA3. Expression vectors for p50, p65, and IκBα (donations from Dr. J.W. Lee, Baylor College of Medicine, Houston, TX) were prepared using commercial kits (Qiagen, Chatsworth, CA). Cells were cultured to 60% to 80% confluence in 60 mm plates. For each plate, each plasmid DNA and 1 μL LipofectAMINE reagent (Life Technologies, Inc., Gaithersburg, MD) were separately diluted in 100 μL serum-free medium, mixed together, and incubated at room temperature for 1 hour. Plates were then washed with serum-free medium, 200 μL serum-free medium was added, and the diluted solution was added to the cells. Plates were incubated at 37°C for 8 hours, after which a growth medium containing 10% serum was added. After 18 hours of additional incubation, cells were then left untreated or were treated with 50 μmol/L etoposide. All experiments were repeated at least thrice with two different batches of purified DNA.

Evaluation of antiapoptotic response in vivo. Tumors were established by injecting C57BL/6 mice, s.c. in the right flank with B16F10 cells (5 × 105 per plug) mixed in Matrigel (23). Tumor growth was assessed at least thrice a week by three-dimensional measurement of tumor volume using a previously described vernier caliper technique (24).

For determination of apoptosis in situ, we used the ApopTag Plus Peroxidase In situ Apoptosis Detection kit (Chemicon, Temecula, CA). Tumor plugs were removed on day 3 and were then fixed in 10% neutral buffered formalin (Sigma Chemical) overnight at 4°C, paraffin embedded, and sectioned at 4 μm. Samples were treated with 20 μg/mL proteinase K for 15 minutes, quenched with 3% H2O2 for 5 minutes at room temperature, and subsequently mixed with digoxigenin-labeled dUTP in the presence of terminal deoxynucleotidyl transferase followed by peroxidase-conjugated antidigoxigenin antibody. Nuclear staining in apoptotic cells was detected using 3,3′-diaminobenzidine substrate. Samples were counterstained with hematoxylin. All magnifications are ×400.

Statistical analysis. Data are represented as the mean ± SE. Statistical significance was determined by the Student's t test when two data sets were analyzed or, alternatively, by ANOVA followed by the appropriate post hoc test for multiple data sets with StatView (version 4.5). All experiments were conducted at least twice. Reproducible results were obtained and representative data are, therefore, shown in the figures.

PAF induces the expression of antiapoptotic factors through NF-κB activation. The role of PAF in the expression of apoptotic factor genes and in protein synthesis in B16F10 melanoma was examined. PAF increased the expression of mRNA (Fig. 1A) and protein synthesis (Fig. 1B) of antiapoptotic factors, such as Bcl-2 and Bcl-xL, but did not increase mRNA expression or protein synthesis of proapoptotic factor, Bax. The mRNA and protein levels of antiapoptotic factors peaked at 4 and 16 hours, respectively. To clarify the relevance of PAF in B16F10 melanoma cells, we did immunostaining and immunoblotting analysis using anti-PAFR antibody. Cells were strongly stained with anti-PAFR antibody, but not control antibody, and immunoblotting analysis of whole-cell extracts revealed two bands: a weak band of ∼39 kDa and a strong band of ∼69 kDa, indicating that B16F10 melanoma cells express PAFR (data not shown).

Figure 1.

PAF induces the expression of antiapoptotic factors in B16F10 melanoma cells. Cells (1 × 106) were pretreated with a PAF antagonist, WEB 2170 (WEB), 30 minutes before PAF (3 μmol/L) treatment. A, RNA was prepared 4 hours after PAF treatment and real-time reverse transcription-PCR (RT-PCR) was done as described in Materials and Methods. Columns, mean; bars, SE. *, P < 0.0001 compared with the control group; **, P < 0.0001 compared with the PAF-treated group. B, proteins were prepared 16 hours after PAF treatment and assessed by immunoblotting analysis using anti-Bcl-2, anti-Bcl-xL, and anti-Bax antibody. To confirm equal protein loading, the same blot was stripped and reprobed with an anti-β-actin antibody.

Figure 1.

PAF induces the expression of antiapoptotic factors in B16F10 melanoma cells. Cells (1 × 106) were pretreated with a PAF antagonist, WEB 2170 (WEB), 30 minutes before PAF (3 μmol/L) treatment. A, RNA was prepared 4 hours after PAF treatment and real-time reverse transcription-PCR (RT-PCR) was done as described in Materials and Methods. Columns, mean; bars, SE. *, P < 0.0001 compared with the control group; **, P < 0.0001 compared with the PAF-treated group. B, proteins were prepared 16 hours after PAF treatment and assessed by immunoblotting analysis using anti-Bcl-2, anti-Bcl-xL, and anti-Bax antibody. To confirm equal protein loading, the same blot was stripped and reprobed with an anti-β-actin antibody.

Close modal

We investigated the association of NF-κB activity with the effects of PAF described in Fig. 1. B16F10 cells were treated with 3 μmol/L PAF for various time intervals. Treatment of PAF resulted in the activation of NF-κB, which was evident as early as 1 hour after treatment, reached a maximal level of binding at 2 hours, but gradually declined thereafter (Fig. 2A). Supershift assay revealed that the p50/p65 heterodimer is the activated form of the NF-κB induced by PAF (Fig. 2A). Treatment with either the PAF antagonist, WEB 2170, or the selective NF-κB inhibitor, parthenolide, 30 minutes before PAF treatment inhibited PAF-induced NF-κB activation (Fig. 2A). Parthenolide blocked the PAF-induced increase in mRNA expression (Fig. 2B) and protein synthesis (Fig. 2C) of antiapoptotic factors, but did not affect the expression of Bax (Fig. 2B and C).

Figure 2.

PAF induces the expression of antiapoptotic factors through NF-κB activation. A, B16F10 cells (1 × 106) were pretreated with a PAF antagonist, WEB 2170 (10 μg/mL), or NF-κB inhibitor, parthenolide (Parth, 5 μmol/L), 30 minutes before PAF (3 μmol/L) treatment. Nuclear extracts were prepared at the indicated times, incubated with α-32P-labeled κB or cAMP response element (CRE) oligonucleotide, and electrophoresed on a 5% polyacrylamide gel. Supershift/inhibition assay was conducted with 1 μg anti-p50, p65, or control rabbit antibody. B, real-time RT-PCR was done as described in Materials and Methods. Columns, mean; bars, SE. *, P < 0.0001 compared with the control group; **, P < 0.0001 compared with the PAF-treated group. C, proteins were prepared 16 hours after PAF treatment and assessed by immunoblotting analysis using anti-Bcl-2, anti-Bcl-xL, and anti-Bax antibody. β-actin was used as a loading control.

Figure 2.

PAF induces the expression of antiapoptotic factors through NF-κB activation. A, B16F10 cells (1 × 106) were pretreated with a PAF antagonist, WEB 2170 (10 μg/mL), or NF-κB inhibitor, parthenolide (Parth, 5 μmol/L), 30 minutes before PAF (3 μmol/L) treatment. Nuclear extracts were prepared at the indicated times, incubated with α-32P-labeled κB or cAMP response element (CRE) oligonucleotide, and electrophoresed on a 5% polyacrylamide gel. Supershift/inhibition assay was conducted with 1 μg anti-p50, p65, or control rabbit antibody. B, real-time RT-PCR was done as described in Materials and Methods. Columns, mean; bars, SE. *, P < 0.0001 compared with the control group; **, P < 0.0001 compared with the PAF-treated group. C, proteins were prepared 16 hours after PAF treatment and assessed by immunoblotting analysis using anti-Bcl-2, anti-Bcl-xL, and anti-Bax antibody. β-actin was used as a loading control.

Close modal

PAF inhibits etoposide-induced apoptosis. We next examined how PAF modulates antineoplastic drug-induced apoptosis. The topoisomerase II inhibitor, etoposide, is an antineoplastic drug that has been widely used to couple DNA damage to induce apoptosis (25). The kinetics of etoposide-induced apoptosis of B16F10 cells were examined using distinct markers of apoptosis (i.e., cell death and activation of caspase-3, caspase-8, and caspase-9). Etoposide significantly induced caspase activities and cell death at 6 and 16 hours, respectively (data not shown). To investigate the dose response of etoposide-induced apoptosis, B16F10 cells were treated with various doses of etoposide. Etoposide increased caspase-3, caspase-8, and caspase-9 activities (Fig. 3A) and cell death (Fig. 3B) in a dose-dependent manner. Pretreatment of the cells with PAF 4 hours (an optimum time point in preliminary experiments) before etoposide treatment resulted in inhibition of the etoposide-induced increases in caspase activities (Fig. 3A and D) and cell death (Fig. 3B and E). Additionally, the increased DNA fragmentation induced by etoposide was inhibited by PAF (Fig. 3C). Parthenolide eliminated the antiapoptotic effect of PAF (Fig. 3D and E), indicating the NF-κB dependency of the PAF effect.

Figure 3.

PAF inhibits etoposide-induced apoptosis in a NF-κB-dependent manner. To examine the effect of PAF in etoposide-induced apoptosis, B16F10 cells were pretreated with WEB 2170 (10 μg/mL) or parthenolide (5 μmol/L) 30 minutes before PAF (3 μmol/L) treatment for 4 hours. The cells were then treated with indicated concentrations (A and B) or 50 μmol/L (C-E) of etoposide (Etopo). A and D, 6 hours after etoposide treatment, caspase activities using the specific substrates were determined at 405 nm. B and E, apoptosis was detected with the Cell Death Detection ELISA kit 16 hours after etoposide treatment. Columns, mean; bars, SE. *, P < 0.0001 compared with the control group; **, P < 0.001 compared with the etoposide-treated group; ***, P < 0.001 compared with the PAF and etoposide-treated group. C, agarose gel analysis of DNA isolated from both the PAF and etoposide-treated cells.

Figure 3.

PAF inhibits etoposide-induced apoptosis in a NF-κB-dependent manner. To examine the effect of PAF in etoposide-induced apoptosis, B16F10 cells were pretreated with WEB 2170 (10 μg/mL) or parthenolide (5 μmol/L) 30 minutes before PAF (3 μmol/L) treatment for 4 hours. The cells were then treated with indicated concentrations (A and B) or 50 μmol/L (C-E) of etoposide (Etopo). A and D, 6 hours after etoposide treatment, caspase activities using the specific substrates were determined at 405 nm. B and E, apoptosis was detected with the Cell Death Detection ELISA kit 16 hours after etoposide treatment. Columns, mean; bars, SE. *, P < 0.0001 compared with the control group; **, P < 0.001 compared with the etoposide-treated group; ***, P < 0.001 compared with the PAF and etoposide-treated group. C, agarose gel analysis of DNA isolated from both the PAF and etoposide-treated cells.

Close modal

To establish a direct relationship between NF-κB and antiapopototic activity, the cells were treated with p50/p65 heterodimer. We then examined the effect of p50/p65 heterodimer on the mRNA expression of antiapopototic factors, and on etoposide-induced caspase activities and cell death. p50/p65 heterodimer increased the mRNA expression of Bcl-2 and Bcl-xL, but did not increase the mRNA expression of Bax (Fig. 4A). p50/p65 heterdimer also significantly decreased etoposide-induced caspase activities (Fig. 4B) and cell death (Fig. 4C). These data indicate that NF-κB plays a critical role in the regulation of antiapopototic factors.

Figure 4.

p50 and p65 subunits of NF-κB induce mRNA expression of antiapopototic factors and inhibit etoposide-induced apoptosis. B16F10 cells (5 × 105) were transfected with plasmid expressing p50/p65 (0.1/0.2 μg) and/or 0.2 μg IκBα expression vector and treated with etoposide (50 μmol/L) at 18 hours. A, real-time RT-PCR was done as described in Materials and Methods. Columns, mean; bars, SE. *, P < 0.0001 compared with the control group; **, P < 0.0001 compared with the p50/p65–transfected group. B, 6 hours after etoposide treatment, caspase activities using the specific substrates were determined at 405 nm. C, apoptosis was detected with the Cell Death Detection ELISA kit 16 hours after etoposide treatment. Columns, mean; bars, SE. *, P < 0.0001 compared with the control group; **, P < 0.001 compared with the etoposide-treated group; ***, P < 0.001 compared with the etoposide and p50/p65–treated group.

Figure 4.

p50 and p65 subunits of NF-κB induce mRNA expression of antiapopototic factors and inhibit etoposide-induced apoptosis. B16F10 cells (5 × 105) were transfected with plasmid expressing p50/p65 (0.1/0.2 μg) and/or 0.2 μg IκBα expression vector and treated with etoposide (50 μmol/L) at 18 hours. A, real-time RT-PCR was done as described in Materials and Methods. Columns, mean; bars, SE. *, P < 0.0001 compared with the control group; **, P < 0.0001 compared with the p50/p65–transfected group. B, 6 hours after etoposide treatment, caspase activities using the specific substrates were determined at 405 nm. C, apoptosis was detected with the Cell Death Detection ELISA kit 16 hours after etoposide treatment. Columns, mean; bars, SE. *, P < 0.0001 compared with the control group; **, P < 0.001 compared with the etoposide-treated group; ***, P < 0.001 compared with the etoposide and p50/p65–treated group.

Close modal

PAF inhibits apoptosis of B16F10 cells in vivo. We finally examined the in vivo effect of PAF on apoptosis of B16F10 cells in a murine model in which Matrigel was injected s.c. Tumor volume progressively increased over the observation period when Matrigel containing B16F10 cells was injected, and the simultaneous injection of PAF with the cells further increased the growth of B16F10 cells (Fig. 5A). As expected, etoposide caused a marked inhibition of the growth of B16F10 cells during the late stage of the observation period, and the inhibitory effect of etoposide was again almost completely blocked by PAF (Fig. 5A). The magnitude of apoptosis of the Matrigel section was compared using in situ internucleosomal DNA fragmentation by terminal deoxyribonucleotide transferase–mediated nick-end (TUNEL) staining. The percentage of TUNEL-positive B16F10 cells in a plug on day 3 was 4.5%, and the percentage of apoptotic cells decreased to 0.1% in Matrigel containing PAF. Etoposide increased the percentage of TUNEL-positive cells in a plug to almost 35%. This increase was completely blocked by PAF and decreased the percentage of cells to 2.5% (Fig. 5B). These data clearly indicate that PAF is capable of augmenting the growth of tumor cells by blocking apoptosis or by attenuating the cytotoxic effects of chemotherapeutic agents.

Figure 5.

PAF inhibits apoptosis of B16F10 melanoma cells in vivo. A, B16F10 cells (5 × 105) mixed in Matrigel containing 3 μmol/L PAF and/or 50 μmol/L etoposide were injected s.c., and tumor volumes were determined after growing tumors became visible. Points, mean tumor volumes of groups of five mice; bars, SE. B, tumor plugs were stained for internucleosomal DNA fragmentation using the ApopTag Plus Peroxidase In situ Apoptosis Detection kit. Arrowheads, apoptotic bodies. a, control group; b, PAF-treated group; c, etoposide-treated group; d, PAF- and etoposide-treated group.

Figure 5.

PAF inhibits apoptosis of B16F10 melanoma cells in vivo. A, B16F10 cells (5 × 105) mixed in Matrigel containing 3 μmol/L PAF and/or 50 μmol/L etoposide were injected s.c., and tumor volumes were determined after growing tumors became visible. Points, mean tumor volumes of groups of five mice; bars, SE. B, tumor plugs were stained for internucleosomal DNA fragmentation using the ApopTag Plus Peroxidase In situ Apoptosis Detection kit. Arrowheads, apoptotic bodies. a, control group; b, PAF-treated group; c, etoposide-treated group; d, PAF- and etoposide-treated group.

Close modal

PAF is known to play an important role in the growth of tumors (20, 21) and apoptosis is an important process in tumor growth. In this study, we showed that PAF up-regulated gene expression and protein synthesis of antiapoptotic factors, such as Bcl-2 and Bcl-xL. Our results are in accordance with the fact that PAF enhances the expression of Bcl-xL in human brain cells (26). However, experiments regarding the influence of PAF on apoptosis had rather controversial results. PAF has also been shown to protect B lymphocytes from apoptosis that occurs during immune cell maturation (12). In a similar manner, PAF can protect epidermal cells from tumor necrosis factor-α (TNF-α) and TNF-related apoptosis-inducing ligand (TRAIL)–induced apoptosis (27). In contrast, other groups have shown that PAF can augment UV radiation–induced apoptosis in epidermal cells (14), ionophore-induced apoptosis in T cells (28), and ischemia-reperfusion–induced mucosal apoptosis (29). This information suggests that PAF differently regulates apoptosis depending on cell types. This is further supported by the fact that PAF regulates apoptosis in a cell-specific manner (30, 31).

NF-κB modulates the expression of factors responsible for growth as well as apoptosis. Different NF-κB activation pathways may cause the expression of proteins that promote or inhibit apoptosis. Constitutive activation of NF-κB is generally seen as an emerging hallmark of various types of tumors, including those of the breast, colon, pancreatic, ovarian, and melanoma (3236). In melanoma, NF-κB has been shown to activate expression of antiapoptotic proteins, such as TNF receptor–associated factor 1 (TRAF1), TRAF2, and the inhibitor-of-apoptosis proteins, cIAP, cIAP2, and melanoma inhibitor of apoptosis, as well as Bcl-2-like proteins (3741). Indeed, TRAIL-induced apoptosis is suppressed by NF-κB-dependent transcription in tumor cell lines (40, 42). Alternatively, suppression of NF-κB activity switches the prevailing death pathway in melanoma from Fas ligand– to TNF-mediated apoptosis (43). Therefore, given that PAF is a potent inducer of NF-κB in vivo (22, 44, 45), our data that show a blocking of the PAF-induced up-regulation of antiapoptotic factors by a NF-κB inhibitor indicate that PAF exerted its effect via NF-κB activation.

Most chemotherapeutic agents exert their cytotoxic effects in part through the induction of apoptosis. Thus, we also assessed whether PAF could attenuate the cytotoxic effect of the chemotherapeutic agent, etoposide. Etoposide increased apoptosis and caspase activities in B16F10 cells, and these effects of etoposide were inhibited by PAF. These results confirm the inhibitory effect of PAF on apoptosis and suggest that PAF can attenuate the cytotoxic effect of chemotherapeutic agents. PAF is produced by a variety of inflammatory cells, including neutrophils, basophils, eosinophils, monocytes, and macrophages (46, 47). Additionally, PAF is released from certain tumor cells (19, 20, 48) as well as from tumor-infiltrating cells such as macrophages (49). Thus, it is possible that endogenous PAF may potentiate tumor cell growth, either through its inhibitory effect on apoptosis or by attenuating the cytotoxic effects of chemotherapeutic agents.

In summary, this study showed that PAF induced up-regulation of antiapoptotic factors in a NF-κB-dependent way and attenuated etoposide-induced apoptosis in a tumor cell line, and will provide important information to improve understanding of the mechanisms underlying apoptosis in tumors.

Note: K.H. Seo and H.M. Ko contributed equally to this work.

Grant support: Chonnam National University in the program, Post-Doc 2004 (K.H. Seo), and National R&D Program for Cancer Control, Ministry of Health and Welfare, Republic of Korea (0420060-1; S.Y. Im).

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.

1
Hockenbery D. Defining apoptosis.
Am J Pathol
1995
;
146
:
16
–9.
2
Thompson CB. Apoptosis in the pathogenesis and treatment of disease.
Science
1995
;
267
:
1456
–62.
3
Kuriyama H, Lamborn KR, O'Fallon JR, et al. Prognostic significance of an apoptotic index and apoptosis/proliferation ratio for patients with high grade astrocytomas.
Neuro-oncol
2002
;
4
:
179
–86.
4
Lipponen P, Aaltomaa S, Kosma VM, Syrjanen K. Apoptosis in breast cancer as related to histopathological characteristics and prognosis.
Eur J Cancer
1994
;
30A
:
2068
–73.
5
Sinicrope FA, Hart J, Hsu HA, Lemoine M, Michelassi F, Stephens LC. Apoptotic and mitotic indices predict survival rates in lymph node-negative colon carcinomas.
Clin Cancer Res
1999
;
5
:
1793
–804.
6
Vindigni C, Miracco C, Spina D, et al. Cell proliferation, cell death and angiogenesis in early and advanced gastric cancer of intestinal type.
Int J Cancer
1997
;
74
:
637
–41.
7
Kroemer G. The proto-oncogene Bcl-2 and its role in regulating apoptosis.
Nat Med
1997
;
3
:
614
–20.
8
Reed JC. Bcl-2 and the regulation of programmed cell death.
J Cell Biol
1994
;
124
:
1
–6.
9
Boise LH, Gonzalez-Garcia M, Postema CE, et al. Bcl-x, a bcl-2-related gene that functions as a dominant regulator of apoptotic cell death.
Cell
1993
;
74
:
597
–608.
10
Fukuda AI, Breuel KF. Effect of platelet-activating factor on embryonic development and implantation in the mouse.
Hum Reprod
1996
;
11
:
2746
–9.
11
Nilsson G, Metcalfe DD, Taub DD. Demonstration that platelet-activating factor is capable of activating mast cells and inducing a chemotactic response.
Immunology
2000
;
99
:
314
–9.
12
Toledano BJ, Bastien Y, Noya F, Mazer B. Characterization of B lymphocytes rescued from apoptosis by platelet-activating factor.
Cell Immunol
1999
;
191
:
60
–8.
13
Toledano BJ, Bastien Y, Noya F, Baruchel S, Mazer B. Platelet-activating factor abrogates apoptosis induced by cross-linking of the surface IgM receptor in a human B lymphoblastoid cell line.
J Immunol
1997
;
158
:
3705
–15.
14
Barber LA, Spandau DF, Rathman SC, et al. Expression of the platelet-activating factor receptor results in enhanced ultraviolet B radiation-induced apoptosis in a human epidermal cell line.
J Biol Chem
1998
;
273
:
18891
–7.
15
Hostettler ME, Knapp PE, Carlson SL. Platelet-activating factor induces cell death in cultured astrocytes and oligodendrocytes: involvement of caspase-3.
Glia
2002
;
38
:
228
–39.
16
Shi LC, Wang HY, Friedman E. Involvement of platelet-activating factor in cell death induced under ischemia/post-ischemia-like conditions in an immortalized hippocampal cell line.
J Neurochem
1998
;
70
:
1035
–44.
17
Reuther-Madrid JY, Kashatus D, Chen S, et al. The p65/RelA subunit of NF-κB suppresses the sustained, antiapoptotic activity of Jun kinase induced by tumor necrosis factor.
Mol Cell Biol
2002
;
22
:
8175
–83.
18
Khreiss T, Jozsef L, Chan JS, Filep JG. Activation of extracellular signal-regulated kinase couples platelet-activating factor-induced adhesion and delayed apoptosis of human neutrophils.
Cell Signal
2004
;
16
:
801
–10.
19
Bussolino F, Arese M, Montrucchio G, et al. Platelet-activating factor produced in vitro by Kaposi's sarcoma cells induces and sustains in vivo angiogenesis.
J Clin Invest
1995
;
96
:
940
–52.
20
Montrucchio G, Sapino A, Bussolati B, et al. Potential angiogenic role of platelet-activating factor in human breast cancer.
Am J Pathol
1998
;
153
:
1589
–96.
21
Biancone L, Cantaluppi V, Del Sorbo L, Russo S, Tjoelker LW, Camussi G. Platelet-activating factor inactivation by local expression of platelet-activating factor acetyl-hydrolase modifies tumor vascularization and growth.
Clin Cancer Res
2003
;
9
:
4214
–20.
22
Im SY, Han SJ, Ko HM, et al. Involvement of nuclear factor-κB in platelet-activating factor-mediated tumor necrosis factor-α expression.
Eur J Immunol
1997
;
27
:
2800
–4.
23
Prewett M, Huber J, Li Y, et al. Antivascular endothelial growth factor receptor (fetal liver kinase 1) monoclonal antibody inhibits tumor angiogenesis and growth of several mouse and human tumor.
Cancer Res
1999
;
59
:
5209
–18.
24
Im SY, Ko HM, Ko YS, et al. Augmentation of tumor metastasis by platelet-activating factor.
Cancer Res
1996
;
56
:
2662
–5.
25
Mizumoto K, Rothman RJ, Farber JL. Programmed cell death (apoptosis) of mouse fibroblasts is induced by the topoisomerase II inhibitor etoposide.
Mol Pharmacol
1994
;
46
:
890
–5.
26
Pulliam L, Zhou M, Stubblebine M, Bitler CM. Differential modulation of cell death proteins in human brain cells by tumor necrosis factor α and platelet-activating factor.
J Neurosci Res
1998
;
54
:
530
–8.
27
Southall MD, Isenberg JS, Nakshatri H, et al. The platelet-activating factor receptor protects epidermal cells from tumor necrosis factor (TNF) α and TNF-related apoptosis-inducing ligand-induced apoptosis through an NF-κB-dependent process.
J Biol Chem
2001
;
276
:
45548
–54.
28
Azzouzi B, Jurgens P, Benveniste J, Thomas Y. Immunoregulatory functions of PAF-acether. IX. Modulation of apoptosis in an immature T cell line.
Biochem Biophys Res Commun
1993
;
190
:
320
–4.
29
Wu B, Iwakiri R, Ootani A, Fujise T, Tsunada S, Fujimoto K. Platelet-activating factor promotes mucosal apoptosis via FasL-mediating caspase-9 active pathway in rat small intestine after ischemia-reperfusion.
FASEB J
2003
;
17
:
1156
–8.
30
Chandrasekher G, Ma X, Lallier TE, Bazan HEP. Delay of corneal epithelial wound healing and induction of keratocyte apoptosis by platelet-activating factor.
Invest Ophthalmol Vis Sci
2002
;
43
:
1422
–8.
31
Brewer C, Bonin F, Bullock P, et al. Platelet activating factor-induced apoptosis is inhibited by ectopic expression of the platelet activating factor G-protein coupled receptor.
J Neurochem
2002
;
82
:
1502
–11.
32
Wang W, Abbruzzese JL, Evans DB, Larry L, Cleary KR, Chiao PJ. The nuclear factor-κB RelA transcription factor is constitutively activated in human pancreatic adenocarcinoma cells.
Clin Cancer Res
1999
;
5
:
119
–27.
33
Bours V, Dejardin E, Goujon-Letawe F, Merville MP, Castronovo V. The NF-κB transcription factor and cancer: High expression of NF-κB- and IκB-related proteins in tumor cell lines.
Biochem Pharmacol
1994
;
47
:
145
–9.
34
Dejardin E, Bonizzi G, Bellahcene A, Castronovo V, Merville MP, Bours V. Highly-expressed p100/p52 (NFKB2) sequesters other NF-κB-related proteins in the cytoplasm of human breast cancer cells.
Oncogene
1995
;
11
:
1835
–41.
35
Duffey DC, Chen Z, Dong G, et al. Expression of a dominant-negative mutant inhibitor-κBα of nuclear factor-κB in human head and neck squamous cell carcinoma inhibits survival, proinflammatory cytokine expression, and tumor growth in vivo.
Cancer Res
1999
;
59
:
3468
–74.
36
Nakshatri H, Bhat-Nakshatri P, Martin DA, Goulet RJ, Jr., Sledge GW, Jr. Constitutive activation of NF-κB during progression of breast cancer to hormone-independent growth.
Mol Cell Biol
1997
;
17
:
3629
–39.
37
Deveraux QL, Roy N, Stennicke HR, et al. IAPs block apoptotic events induced by caspase-8 and cytochrome c by direct inhibition of distinct caspases.
EMBO J
1998
;
17
:
2215
–23.
38
Wang CY, Mayo MW, Korneluk RG, Goeddel DV, Baldwin AS, Jr. NF-κB anti-apoptosis: induction of TRAF1 and TRAF2 and c-IAP1 and c-IAP2 to suppress caspase-8 activation.
Science
1998
;
281
:
1680
–3.
39
Baldwin AS, Jr. The NF-κB and IκB proteins: New discoveries and insights.
Annu Rev Immunol
1996
;
14
:
649
–83.
40
Ravi R, Bedi GC, Engstrom LW, et al. Regulation of death receptor expression and TRAIL/Apo2L-induced apoptosis by NF-κB.
Nat Cell Biol
2001
;
3
:
409
–16.
41
Catz SD, Johnson JL. Transcriptional regulation of bcl-2 by nuclear factor κB and its significance in prostate cancer.
Oncogene
2001
;
20
:
7342
–51.
42
Franco AV, Zhang XD, Van Berkel E, et al. The role of NF-κB in TNF-related apoptosis-inducing ligand (TRAIL)-induced apoptosis of melanoma cells.
J Immunol
2001
;
166
:
5337
–45.
43
Ivanov VN, Fodstad O, Ronai Z. Expression of ring finger-deleted TRAF2 sensitizes metastatic melanoma cells to apoptosis via up-regulation of p38, TNFα and suppression of NF-κB activities.
Oncogene
2001
;
20
:
2243
–53.
44
Han SJ, Choi JH, Ko HM, et al. Glucocorticoids prevent NF-κB activation by inhibiting the early release of platelet-activating factor in response to lipopolysaccharide.
Eur J Immunol
1999
;
29
:
1334
–41.
45
Choi JH, Ko HM, Kim JW, et al. Platelet-activating factor-induced early activation of NF-κB plays a crucial role for organ clearance of Candida albicans.
J Immunol
2001
;
166
:
5139
–44.
46
Mencia-Huerta JM, Benveniste J. Platelet-activating factor and macrophage. I. Evidence for the release from rat and mouse peritoneal macrophages and not from mastocytes.
Eur J Immunol
1979
;
9
:
409
–15.
47
Camussi G, Aglietta M, Coda R, Bussolino F, Piacibello W, Tetta C. Release of platelet-activating factor (PAF) and histamine. II. The cellular origin of human PAF: monocytes, polymorphonuclear neutrophils and basophils.
Immunology
1981
;
42
:
191
–9.
48
Pitton C, Lanson M, Besson P, et al. Presence of PAF-acether in human breast carcinoma: relation to axillary lymph node metastasis.
J Natl Cancer Inst
1989
;
81
:
1298
–302.
49
Mantovani A, Sozzani S, Locati M, Allavena P, Sica A. Macrophage polarization: tumor-associated macrophages as a paradigm for polarized M2 mononuclear phagocytes.
Trends Immunol
2002
;
23
:
549
–55.