Tyrosine Phosphorylation of Protein Kinase Cδ Is Essential for Its Apoptotic Effect in Response to Etoposide

Materials.

An affinity-purified polyclonal anti-PKCɛ antibody against a polypeptide corresponding to amino acids 726 to 737 of PKCɛ was purchased from GIBCO-BRL Life Technologies (Gaithersburg, Md.). Monoclonal anti-PKCδ antibody directed against the regulatory domain was obtained from Transduction Laboratories (Lexington, Ky.), and polyclonal anti-PKC antibodies were from Santa Cruz (Santa Cruz, Calif.). Etoposide was from Alexis Co. (San Diego, Calif.), and an anti-active caspase 3 antibody was obtained from New England Biolabs (Beverly, Mass.). The caspase inhibitors DEVD-FMK, Z-VAD-FMK, and YVAD were obtained from Calbiochem (La Jolla, Calif.). Leupeptin, aprotinin, phenylmethylsulfonyl fluoride (PMSF), and sodium vanadate were obtained from Sigma Chemical Co. (St. Louis, Mo.). The Caspase 3 Cellular Activity Assay Kit PLUS was obtained from BIOMOL (Plymouth Meeting, Pa.), and the Cell Death Detection enzyme-linked immunosorbent assay (ELISA) Kit was from Roche Molecular Biochemicals.

Generation of PKC chimeras.

The PKC chimeras were generated by exchanging the regulatory and catalytic domains of PKCα and PKCδ as previously described (1). PKCα/δ refers to the chimera with the PKCα regulatory domain and the PKCδ catalytic domain, and PKCδ/α refers to the reciprocal chimera. The PKC cDNAs were subcloned into the metallothionein promoter-driven eukaryotic expression vector (MTH). The vector sequence encodes a C-terminal PKCɛ-derived 12-amino-acid tag (ɛMTH) that is added to the expressed proteins (47). The expression of these chimeras and their activities in C6 cells were recently described (5).

Site-directed mutagenesis of PKCδ.

Mouse PKCδ was cloned into the pGEM-T vector (Promega, Madison, Wis.) as described previously (5). This plasmid served as our “master” vector for the site-directed mutagenesis, using the Transformer Site-Directed Mutagenesis Kit from Clontech (Palo Alto, Calif.). Conversion of tyrosine residues at sites 52, 64, 155, 187, and 565 into phenylalanine was performed as previously described (5). PKCδ and the PKCδ mutants were subcloned into the metallothionein promoter-driven eukaryotic expression vector (ɛMTH). A PKCδ K376R dominant mutant was generated as previously described (36).

Construction of PKCδ-GFP fusion protein.

cDNAs encoding the murine PKCδ and the PKCδ5 mutant were fused into the N-terminal enhanced green fluorescent protein (GFP) vector pEGFP-N1 (Clontech Laboratories). The original pEGFP-N1 vector was modified by the insertion of an MluI site into the plasmid polylinker. The restriction site was created by ligating a phosphorylated linker containing the MluI site into pEGFP-N1 digested with SmaI. The construct was verified by sequencing. The clones containing GFP-PKCδ or GFP-PKCδ5 were constructed by the excision of PKCδ or PKCδ5 from MTH-PKC plasmids by digestion with XhoI and MluI. The inserts were then ligated into the modified GFP vector by using the same restriction sites. DNA sequencing of the GFP-PKC constructs confirmed the intended reading frame.

C6 glial cultures and cell transfection.

C6 cells (10⁵ cells/ml) were seeded on tissue culture dishes (10-cm diameter) and were grown in medium consisting of Dulbecco’s modified Eagle’s medium containing 10% heat-inactivated fetal calf serum, 2 mM glutamine, penicillin (50 U/ml), and streptomycin (0.05 mg/ml). The cells were transfected either with the empty vectors or with the PKCδ and PKCδ5 expression vectors by using Lipofectamine (Gibco-BRL Life Technologies) as previously described (6). Experiments were routinely carried out on a clone of the transfected cells, but all the results were confirmed on one pool and two additional individual clones.

For overexpression of the GFP-PKCδ fusion proteins, C6 cells were seeded onto 40-mm round glass coverslips at a density of 5 × 10⁴ cells/coverslip. Twenty-four hours later, cells were transfected with the different GFP-PKCδ constructs using Lipofectamine Plus reagent according to the manufacturer’s instructions. All experiments were performed 48 h posttransfection.

Measurements of cell apoptosis.

Cell apoptosis was measured using propidium iodide (PI) staining and analysis by flow cytometry and by ELISA (Cell Death Detection ELISA Kit) using anti-histone antibodies. Cells (1 × 10⁶/ml) were plated in six-well plates and treated with the indicated treatments for 24 h. Detached cells and trypsinized adherent cells were pooled, fixed in 70% ethanol for 1 h on ice, washed with phosphate-buffered saline (PBS), and treated for 15 min with RNase (50 μM) at room temperature. Cells were then stained with PI (5 μg/ml) and analyzed on a Becton-Dickinson cell sorter.

For anti-histone ELISA (Cell Death Detection ELISA kit), extracts of cells containing histone-associated DNA fragments were incubated in 96-well plates coated with anti-histone antibodies for 2 h. Plates were then washed and incubated with anti-DNA antibodies conjugated to peroxidase for an additional 2 h. Substrate solution was added and absorbance was measured at 405 nm.

Cell viability was also quantitatively assessed by the measurement of lactate dehydrogenase (LDH) in the medium.

Measurement of caspase 3 activity.

Caspase 3 activity was measured using the caspase 3 colorimetric assay kit obtained from BIOMOL by using Ac-DEVD-pNA as a substrate according to the manufacturer’s recommendations.

Nuclear and cytosolic fractionation.

Nuclear proteins were prepared according to the method described by Haglund and Rothblum (24). Cells (1 × 10⁶ to 5 × 10⁶/ml) were washed once with 1 ml of PBS and once with 1 ml of lysis buffer (10 mM HEPES, 10 mM KCl, 1.5 mM MgCl₂, 0.5 mM PMSF, 10 μg of leupeptin/ml, pH 7.9). Cells were lysed by suspending the cell pellet in 20 μl of lysis buffer containing 0.1% Nonidet P-40 (NP-40) for 10 min on ice. To isolate nuclei, the lysates were microcentrifuged for 5 min at 12,000 × g, and the nuclear pellet was washed with lysis buffer without NP-40. Nuclear proteins were obtained by resuspending the nuclear pellet in 20 μl of extraction buffer (420 mM NaCl, 20 mM HEPES, 1.5 mM MgCl₂, 0.2 mM EDTA, and 25% glycerol, pH 7.9) for 10 min at 4°C. The nuclear suspension was microcentrifuged, the pellet was discarded, and the supernatant was diluted in dilution buffer (50 mM KCl, 20 mM HEPES, 0.2 mM EDTA, and 20% glycerol, pH 7.9). Lack of contamination of the nuclear fraction by the plasma membrane was confirmed using the plasma membrane marker Na-K ATPase.

Preparation of cell homogenates.

Cells were washed and resuspended in serum-free medium. The plates were placed on ice, scraped with a rubber policeman, and centrifuged at 1,400 rpm for 10 min. The supernatants were aspirated, and the cell pellets were resuspended in 100 μl of lysis buffer (25 mM Tris-HCl, pH 7.4, 50 mM NaCl, 0.5% Na deoxycholate, 2% NP-40, 0.2% sodium dodecyl sulfate [SDS], 1 mM PMSF, 50 μg of aprotinin/ml, 50 μM leupeptin, 0.5 mM Na₃VO₄) on ice for 15 min. The cell lysates were centrifuged for 15 min at 14,000 rpm in an Eppendorf microcentrifuge, supernatants were removed, and 2× sample buffer was added.

Immunoblot analysis.

Lysates (40 μg of protein) were resolved by SDS-polyacrylamide gel electrophoresis (PAGE) (10% polyacrylamide) and were transferred to nitrocellulose membranes. The membranes were blocked with 5% dry milk in PBS and subsequently stained with the primary antibody. Specific reactive bands were detected using a goat anti-rabbit or goat anti-mouse IgG conjugated to horseradish peroxidase (Bio-Rad, Hercules, Calif.), and the immunoreactive bands were visualized by the ECL Western blotting detection kit (Amersham, Arlington Heights, Ill.).

Immunoprecipitation.

Immunoprecipitation was performed as previously described (5). Briefly, C6 cells overexpressing PKCδ or PKCδ5 were serum starved overnight and treated for different periods of time with PMA (10 nM) or PDGF (100 ng/ml). The samples were preabsorbed with 25 μl of protein A/G-Sepharose (50%) for 10 min, and immunoprecipitation was performed using 4 μg of antibody/ml for 1 h at 4°C and then incubated with 30 μl of protein A/G-Sepharose for an additional hour. Following washes, the pellets were resuspended in 25 μl of SDS sample buffer and boiled for 5 min. The entire supernatants were subjected to Western blotting. Membranes were incubated with horseradish peroxidase-conjugated secondary antibodies for 1 h at room temperature. The membranes were washed and visualized by the enhanced chemiluminescence (ECL) system.

Immunofluorescence staining.

Cells were grown on glass coverslips. Following etoposide treatment (3 to 24 h), cells were washed with PBS and fixed in 4% paraformaldehyde (PFA) for 10 min. Subsequently, cells were washed in PBS, and after blocking with staining buffer (2% bovine serum albumin [BSA] and 0.1% Triton X-100 in PBS) for 30 min at room temperature, cells were incubated with a mouse (regulatory domain) or a rabbit (catalytic domain) anti-PKC antibody. Following washes in PBS, cells were incubated with an anti-rabbit antibody or with Alexa FluorTM 546 goat anti-mouse IgG for an additional 60 min and were mounted in FluoroGuard antifade reagent. Cells were viewed and photographed using confocal microscopy with ×63 magnification at excitation wavelengths of 543 and 488 nm.

For the translocation of GFP-PKCδ and GFP-PKCδ5, cells were treated for 6 and 24 h with etoposide, were fixed in 4% PFA for 10 min, and were mounted in FluoroGuard antifade reagent.

Statistical analysis.

The results are presented as the mean values ± standard error (SE). Data were analyzed using analysis of variance (ANOVA) and a paired Student’s t test to determine the level of significance between the different groups.

Etoposide induces apoptosis and cell cycle arrest in C6 glioma cells.

Treatment of the cells with etoposide arrested the cells in the G₁/S phase of the cell cycle and induced cell apoptosis (Fig. 1A). Using PI staining and fluorescence-activated cell sorter (FACS) analysis, we found that approximately 22% ± 3.9% of the cells underwent apoptosis in response to etoposide after 24 h of treatment, whereas approximately 63% ± 7.1% of the cells were apoptotic after 48 h. Similar results were obtained using anti-histone ELISA (Fig. 1B) and using measurements of LDH levels in the cell supernatants (data not shown).

Treatment of the cells with etoposide also resulted in lower cell number and in the appearance of rounded and detached cells, which are characteristic of apoptotic cells (Fig. 1C).

Role of PKCδ in apoptotic effect of etoposide.

Recent studies suggested that PKCδ plays a role in the apoptotic effect of etoposide in salivary gland acinar cells (49). To explore the role of PKCδ in the effect of etoposide on C6 cells, we utilized rottlerin, which has been reported to be a PKCδ selective inhibitor (22). Treatment of C6 cells with rottlerin (5 μM) did not affect the basal level of cell apoptosis; however, rottlerin inhibited the apoptotic effect of etoposide (Fig. 2A), reducing cell apoptosis by approximately 50%. Higher concentrations of rottlerin itself induced morphologic changes in the cells and some cell apoptosis and therefore could not be used.

Since the in vitro inhibitory effect of rottlerin on PKCδ activity has recently been subject to controversy (12, 22, 30), we further explored the role of PKCδ in the apoptotic effect of etoposide by employing cells overexpressing a PKCδ dominant-negative mutant (K376R) (36). Cells stably expressing the PKCδ DN mutant were treated with ZnCl₂ for 24 h (to induce overexpression through its MTH promoter), followed by a 48-h treatment with etoposide (50 μM). Overexpression of PKCδ DN did not affect the basal level of C6 cell apoptosis; however, it reduced the apoptosis induced by etoposide by 40% (Fig. 2B).

The role of PKCδ in the apoptotic effect of etoposide was also demonstrated using cells stably overexpressing PKCδ. The expression of PKCδ in these cells was seven- to eightfold higher than control vector cells (35). These cells exhibited an increased rate of apoptosis in response to etoposide (45%) compared to cells expressing the control vector (20%; Fig. 2C).

Etoposide induces nuclear translocation of PKCδ.

Activation of PKC by different stimuli results in distinct patterns of translocation, which are cell and stimulus dependent. To examine the effect of etoposide on the translocation of PKCδ, we treated C6 cells with etoposide for various periods of time and followed the expression of PKCδ using immunofluorescence and confocal microscopy. In control cells, PKCδ was found largely in the cytosol, but with some expression in the nucleus (Fig. 3). Treatment with etoposide induced further translocation of PKCδ to the nucleus. Translocation was observed already after 3 h, and the expression of PKCδ in the nucleus was observed up to 24 h following treatment. We confirmed that etoposide did not induce translocation of PKCδ to the Golgi, endoplasmic reticulum, or the mitochondria, as determined by costaining with these organelle-specific markers (data not shown).

No translocation of PKCα, -β, -ζ, -μ, and -ι was observed in etoposide-treated cells, whereas some translocation of PKCɛ to the perinuclear membrane was observed (data not shown).

Cleavage of PKCδ by etoposide.

PKCδ undergoes cleavage in response to various apoptotic stimuli (21, 49). To examine the effect of etoposide on the cleavage of PKCδ in our system, we treated C6 cells with etoposide (50 μM) for various periods of time and analyzed cell lysates by using Western blotting. The level of PKCδ decreased following 24 h of etoposide treatment, and a 40-kDa cleavage product of PKCδ appeared and started to accumulate (Fig. 4A). No significant cleavage of PKCα, -β, -ɛ, -ζ, and -μ was observed in response to etoposide, and no cleavage products were detected (data not shown).

Since the nuclear translocation of PKCδ preceded its caspase-dependent cleavage, we examined if PKCδ cleavage occurred in the nucleus. We first performed cell fractionation and separated the nuclear and cytosolic fractions. We found that in untreated cells, PKCδ was mainly expressed in the cytosol with some expression in the nucleus. Following etoposide treatment, PKCδ translocated to the nucleus and a cleaved form of PKCδ accumulated only in the nuclear fraction (Fig. 4B). The nuclear marker lamin B was also detected only in the nuclear fraction. Using anti-PKCδ antibodies directed against the regulatory and catalytic domains, we found that the immunostaining of PKCδ using these two antibodies resulted in a similar pattern of nuclear translocation (Fig. 4C). The results of these two experiments suggest that PKCδ translocated to the nucleus, where it underwent cleavage. Since some PKCδ was present in the nucleus of the untreated cells without being cleaved, the cleavage must depend on the etoposide treatment itself as well as on its location.

Caspase 3 is involved in cleavage of PKCδ and apoptosis is induced by etoposide.

PKCδ can be cleaved by a caspase-dependent process (21, 34). We therefore examined the effects of the cell-permeable caspase 3 inhibitor DEVD.FMK (20 μM) on the cleavage of PKCδ in the C6 cells. As seen in Fig. 5A, pretreatment of the cells for 1 h with DEVD.FMK significantly reduced the accumulation of the PKCδ cleavage product in response to etoposide. Similar results were obtained with another caspase inhibitor, Z-VAD.FMK (data not shown).

We also examined the ability of the caspase inhibitors to block the apoptosis induced by etoposide in C6 cells. Pretreatment of the cells with either DEVD.FMK or Z-VAD.FMK reduced the apoptotic effect of etoposide by approximately 60% (Fig. 5B), suggesting a role for caspase 3 in the apoptotic response. In contrast, the caspase 1 inhibitor YVAD did not affect etoposide effects in these systems (data not shown).

Since the cleavage of PKCδ occurred in the nucleus, we examined whether active caspase 3 was also expressed in the nucleus in etoposide-treated cells. Using immunofluorescence staining, we found that the active form of caspase 3 was expressed in the nucleus of etoposide-treated cells, whereas a very low expression of active caspase 3 was detected in control cells (Fig. 5C). Similar results were obtained using Western blot analysis of nuclear and cytosolic fractions (Fig. 5D).

Inhibition of PKCδ blocks cleavage of caspase 3 and PKCδ.

To examine the importance of PKCδ activity in its ability to undergo cleavage, we employed the selective PKCδ inhibitor rottlerin. Pretreatment of the cells with rottlerin (5 μM) reduced the cleavage of caspase 3 in response to etoposide (Fig. 6A). Similarly, cells overexpressing PKCδ DN exhibited a significantly lower caspase 3 activity than control vector cells (Fig. 6B). Finally, the cleavage of PKCδ and the accumulation of the PKCδ catalytic domain were reduced in rottlerin-treated cells (Fig. 6C), suggesting that the cleavage of both caspase 3 and PKCδ is dependent on an active PKCδ.

Both regulatory and catalytic domains of PKCδ are required for apoptotic effect induced by etoposide.

In a recent study, we demonstrated that the regulatory domain of PKCδ mediated its inhibitory effect on C6 cell proliferation and on the expression of the astrocytic marker GS (5). The regulatory domain of PKCδ was phosphorylated on tyrosines 155 and 187 in response to PMA and PDGF, respectively (5, 35), and the translocation of the PKCδ/α chimera resembled that of PKCδ (40). To examine the relative contributions of the regulatory and catalytic domains of PKCδ to its apoptotic effects, we used chimeras between the regulatory and catalytic domains of PKCα and -δ combined at the highly conserved hinge region. The expression and activity of C6 cells overexpressing the different chimeras were already described (5).

Cells were pretreated for 24 h with ZnCl₂, followed by etoposide treatment (50 μM) for an additional 48 h. Cells overexpressing PKCδ/δ exhibited levels of apoptosis similar to those of control vector-transfected cells. Treatment of the control vector cells with etoposide induced about 60% apoptosis, similar to the results obtained with the parental C6 cells, whereas cells overexpressing PKCδ exhibited increased levels of apoptosis. Cells overexpressing PKCα/α exhibited a rate of cell apoptosis similar to that of the control vector cells. Interestingly, cells overexpressing the chimera containing the regulatory domain of PKCδ (δ/α) or the chimera containing the catalytic domain of PKCδ (α/δ) exhibited a similar level of apoptosis to that of control vector cells and did not resemble cells overexpressing PKCδ (Fig. 7). These results suggest that both domains are required for the apoptotic effect of PKCδ.

Etoposide induces tyrosine phosphorylation of PKCδ.

PKCδ has been shown to undergo tyrosine phosphorylation in response to PMA, growth factors, and apoptotic stimuli, such as H₂O₂ and γ-irradiation (5, 35, 58). To examine the effect of etoposide on tyrosine phosphorylation of PKCδ, we treated C6 cells with etoposide for 15 to 180 min. PKCδ was immunoprecipitated, and the membrane was blotted with anti-phosphotyrosine antibody. As illustrated in Fig. 8A, a low basal level of tyrosine phosphorylation was observed in untreated cells. Etoposide induced tyrosine phosphorylation of PKCδ in a time-dependent manner. Initial phosphorylation was observed after 45 min (data not shown), maximal phosphorylation was obtained after 60 min and was decreased thereafter (Fig. 8A). Tyrosine phosphorylation of PKCδ in response to etoposide was lower than that induced by PMA and exhibited slower kinetics (5, 35). The tyrosine phosphorylation induced by etoposide was specific for PKCδ. We did not observe tyrosine phosphorylation of any of the other isoforms (data not shown).

We next examined whether the tyrosine phosphorylation in response to etoposide occurred in the regulatory or catalytic domains. For these experiments, we used the PKC chimeras α/δ and δ/α. Treatment of the chimeras with etoposide for 60 min resulted in the phosphorylation of PKCδ/δ and of the chimera containing the regulatory domain of PKCδ (PKCδ/α). The chimera containing the catalytic domain of PKCδ did not exhibit an increase in tyrosine phosphorylation in response to etoposide (Fig. 8B), suggesting that the tyrosine phosphorylation occurred in the regulatory domain.

Tyrosine phosphorylation of PKCδ is essential for its apoptotic effect.

In previous studies, we identified five putative tyrosine phosphorylation sites in PKCδ and generated a PKCδ5 mutant, in which these five tyrosine phosphorylation sites were mutated to phenylalanine (5). The expression and activity of C6 cells overexpressing this mutant were already described (5). Overexpression of the PKCδ5 mutant in C6 cells abolished the decrease in the expression of the astrocytic marker GS induced by PMA or PDGF and the decrease in cell proliferation induced by PMA. Expression of the PKCδ5 mutant also resulted in a lower level of tyrosine phosphorylation of PKCδ in response to these treatments (5). Using cells expressing PKCδ5, we found that etoposide did not induce a significant increase in tyrosine phosphorylation of PKCδ5 (Fig. 9A). Similarly, treatment of these cells with etoposide resulted in low levels of apoptosis similar to the response observed in the control vector cells when measured after 24 h (Fig. 9B) and in lower levels of apoptosis than control vector cells after 48 h of treatment (Fig. 9C). Thus, the tyrosine phosphorylation of PKCδ in the regulatory domain induced by etoposide was essential for the apoptotic effect of PKCδ in response to this drug.

Cleavage of caspase 3 and PKCδ in cells overexpressing PKCδ5 mutant.

To further explore the role of tyrosine phosphorylation of PKCδ in the apoptosis induced by etoposide, we compared the activation of caspase 3 in cells overexpressing control vector, PKCδ, and the PKCδ5 mutant. Using a specific antibody recognizing the cleaved product (17 kDa) of caspase 3, we found that etoposide induced caspase 3 cleavage in the control vector cells. Cells overexpressing PKCδ showed a larger amount of the cleaved product, whereas cells overexpressing the PKCδ5 mutant exhibited very low levels of the 17-kDa fragment (Fig. 10A). Similar results were obtained for caspase 3 activity as measured by a caspase 3 colorimetric assay (Fig. 10B). These results suggest that the activation of caspase 3 required a tyrosine phosphorylated form of PKCδ.

We then examined the cleavage of PKCδ and PKCδ5 in response to etoposide. Using the ɛ tag, we were able to detect the cleaved catalytic domain of the exogenous PKCδ and PKCδ5. Using the anti-PKCɛ antibody, we found no detectable cleaved product in the etoposide-treated control vector cells. Accumulation of the catalytic fragment was observed in cells overexpressing PKCδ, whereas cells overexpressing the PKCδ5 mutant displayed no detectable cleavage (Fig. 10C). Since the kinetics of PKCδ phosphorylation is faster than the cleavage and is transient, the phosphorylation presumably plays a role upstream from the actual cleavage.

Translocation of PKCδ and the PKCδ 5 mutant in response to etoposide.

One possible explanation for the differential effects of PKCδ and the PKCδ5 mutant on cell apoptosis is their differential translocation following etoposide treatment. We therefore examined the translocation of PKCδ and the PKCδ5 mutant in response to etoposide. For these experiments, we used GFP-tagged PKCδ wild type or the PKCδ5 mutant. Cells were transiently transfected with the specific construct, and the response of the cells to etoposide was monitored after 6 and 24 h.

We found that etoposide induced a similar pattern of translocation for both PKCδ and PKCδ5 (Fig. 11). Thus, stimulation of the cells with etoposide for 6 h induced translocation of PKCδ-GFP to the nucleus in about 90 to 95% of the cells, similar to the results obtained using immunofluorescent staining of the endogenous PKCδ. Likewise, PKCδ5-GFP translocated to the nucleus in response to etoposide in about 90% of the cells. A similar pattern of nuclear translocation of PKCδ and PKCδ5 was also obtained after 24 h of etoposide treatment (data not shown).

Role of single tyrosine mutants in apoptotic response of etoposide.

To identify the specific tyrosines that are involved in the inhibitory effect of PKCδ on cell apoptosis, we examined the role of the single tyrosines 52, 64, 155, 187, and 565. C6 cells were stably transfected with the different PKCδ mutants in which each one of these tyrosines was individually mutated to phenylalanine. The expression and activity of C6 cells overexpressing the different PKC mutants were previously described (35).

The apoptotic effect of etoposide was examined in cells overexpressing the different mutants using PI staining and FACS analysis. We found that cells overexpressing PKCδY52F, PKCδY155F, and PKCδY565F exhibited an enhanced apoptotic response to etoposide similar to that of cells overexpressing PKCδ. In contrast, treatment with etoposide of cells overexpressing PKCδY64F or PKCδY187F resulted in a lower apoptotic response, similar to the response observed with cells overexpressing the PKCδ5 mutant (Fig. 12A). Similar results were obtained with anti-histone ELISA (Fig. 12B).

We also found that etoposide did not induce a significant increase in the tyrosine phosphorylation of PKCδY64F and PKCδY187F, similar to the results obtained with PKCδ5 (Fig. 12C). Finally, we found no detectable cleaved product of PKCδ in the etoposide-treated PKCδY64F and PKCδY187F overexpressors (data not shown).

Apoptosis induced by etoposide is not inhibited by Src-kinase inhibitors PP1 and PP2.

In a previous study, we showed that Fyn and Lyn can associate with PKCδ and that Fyn associates with PKCδ via tyrosine 187 (35). To examine the role of Src-related kinases in the apoptotic response of etoposide, we employed the Src-kinase inhibitors PP1 and PP2. Pretreatment of the cells with either PP1 (10 μM) or PP2 (10 μM) did not affect the apoptotic response of etoposide (data not shown), suggesting that Src-related kinases are probably not involved in the tyrosine phosphorylation of PKCδ and in the apoptotic response of etoposide.