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TOXICOLOGY
Division of Toxicology, Leiden/Amsterdam Center for Drug Research, Leiden University, Leiden, The Netherlands
Received June 16, 2004; accepted September 17, 2004.
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Abstract |
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-catenin in association with PKC-mediated phosphorylation of the actincapping protein adducin. These events preceded and were independent of caspase activation. -Catenin did not dissociate from E-cadherin. Cisplatin-induced loss of cell-cell contacts was associated with the increased formation of F-actin stress fibers, which was inhibited by Bis I and Gö6983 as well as dominant-negative PKC-
. Also, the loss of cell-cell adhesions by cisplatin was prevented by Bis I and Gö6983. Activation of protein kinase C with phorbol esters promoted cisplatin-induced loss of cell-cell adhesions as well as apoptosis. In conclusion, the combined data fit a model whereby protein kinase C mediates the cisplatin-induced loss of cellular interactions. Such a loss of these interactions has a role in the onset of apoptosis.
; Lieberthal and Levine, 1996). Cisplatin causes various biochemical and cellular perturbations in renal cells, including DNA cross-linking (Yasumasu et al., 1992); decreased capacity of several transport systems such as Na+/K+-ATPase activity (Nowak, 2002); reduced activity of antioxidant enzymes such as manganese superoxide dismutase, glutathione peroxidase, and glutathione transferase (Sadzuka et al., 1994) in association with lipid peroxidation; mitochondrial dysfunction (Kruidering et al., 1997); cytoskeletal reorganization (van de Water et al., 1994a); and changes in cell adhesion (Kruidering et al., 1998). Cisplatin-induced oxidative stress is important in the necrosis of PTC. Cisplatin-induced renal cell apoptosis follows the common degradation pathway, which involves the activation of caspase-9 and caspase-3 followed by caspase substrate cleavage, nuclear condensation, and DNA fragmentation and is blocked by Bcl-2 (Kruidering et al., 1997; Kaushal et al., 2001). So far, not much is known about the more upstream cellular perturbations that link cisplatin injury to the onset of apoptosis.
There is increasing evidence that injury to PTC caused by either ischemia-related ATP depletion or nephrotoxic chemicals results in impaired adhesion to the extracellular matrix (ECM) (cell-ECM adhesion) as well as to neighboring cells (cell-cell adhesion) (Racusen et al., 1991; Kroshian et al., 1994). Both cell-ECM as well as cell-cell interactions are vitally important in providing the renal epithelial cells with survival cues to prevent the onset of apoptosis; these are largely mediated through the activation of the phosphoinositide-3 kinase/protein kinase B pathway (King et al., 1997). Cell-ECM adhesions are mediated by integrin receptors that link the ECM to the cellular actin cytoskeleton (Richardson and Parsons, 1995; Schwartz, 1997). Cell-cell contacts are mediated by E-cadherins that form transdimers at the adherens junctions (Steinberg and McNutt, 1999). E-Cadherins are indirectly linked to the cortical actin cytoskeleton through -catenin and 
-actinin. Cisplatin-induced apoptosis is also associated with changes in cell adhesion. In primary cultured pig proximal tubular epithelial cells, cisplatin causes a drastic decrease of ECM expression and a reorganization of the actin cytoskeletal network (Kruidering et al., 1994), yet cell-ECM signaling does not seem to be a major component that controls cisplatin-induced apoptosis (van de Water et al., 2000). In primary cultures of rat renal proximal tubular epithelial (RPTE) cells, cisplatin causes early disruption of cell-cell interactions that are independent of caspase activity and precede the onset of apoptosis. It is not clear which molecular mechanisms are involved in this process and what role the loss of these interactions plays in cisplatin-induced apoptosis of PTC.
The cortical actin cytoskeleton is central in the regulation of cell-cell adhesions, which is tightly regulated by various signal transduction molecules. Protein kinase C (PKC) is a family of signaling molecules that is involved in the regulation of a variety of processes, including proliferation, differentiation, transformation, and apoptosis (Kiley et al., 1996; Mellor and Parker, 1998). These biological effects are linked to the differential expression profile and cellular location of PKC isoenzymes. These isoenzymes are divided into three subclasses, i.e., classic, novel, and atypical, according to their requirement of Ca2+, phosphatidylserine, and diacylglycerol. PKCs are important in the regulation of the actin cytoskeleton. Thus, the actin-capping protein adducin was identified as a major substrate for PKC and its phosphorylation on serine residue 726, which results in the dissociation from the actin filaments and translocation to the cytosol. Previously, we found that adducin is phosphorylated on serine 726 in RPTE cells after cisplatin treatment (van de Water et al., 2000). In rabbit proximal tubular cells, cisplatin causes mitochondrial injury that is associated with increased phosphorylation of PKC-, indicative of activation (Nowak, 2002). So far, a role for PKC in the control of cell-cell interactions in relation to cisplatin-induced apoptosis has not been investigated. In the current study, we establish such a role.
Using the renal LLC-PK1, we demonstrate that cisplatin causes increased formation of F-actin stress fibers in conjunction with subtle changes in the organization of adherens junctions preceded by caspase activation and apoptosis. Pharmacological inhibitors of novel PKC, but not inhibitors of classic PKC, protect against cisplatin-induced apoptosis. This is associated with protection against the reorganization of the actin cytoskeleton and loss of adherens junctions. Together, the data indicate an important role for PKC controlled restructuring of and signaling by cell-cell adhesions in cisplatin-induced apoptosis.
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Materials and Methods |
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Cell Culture, Transfection, and Treatment. LLC-PK1 cells of porcine proximal tubular origin (American Type Culture Collection, Manassas, VA) were cultured in DMEM supplemented with 10% (v/v) FBS and 50 U of penicillin/liter and 50 mg of streptomycin/liter (penicillin/streptomycin). Cultures were maintained in a humidified incubator gassed with 5% CO2 and 95% air at 37°C. The medium was changed every other day.
For experiments, the cells were cultured at a density of 100,000 cells/ml in 24-well dishes containing collagen-coated glass coverslips, six-well dishes (Corning Costar, Acton, MA), or 10-cm dishes (Greiner, Frickenhausen, Germany). After 5 days, experiments were performed when the cells reached 100% confluence.
For transient transfection, LLC-PK1 cells were plated overnight on collagen-coated coverslips in complete medium without antibiotics. Cells were transfected with dominant-negative (DN)-PKC- and DN-PKC- (Dr. P. J. Parker, Cancer Research Institute, London, UK) using LipofectAMINE 2000 (Invitrogen) as indicated by the supplier. Forty hours after transfection, cells were treated with cisplatin as indicated above followed by immunofluorescent staining. Cells were treated with cisplatin in the absence or presence of the indicated concentrations of Bis I (1 mM stock in DMSO), Gö6976 (1 mM stock in DMSO), Gö6983 (1 mM stock in DMSO), or PDBu (2 mM stock in DMSO) in DMEM supplemented with 10% (v/v) FBS and 50 U of penicillin/liter and 50 mg of streptomycin/liter.
Isolation and Culturing of Rat Renal Proximal Tubular Epithelial Cells. RPTE cells were isolated by collagenase perfusion and separated by density centrifugation using Nycodenz as previously described (van de Water et al., 1994b). Cells were cultured on rat tail collagen (Collaborative Research, Bedford, MA)-coated dishes in DMEM/F-12 containing 1% (v/v) FBS, 0.5 mg/ml BSA, 10 µg/ml insulin, 10 ng/ml epidermal growth factor, 10 ng/ml cholera toxin, and antibiotics as previously described [complete medium A (van de Water et al., 1999)]. Cells were maintained at 37°C in a humidified atmosphere of 95% air and 5% carbon dioxide and fed every other day. RPTE cells were used after they had reached confluence, 6 to 9 days after plating. Cells were treated in complete culture medium with indicated concentrations of cisplatin in the presence or absence of inhibitors of protein kinase C, and after 24 h apoptosis was determined by cell cycle analysis as well as determination of caspase-3-like activity. Necrosis and secondary apoptosis were determined after 24 and 48 h by determination of the percentage of lactate dehydrogenase (LDH) release.
Cell Cycle Analysis. Apoptosis was determined by cell cycle analysis. Medium containing floating cells was collected. Adherent cells were washed twice with PBS containing 1 mM EDTA and trypsinized. Floating cells and adherent cells that were trypsinized were pooled, centrifuged for 10 min at 2000 rpm, and resuspended in 100 µl of PBS followed by fixation in 90% ethanol (–20°C). Fixed cells were centrifuged and washed once with PBS followed by resuspension in PBS-EDTA containing 7.5 µM propidium iodide and 10 µg/ml RNase A. After 30 min at room temperature (RT), cells were analyzed by flow cytometry (FACSCalibur; BD Biosciences, Franklin Lakes, NJ). The amount of cells in subG0/G1 was calculated using the CellQuest software (BD Biosciences).
Determination of LDH Release. Cell death was measured by the release of LDH from cells in the culture medium as previously described (Chen et al., 1990). The percentage of cell death was calculated from the amount of LDH release caused by treatment with toxicants relative to the amount to that released by 0.1% (w/v) Triton X-100 (i.e., 100% release).
Caspase Activity Measurement. Cells were scraped in medium and collected by centrifugation together with floating cells. The cell pellet was taken up in lysis buffer (10 mM HEPES, 40 mM -glycerophosphate, 50 mM NaCl, 2 mM MgCl2, and 5 mM EGTA) and subjected to three cycles of freezing and thawing in liquid nitrogen. The suspension was centrifuged at 13,000 rpm in a microfuge for 30 min. The supernatant was collected and used to determine the protein concentration using the Bradford protein assay using IgG as a standard. Equal amounts of cell protein (10 µg) were used for measuring caspase activity using Ac-DEVD-AMC as a substrate (25 µM). AMC fluorescence was followed in time using a fluorescence plate reader (HTS 7000 BioAssay Reader; PerkinElmer Life and Analytical Sciences, Boston, MA). Caspase activity was calculated as picomoles per milligram of cell protein per minute using AMC as a standard.
Paracellular Transport of Fluorescein-Labeled Dextran. LLC-PK1 cells were cultured on 12-well transwell dishes with a pore size of 0.4 µm (Corning Costar). Cells were allowed to form a tight monolayer for 7 days. Thereafter, cells were treated with cisplatin in complete medium both in the apical (0.5 ml) and basolateral (1.5 ml) compartment. After 8 h of treatment, cisplatin was removed from both compartments, and cells were incubated for an additional period with fluorescein-labeled 4-kDa dextran (4FD; 200 µg/ml; Sigma-Aldrich) dissolved in Hanks'/HEPES buffer in the basolateral compartment. After 4 h, 100-µl samples were taken from both the basolateral and apical compartment, and the 4FD was analyzed using a fluorescent plate reader (HTS 7000 BioAssay Reader; PerkinElmer Life and Analytical Sciences).
Preparation of Cytoskeletal and Soluble Cellular Fractions. To obtain soluble and cytoskeletal fractions, the Triton X-100 extraction method was used (Dong et al., 1995). Briefly, RPTE cells cultured in 10-cm dishes were treated with cisplatin as described above. The medium was removed, and adherent cells were washed twice with PBS and a final wash with microtubule stabilization buffer (MSB; 100 mM PIPES, 2 M glycerol, 1 mM EGTA, and 1 mM magnesium acetate, pH 6.9). Adherent cells were scraped in 500 µlof MSB buffer containing protease and phosphatase inhibitors and 0.2% (w/v) Triton X-100 (MSB Plus). Floating cells were collected from the pooled washes by centrifugation for 5 min at 500g. Pelleted cells were mixed with the adherent cells in MSB buffer. Cells were extracted for 4 min at room temperature. The homogenate was centrifuged at 20,000g for 20 min at 15°C. The supernatant (soluble fraction) was removed, and the pellet (cytoskeleton fraction) was resuspended in MSB Plus. Equal amounts of protein were separated by SDS-PAGE followed by Western blotting.
Gel Electrophoresis and Immunoblotting. Medium with floating cells was collected and centrifuged (5 min, 500g, 4°C). Adherent cells were kept on ice and washed twice in ice-cold PBS and once in lysis buffer (10 mM Tris-HCL, 250 mM sucrose, and 1 mM EGTA, pH 7.4). Cells were then scraped and resuspended in lysis buffer containing 1 mM dithiothreitol, 10 µg/ml leupeptin, 10 µg/ml aprotinin, 1 mM sodium vanadate, 50 mM sodium fluoride, and 1 mM phenylmethylsulfonylfluoride. Floating cells in pellet were pooled with scraped cells. Cells were then sonicated, and the protein concentration was determined using the Bradford protein assay using IgG as a standard. Equal amounts of total cellular protein were separated by SDS-PAGE and transferred to polyvinylidene difluoride membrane (Millipore Corporation, Bedford, MA). Membranes were blocked for 1 h in the following blocking buffer: 5% (w/v) BSA in Tris-buffered saline-Tween 20 [0.5 M NaCl, 20 mM Tris-HCl, and 0.05% (v/v) Tween 20, pH 7.4]. Primary antibody incubation was performed overnight at 4°C for active caspase-3 (CM-1 kindly provided by Dr. A. Srinivasan, Idun Pharmaceuticals, La Jolla, CA); -catenin (clone 14; Transduction Laboratories, Lexington, KY); panphospho-Ser660-PKC (Cell Signaling Technology Inc., Beverly, MA); phospho-Ser729-PKC- (BioSource International, Camarillo, CA); PKC- (clone M1; Upstate Biotechnology); PKC- (clone 21; BD Biosciences); adducin and phospho-adducin (Fowler et al., 1998) (kindly provided by Dr. S. Jaken, Lilly Research Laboratories, Indianapolis, IN); -tubulin (Sigma-Aldrich); and protein disulfide isomerase (PDI; Stressgen Biotechnologies, Inc., San Diego, CA). Secondary antibody incubation was performed using either horseradish peroxidase-coupled secondary antibodies or Cy-5-labeled secondary antibodies. Visualization was performed with the ECL Plus reagent (Amersham Biosciences Inc., Piscataway, NJ) for the horseradish peroxidase-coupled antibodies or directly with the Cy-5-labeled antibodies, followed by imaging with the Typhoon 9400 multilabel imager (Amersham Biosciences Inc.).
Immunofluorescence and Fluorescence Microscopy. Cells were cultured on 12-mm collagen-coated glass coverslips and fixated in 3.7% (w/v) formaldehyde in PBS. Coverslips were blocked in TBP [0.5% (w/v) BSA and 0.1% (v/v) Triton X-100 in PBS, pH 7.4 (1 h, RT)] and subsequently incubated with primary antibody in TBP (O/N, 4°C). Coverslips were washed three times in TBP and incubated with Alexa-488, Cy-3, or Cy-5-conjugated secondary antibodies or 0.2 U/ml rhodamine phalloidin (Molecular Probes, Eugene, OR) in TBP (1 h, RT). After washing with TBP, coverslips were incubated with 2 µg/ml Hoechst 33258 in PBS (15 min, RT), washed in PBS, and mounted in Aqua Polymount (Polysciences, Warrington, PA). Cells were viewed using a Bio-Rad Radiance 2100 MP confocal laser scanning system (Bio-Rad, Hercules, CA) equipped with a Nikon Eclipse TE2000-U inverted fluorescence microscope and a 60x Nikon objective (Nikon, Melville, NY). Images were processed with Paint Shop Pro 7.02.
Statistical Analysis. Student's t test was used to determine whether there was a significant difference between two means (p < 0.05); statistical differences are indicated with an asterisk. When multiple means were compared, significance was determined by oneway analysis of variance (p < 0.05). For analysis of variance, letter designations are used to indicate significant differences: common letters means not different; different letters means statistically different.
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Results |
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; data not shown). At 25 µM, the process of apoptosis started after 8 h. This apoptosis was associated with a time- and concentration-dependent increase in the enzyme activity of caspase-3, with a maximal activity after 24 h; this was associated with the formation of the active form of caspase-3 as well as the formation of cleavage products of caspase-3 substrates such as -catenin, as determined by Western blotting (Fig. 1C). Importantly, the general caspase inhibitor zVADfmk blocked all of the events typical of apoptosis, indicating the requirement for caspase activity (Fig. 1C). Since we wanted to study the effect of cisplatin on processes that take place before the cells become apoptotic, a concentration of 25 µM was chosen throughout our studies.
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Renal proximal tubular epithelial cells are the most important targets for cisplatin toxicity within the kidney. In most cases of nephrotoxicity, kidney damage is caused by an impaired adhesion of proximal tubular epithelial cells. Since cell-cell interactions are essential in providing cells with survival signals and suppressing apoptosis, we hypothesized that perturbations of such interactions could be observed after cisplatin treatment. Therefore, we studied the process of cisplatin-induced cell detachment in LLC-PK1 cells. Confluent monolayers of LLC-PK1 cells form so-called domes (Fig. 2). The formation of these domes is dependent on well formed cell-cell interactions; consequently, the loss of these interactions results in the loss of domes. To study the effect of cisplatin on dome formation, LLC-PK1 cells were exposed to 25 µM cisplatin, and the domes were counted at different time points. A decrease in dome size and quantity started after 8 h of treatment (Fig. 2). After 16 and 24 h of exposure to cisplatin, domes were no longer seen. In fact, many cells were now detached and floating in the medium. These floating cells represent apoptotic cells, which is consistent with the apoptosis results (Fig. 1). Although it was apparent by light microscopy that cells lose cell-cell interactions, we wanted to verify that the loss of domes was indeed related to the loss of cell-cell adhesion structures and not to perturbations of transcellular water transport. Since tight cell-cell interactions in epithelial cell monolayers prevent paracellular transport/diffusion, we reasoned that if cisplatin causes the loss of cell-cell interactions, paracellular diffusion would be facilitated. To test this hypothesis, we performed transport studies with 4FD molecules using transwell culture dishes. Although hardly any 4FD was transported from the basolateral site to the apical site of the transwell under control conditions, treatment with cisplatin caused a collapse of the barrier and allowed the paracellular diffusion of 4FD (Table 1). Since various caspase-3 substrates control cellular adhesions (in) directly, including -catenin and E-cadherin (Schmeiser and Grand, 1999; Van de Water et al., 1999), the possibility existed that the loss of domes is purely related to cisplatin-induced caspase activity. To investigate this possibility, we exposed LLC-PK1 cells to cisplatin in combination with zVADfmk. Although zVADfmk prevented cisplatin-induced apoptosis and -catenin cleavage (Fig. 1), it did not block the loss of domes after cisplatin treatment (Fig. 2).
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Since the early loss of domes suggested changes in the organization of cell-cell interaction, we studied these interactions in more detail. The most important cell-cell contacts are formed at the adherens junctions by intercellular E-cadherin homodimer formation. E-cadherin is linked to the actin cytoskeletal network through -catenin, -catenin, and actinin. Since changes in both the localization of these molecules as well as the actin cytoskeletal network indicate changes in the cell-cell interactions, their localization and organization were studied using immunofluorescent staining and confocal laser scanning microscopy (CLSM) (Fig. 3). After 8 h, cells exposed to cisplatin showed a subtle but clear reorganization of -catenin-containing adherens junctions from large plaques to thinner structures (Fig. 3). In addition, the F-actin network shifted from a more intense cortical actin cytoskeletal network with less stress fibers to a condition with a less intense cortical actin and more stress fibers at the basement membrane plane after cisplatin treatment (Fig. 3). After 24 h of cisplatin treatment, most of the -catenin had disappeared from the sharp cell-cell boundaries. Since apoptosis is already present at this time point and -catenin is a caspase-3 substrate, we wanted to make sure that this effect was not related to increased caspase-3 activity. Therefore, cells were also immunostained for active caspase-3. Importantly, those cells that had lost -catenin from the membrane did not yet have activated caspase-3. Treatment with the pan-caspase inhibitor zVADfmk, which inhibited cisplatin-induced -catenin cleavage (Fig. 1), did not affect cisplatin-induced -catenin translocation (Fig. 3). Together, these data indicate that the reorganization of -catenin at the cell membrane is not caused by caspase-3-mediated cleavage of -catenin.
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Critical Role of PKC in Cisplatin-Induced Apoptosis of Renal Epithelial Cells. PKC is known to phosphorylate and thereby regulate several cytoskeletal-associated proteins. Previously, we showed that phosphorylation of the actin-capping protein adducin is preceded by its cleavage in RPTE cells (Dong et al., 1995; van de Water et al., 2000). Also, in primary rabbit proximal tubular epithelial cells, cisplatin exposure leads to the phosphorylation of PKC-, indicative of activation (Nowak, 2002). Therefore, PKC might be a good candidate involved in the weakening of the cellular adhesion. Such a weakening of the cell adhesion may facilitate the onset of apoptosis. First, we determined the effect of cisplatin on PKC. PKC function is regulated in a complex manner by multiple phosphorylation sites in the protein. Thus, the serine/threonine residue homologues to serine 660 in PKC-II is a critical autophosphorylation site that is present in several PKC isoforms and modulates the activity/localization/turnover of PKCs (Keranen et al., 1995; Feng and Hannun, 1998). We used a phospho state-specific antibody that binds the phosphorylated form of this residue present in various PKC isoforms, including , , 
, and . In LLC-PK1, no direct effect of cisplatin (either 25 or 100 uM) was observed after 8 h on the phosphorylation of PKC using either pan-phospho-PKC antibody or phospho-PKC- antibodies (Fig. 4A). Since PKC activity is also associated with translocation, we also determined the effect of cisplatin on the localization of phosphorylated PKC. Cisplatin (25 uM) caused an increased localization of phosphorylated PKC at the cell-cell junction, indicative for the modulation of PKC (Fig. 4B). To further study a role for PKC in cisplatin-induced apoptosis, we used several pharmacological inhibitors of PKC: Gö6976, a selective inhibitor of the Ca2+-dependent PKC isozymes and I; and Bis I and Gö6983, two more general PKC inhibitors (Kiley et al., 1999; Way et al., 2000). Dose finding experiments indicated that the most optimal concentration of these inhibitors for the inhibition of cytotoxicity was 1 µM. At this concentration, Bis I and Gö6983 both significantly reduced the cisplatin-induced apoptotic cell death after 24 h (Fig. 5, A and B). In contrast, Gö6976 did not affect the cisplatin-induced apoptosis (Fig. 5C). To verify whether this protection against apoptosis was related to the inhibition of caspase-3 activation, we also measured the caspase-3 enzyme activity. Indeed, both Bis I and Gö6983, but not Gö6976, inhibited the activation of caspase-3 caused by cisplatin (Fig. 5, D–F). The combined data suggest that PKC plays an important role in the cisplatin-induced apoptotic pathway. We reasoned that if this is indeed the case, then the activation of PKC by the addition of phorbol esters would promote cisplatin-induced apoptosis. Indeed, cotreatment of LLC-PK1 cells with PDBu (200 nM) increased both the time point of the onset as well as the percentage of apoptosis caused by cisplatin, further supporting a role of PKC in the regulation of cisplatin-induced apoptosis (Fig. 6, A and B).
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In our previous work, we demonstrated that cisplatin causes the phosphorylation of the PKC substrate adducin before the onset of apoptosis in primary cultured RPTE cells (van de Water et al., 2000). We first determined the effect of cisplatin on PKC phosphorylation and localization in RPTE cells. Cisplatin caused a time-dependent increase of phosphorylated PKC in RPTE cells, with a maximum increase after 16 h (Fig. 4C). After 24 h, phosphorylated PKC was decreased. To verify that the phosphorylation was related to activation, cells were also treated with an activator of PKC, PDBu. PDBu caused an increase in the phosphorylation of PKC after 30 min, which already decreased after 1 h and remained lower until 8 h (Fig. 4C). This implies a rapid turnover of phosphorylated PKC upon activation. Next, we also determined the localization of phosphorylated PKC in RPTE cells after cisplatin treatment. Although phosphorylated PKC was located primarily in the cytosol and nucleus in untreated cells, after cisplatin treatment, phosphorylated PKC was also associated with cell-cell junctions in a similar fashion as observed for LLC-PK1 cells (data not shown). To verify whether similar findings were observed with another phospho-PKC antibody, we also used a phospho state-specific antibody that specifically recognizes PKC- when phosphorylated on serine 729. Cisplatin also caused a time-dependent phosphorylation of PKC- in a similar fashion as for the pan-phospho-PKC antibody. To check whether PKC is also involved in the regulation of cisplatin-induced apoptosis in primary cultured RPTE cells, these cells were treated with cisplatin in combination with the PKC inhibitors Bis I, Gö6983, and Gö6976. Bis I, Gö6983, and Gö6976 all inhibited the apoptosis of RPTE cells, as determined by cell cycle analysis and caspase-3 activity measurement (Fig. 7). Since apoptosis is generally followed by secondary necrosis, which is associated with the leakage of plasma membrane, we also measured the release of LDH from the cells in the medium. At 24 h, no significant difference in the levels of LDH release was observed between controls (data not shown). After 48 h, cisplatin caused the release of LDH, indicative of (secondary) necrosis, which was prevented by both Bis I, Gö6983, and Gö6976 (Fig. 8). In RPTE cells, activation of PKC by PDBu also caused an increase in cisplatin-induced cell death (Fig. 8). Finally, we wanted to exclude a possible required cooperation of growth factor present in the medium for the cisplatin effects. Therefore, in an other set of experiments, we treated RTPE cells with 25, 50, or 100 µM in Hanks'/HEPES buffer for 8 h followed by a recovery in complete medium until 24 h in the presence of the above inhibitors of PKC. Under these conditions, protection against cisplatin-induced cell death was also observed by the inhibition of PKC (data not shown). Together, the combined data indicate a critical role for PKC in the control of cisplatin-induced apoptosis of renal proximal tubular epithelial cells.
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PKC Is Involved in Cisplatin-Induced Loss of Cell-Cell Contacts and F-Actin Reorganization. The above data indicated that cisplatin causes an early loss of domes in association with a reorganization of adherens junctions preceding the onset of apoptosis. The fact that PKC is important in the regulation of cell-cell adhesions prompted us to investigate whether PKC also regulates the cisplatin-induced loss of cell-cell adhesion. Both Bis I and Gö6983 inhibited the rapid loss of domes caused by cisplatin (Fig. 9, A and B). Importantly, there were still domes detected even after 24 h, although the domes were somewhat smaller than in the control situation. Gö6976 did again not protect against the rapid cisplatin-induced loss of domes. In conjunction with the apoptosis data, the combined treatment of cisplatin and PDBu accelerated the loss of domes (data not shown).
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Since PKC mediates the phosphorylation of cytoskeletal components that on their turn control, among others, the cortical actin cytoskeletal network organization/turnover, it is most likely that the inhibition of PKC also protected against a reorganization of the cortical actin network and/or localization of adherens junction components, such as E-cadherin and -catenin. When LLC-PK1 cells were treated with Bis I and Gö6983 in combination with cisplatin, F-actin stress fiber formation was inhibited, and the localization of -catenin was maintained at the site of the cell membrane (Fig. 10A); however, Gö6976 did not protect against the translocation of -catenin and the formation of F-actin stress fibers (Fig. 10B). When cells were treated with PDBu and cisplatin, the organization of adherens junctions was lost already after 8 h, and -catenin staining was drastically decreased (Fig. 8B). This was independent of caspase activity since zVADfmk did not block this effect of PDBu (data not shown). Since cisplatin-induced perturbations of cellular interactions were associated with increased actin stress fiber formation, we wanted to determine the relationship between PKC activity and F-actin formation after cisplatin treatment. Although cisplatin itself caused a pronounced increase in the number of actin filaments at the basement membrane (see Fig. 3), less stress fibers were detected when PKC was inhibited by either Gö6983 or Bis I (Fig. 10A). As an alternative approach to determine the role of PKC in cisplatin-induced cytoskeletal reorganization, we also transfected LLC-PK1 cells with dominant-negative-acting PKC- and PKC-, followed by an analysis of F-actin rearrangement in transfected cells. Both DN-PKC- and DN-PKC--positive cells were clearly distinguishable (Fig. 11). DN-PKC- overexpression inhibited F-actin stress fiber formation caused by cisplatin compared with untransfected cells. This effect was not observed for DN-PKC-. Neither DN-PKC- nor DN-PKC- affected the actin cytoskeleton in untreated cells (Fig. 11). Together, the data indicate that PKC is involved in cisplatin-induced loss of the cell-cell contacts in direct relation to the increased formation of actin stress fibers.
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Cisplatin Increases Phosphorylation of Adducin without Affecting Cytoskeletal Association of -Catenin. The above data indicate that cisplatin affects the localization of -catenin as determined by immunofluorescence. In RPTE cells, cisplatin causes a phosphorylation of the actin-capping protein adducin. Therefore, we next investigated whether -adducin is also phosphorylated in LLC-PK1 cells after cisplatin treatment and whether this is associated with a translocation of -catenin from the cytoskeleton to the cytosol. For this purpose, LLC-PK1 cells were exposed to cisplatin for different time points, and cytoskeletal and soluble cellular fractions were prepared using Triton X-100 extraction. The endoplasmic reticulin-associated protein PDI was used as a control for the extraction procedure (Fig. 12A). Cisplatin caused an increase in the phosphorylation of the serine residue 726 of both the - and 
-adducin isoform. This increased phosphorylated adducin was primarily observed in the cytoskeleton fraction; after 8 h, no increased phosphorylated adducin was present in the cytosol (Fig. 12A). Despite the changes in adducin phosphorylation and F-actin organization after cisplatin treatment, no obvious change in the association of -catenin with the cytoskeleton or its location in the cytosol was observed. Next, we evaluated the effect of both Bis I and Gö6983 on the phosphorylation status of adducin and -catenin localization in LLC-PK1 cells. Both Bis I and Gö6983 reduced the phosphorylation of adducin, indicating that PKC is involved in this process. These two PKC inhibitors did not affect the association of -catenin with the cytoskeleton.
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Finally, we also checked whether the cisplatin-induced reorganization of -catenin was associated with changes in the binding of -catenin and E-cadherin using immunoprecipitation. Cisplatin did not lead to any major change in the interaction between -catenin and E-cadherin after either 8 or 12 h (Fig. 12B). Together, this suggests a role for PKC in the reorganization of the actin cytoskeleton through phosphorylation of substrates that regulate the actin cytoskeleton.
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) and that this is associated with an early phosphorylation of the PKC substrate -adducin (van de Water et al., 2000). In the present study we found that 1) PKC activity is essential for the induction of apoptosis of PTC by cisplatin; 2) cisplatin causes an early formation of F-actin stress fibers and loss of cell-cell interactions; and 3) PKC is directly involved in the cisplatin-induced cytoskeletal reorganization and loss of adherens junctions. The early reorganization of the cytoskeletal network and relocation of -catenin from cell-cell contacts occurred prior to the activation of caspases; moreover, inhibition of caspases did not affect these changes caused by cisplatin. Therefore, we propose that an early PKC-mediated reorganization of the actin cytoskeletal network is involved in the reorganization of adherens junctions and decreased cell-cell adhesion. This may provide a condition that resembles an anoikis status and, hence, facilitates the activation of the apoptotic machinery.
Previous studies have identified various cell adhesion molecules that by themselves are a target for proteolytic cleavage by the group of executioner caspases. These include both cell-matrix adhesion (cytoskeletal) components such as focal adhesion kinase and proteins that are involved in the regulation of cell-cell contacts, such as -catenin (Wijnhoven et al., 2000) and -adducin (van de Water et al., 2000). Our data in LLC-PK1 cells indicate that the rearrangement of the actin cytoskeletal network is independent of caspase activity. Thus, caspase-3 activation caused by cisplatin could be blocked by zVADfmk. Despite this blockade and protection against -catenin cleavage, zVADfmk did not prevent the loss of -catenin from the sites of cell-cell contacts, indicating that this process is independent of caspase-3 activation. Furthermore, cisplatin increased the formation of actin stress fibers in the cells in a caspase-3-independent fashion. Together, these data indicate that other mechanisms that are activated after exposure to cisplatin induced the reorganization of the cytoskeleton and adherens junctions that by itself may directly be involved in the onset of apoptosis.
Our findings indicate that PKC activity is critical for the induction of apoptosis by cisplatin both in LLC-PK1 cells and primary cultured RPTE cells. Thus, Bis I, a more general inhibitor of PKC isozymes, significantly down-regulated cisplatin-induced apoptosis in LLC-PK1 cells as well as RPTE cells. In addition, increased activation of PKC with PDBu potentiated the cisplatin-induced caspase activation and cell death. Since Bis I is a general PKC inhibitor, we used other, more PKC isozyme-specific inhibitors Gö6983 and Gö6976 to check whether inhibition of apoptosis is mediated through certain isozymes of PKC. Gö6983, which is selective for novel PKC isozymes, showed significant inhibition of apoptosis comparable to Bis I in LLC-PK1 cells. In contrast, such an effect was not observed in these cells with Gö6976, which is more specific for PKC- (Kiley et al., 1999; Way et al., 2000; Nowak, 2002); this suggests that PKC- has no direct role in the regulation of apoptosis caused by cisplatin in LLC-PK1 cells. In contrast, in RPTE cells, Gö6976 protected against cisplatin-induced apoptosis, which was comparable to the effect of Gö6976 reported by Nowak (2002). This discrepancy between LLC-PK1 and primary cultured rat and rabbit PTC may be related to different species. Since LLC-PK1 are derived from pig kidneys that more closely resemble human physiology than rat and rabbit kidney, the question remains whether cisplatin-induced nephrotoxicity in human PTC is inhibited by either Gö6976 or Gö6983. Alternatively, the difference may be explained by the fact that RPTE cell cultures already have a dense F-actin stress fiber network and relatively little cortical actin under control conditions (van de Water et al., 1999), whereas LLC-PK1 cells have a strong cortical F-actin with little stress fibers under control conditions. Regardless of the differences, the combined studies clearly indicate an essential role of PKC in cisplatin-induced cytotoxicity. Such a role for PKC may also bear relevance for the in vivo situation: cisplatin treatment of rats causes elevated activity of PKC in proximal tubulus cells, whereas nephrotoxicity was inhibited by H-7, a very general inhibitor of PKC as well as other serine/threonine protein kinases (Ikeda et al., 1999). It will be important to determine next whether other, more specific inhibitors of PKC also inhibit cisplatin-induced renal failure.
Inhibition of cisplatin-induced apoptosis by Bis I and Gö6983 was associated with almost complete protection against the loss of cell-cell interactions in LLC-PK1 cells. These interactions are tightly controlled by the organization of the actin network, suggesting that PKC may directly act on this network after cisplatin treatment. This would be in line with the work of Dong and coworkers, who identified various cytoskeletal-associated proteins that are substrates of and interact with PKC (so-called STICKS; substrates that interact with C kinases); these include -adducin, -adducin, myristoylated alanine-rich C kinase substrate (MARCKS), and desmoyokin (Dong et al., 1995). Interestingly, in our previous study and herein, we identified that one of these STICKS, -adducin, has an increased phosphorylation on serine residue 726 after treatment of both LLC-PK1 cells as well as primary cultured rat renal proximal tubular cells with cisplatin (Fig. 12). -Adducin is an actin-capping protein that has a role in the regulation of the cortical actin cytoskeleton and is located at cell-cell contacts. Phosphorylation of adducin leads to the weakening of its interaction with the actin cytoskeleton and may thereby affect its capping activity and allow for changes in the actin cytoskeletal turnover. This is consistent with our data that cisplatin causes increased stress fiber formation. Both Bis I and Gö6983 reduced cisplatin-induced phosphorylation of -adducin in the LLC-PK1 cells, which was associated with inhibition of stress fiber formation and maintenance of proper cell-cell interactions. Together, this suggests that an increased phosphorylation of adducin may be related to a more active actin cytoskeletal turnover process at the cortical cytoskeleton after cisplatin treatment. Adducin as well as potential other proteins are direct targets for PKC-mediated regulation of the actin cytoskeletal network.
Our studies show that cisplatin exposure leads to an increase in stress fiber formation at the basement membrane of the cells, which was inhibited, but not completely prevented by, Bis I and Gö6983, but not Gö6976. Activation of PKC with PDBu resulted in increased F-actin stress fiber formation already after 4 h when treated with cisplatin. Together, these data strongly suggest the involvement of PKC in the formation of the stress fibers. The PKC- isozyme can bind actin and colocalizes in some cells with F-actin filaments. In RPTE cells, we found that cisplatin causes an increased mobility phosphorylation of PKC- (Fig. 4). Since both Bis I (general PKC inhibitor) and Gö6983 (inhibitor of PKC- and -), but not Gö6976 (inhibitor of PKC- and -), inhibited F-actin reorganization, the combined observations suggest a possible role for PKC- in the increased formation of the F-actin network after cisplatin treatment. This is supported by our findings that a DN-PKC- inhibits the formation of F-actin stress fibers after cisplatin treatment, whereas DN-PKC- does not (Fig. 11). In rabbit PTC, inhibition of PKC with Gö6976 was associated with protection against mitochondrial dysfunctioning after cisplatin treatment; no protective effect was described at earlier time points (Nowak, 2002). Therefore, it cannot be excluded that the late protection against mitochondrial dysfunctioning merely correlates with the observed cytoprotection against apoptosis. We would propose that the early PKC-dependent cytoskeletal rearrangement and loss of cell-cell contacts cause disturbances in the cellular homeostasis that by themselves may lead to mitochondrial dysfunctioning and/or accelerate the mitochondrial dysfunctioning that will be caused by cisplatin itself (Kruidering et al., 1997).
Our data indicate that the pharmacological inhibition of PKC inhibits cisplatin-induced cytotoxicity at a clinically relevant concentration of cisplatin. In the event that such inhibitors would be used for renal protection, PKC inhibition should not affect the effect of anticancer reagents on the tumors. Importantly, we recently showed that inhibition of PKC with either Gö6983 or Gö6976 strongly potentiated the cytotoxicity of the anticancer agent doxorubicin in the mammary adenoma-carcinoma cell line MTLn3 (Huigsloot et al., 2003). This would indicate that inhibition of PKC has the potential on the one hand to potentiate the anticancer effects of cytostatics and on the other hand prevent the unwanted cytotoxicity on renal cells.
In conclusion, we have identified that cisplatin-induced renal cell apoptosis is related to a PKC-dependent loss of cell-cell interactions. Pharmacological inhibitors of PKC prevent both the loss of cellular interactions as well as the onset of apoptosis. Future studies should further evaluate the possibilities of modulating PKC activity in in vivo conditions, thereby not only preventing cisplatin-induced renal failure, but also enhancing the cytostatic action of the anticancer drugs.
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Acknowledgements |
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and DN-PKC- constructs were kindly provided by Dr. Peter J. Parker. |
Footnotes |
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ABBREVIATIONS: PTC, proximal tubular epithelial cells; ECM, extracellular matrix; RPTE, renal proximal tubular epithelial; PKC, protein kinase C; DMEM, Dulbecco's modified Eagle's medium; FBS, fetal bovine serum; BSA, bovine serum albumin; AMC, 7-amino-4-methylcoumarin; Bis I, bisindolylmaleimide I; Gö6976, 12-(2-cyanoethyl)-6,7,12,13-tetrahydro-13-methyl-5-oxo-5H-indolo(2,3-a)pyrrolo(3,4-c)-carbazole; PDBu, phorbol 12,13-dibutyrate; Ac-DEVD-AMC, acetyl-Asp-Glu-Val-Asp-AMC; DN, dominant-negative; LDH, lactate dehydrogenase; 4FD, fluorescein-labeled 4-kDa dextran; PBS, phosphate-buffered saline; RT, room temperature; MSB, microtubule stabilization buffer; PAGE, polyacrylamide gel electrophoresis; PDI, protein disulfide isomerase; TBP, blocking buffer; CLSM, confocallaser scanning microscopy; Gö6983, 2-[1-(3-dimethylaminopropyl)-5-methoxyindol-3-yl]-3-(1H-indol-3-yl) maleimide.
Address correspondence to: Dr. B. van de Water, Division of Toxicology, Leiden/Amsterdam Center for Drug Research, Gorlaeus Laboratoria, P.O. Box 9502, 2300 RA Leiden, The Netherlands. E-mail: b.water{at}lacdr.leidenuniv.nl
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