Departments of 1Pharmaceutical Sciences and 2Internal Medicine, University of Arkansas for Medical Sciences, Little Rock, Arkansas 72205; and 3Department of Pharmaceutical Sciences, Medical University of South Carolina, Charleston, South Carolina 29425
Submitted 20 June 2002 ; accepted in final form 7 May 2003
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ABSTRACT |
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renal proximal tubular cells; caspase activation; DNA damage; knockout mice
The cyclin-dependent kinase inhibitor p21WAF1/CIP1 plays a pivotal role in cell differentiation, DNA repair, and apoptosis through regulation of the cell cycle. The p21WAF1/CIP1 protein is constitutively present at low levels in the nucleus of most cells as a complex with cyclin, cyclin-dependent kinase, and proliferating cell nuclear antigen (23, 49). p21WAF1/CIP1 is known to be a mediator of p53 tumor suppressor function and has been implicated in apoptosis caused by numerous agents. DNA damage caused by radiation and various chemotherapeutic drugs activates p53, which upregulates p21WAF1/CIP1 mRNA and protein levels. Induction of p21WAF1/CIP1 is required for the p53-dependent arrest in the G1 phase of the cell cycle after DNA damage (2, 46). However, transcriptional and posttranscriptional changes in p21WAF1/CIP1 expression after DNA damage can also be induced in a p53-independent manner (11, 29).
p21WAF1/CIP1 has differential effects on the sensitivity of cancer cells to ionizing radiation, cisplatin, and other antineoplastic agents, such as adriamycin, taxol, and vincristine, protecting against cell death in some models and not in others (7, 8, 37). For example, increased levels of p21WAF1/CIP1 enhance sensitivity of hepatoma and osteosarcoma cells to cisplatin (22). In contrast, in colon cancer and embryonic fibroblast cells, increased sensitivity to cisplatin and other chemotherapeutic agents is associated with the lack of p21WAF1/CIP1 (7, 37).
p21WAF1/CIP1 is rapidly induced in the murine kidney in response to acute renal failure (ARF) produced by ischemia-reperfusion, ureteral obstruction, and cisplatin (29). The p21WAF1/CIP1 induction decreases renal damage after cisplatin- and ischemia-induced ARF in murine models (26, 28). In contrast, the lack of the p21WAF1/CIP1 gene accelerates progression to ARF and chronic renal failure, increases morphological and functional damage to proximal tubular cells and the kidney, and results in a higher mortality rate (26-28). The mechanisms of the protective effects of p21WAF1/CIP1 are not clear. It was shown that cells having DNA damaged by cisplatin administration enter the cell cycle, but cell-cycle progression is inhibited in the presence of p21WAF1/CIP1. In contrast, kidney cells lacking p21WAF1/CIP1 progress from the G1 to S phase of the cell cycle after cisplatin-induced damage (28). Therefore, it has been proposed that p21WAF1/CIP1 decreases cisplatin nephrotoxicity by decreasing the number of injured cells that enter the cell cycle and undergo mitosis with damaged DNA, which results in cell death by either apoptosis or oncosis (28).
The aim of this study was to examine the cellular basis for increased cell death in p21WAF1/CIP1 knockout mice by investigating the temporal activation and the role of caspase-3, caspase-8, and caspase-9 during cisplatin-induced apoptosis in mouse renal proximal tubular cells (MPTC) obtained from p21WAF1/CIP1 knockout and wild-type mice.
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MATERIALS AND METHODS |
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Animals. Adult mice carrying a deletion of a large portion of the p21WAF1/CIP1 gene, in which neither p21WAF1/CIP1 mRNA nor p21WAF1/CIP1 protein is expressed, were obtained from Dr. Philip Leder (Harvard Medical School, Boston, MA) (5). Mice homozygous for the p21WAF1/CIP1 deletion were selected from the offspring of heterozygous matings using Southern blotting of tail DNA as described previously (5). The animals used in these studies were housed at the Veterinary Medical Unit at the John L. McClellan Memorial Veterans Hospital (Little Rock, AR). Wild-type homozygous mice 129Sv (obtained from The Jackson Laboratory, Bar Harbor, ME) were used as controls. Female mice (12-18 wk old) were used in this study.
Isolation of proximal tubules. Mouse renal proximal tubules were isolated by a modification of the method described by Sheridan et al. (40). The basal isolation medium was Hanks' solution and a 50:50 mixture of DMEM and Ham's F-12 nutrient mix (DMEM/F-12) supplemented with 29 mM NaHCO3, 15 mM HEPES, and penicillin G (150 U/ml). The medium was adjusted to pH 7.4 while being gassed with 95% O2-5% CO2 and was diluted to 295 mosmol/kgH2O before filter sterilization.
The animals were anesthetized with halothane, and both kidneys were removed, dissected under sterile conditions, and placed in ice-cold Hanks' solution gassed with 95% O2-5% CO2. Cortical tissue was removed, placed in fresh ice-cold Hanks' solution, and finely minced with a scalpel blade. Minced tissue was incubated for 30 min at 37°C (with shaking) in digestion medium consisting of Hanks' solution, collagenase type 4 (140 U/ml), and soybean trypsin inhibitor (0.75 mg/ml). Large undigested fragments of cortical tissue were separated by gravity after the mixing of equal volumes of the tissue suspension and ice-cold 10% horse serum in Hanks' solution. Following sedimentation of undigested tissue, the supernatant containing cortical tubules was collected and centrifuged for 2 min x 50 g at 4°C. The pellet was washed with ice-cold Hanks' solution, centrifuged for 2 min x 50 g at 4°C, washed again with DMEM/F-12, centrifuged, and resuspended in DMEM/F-12 medium. Cortical tubules were purified by gradient centrifugation in a 40% Percoll/60% DMEM/F-12 at 36,000 g x 20 min at 4°C. The band containing renal proximal tubules was collected and washed twice with DMEM/F-12, and the final pellet was resuspended in DMEM/F-12.
Culture conditions. The culture medium was a 50:50 mixture of DMEM/F-12 nutrient mix without phenol red and pyruvate, supplemented with 15 mM NaHCO3, 15 mM HEPES, 1 mM glucose, and 5 mM lactate (pH 7.4, 295 mosmol/kgH2O). Renal proximal tubule segments were plated in 35-mm culture dishes (0.3 mg protein/dish) and grown in optimized conditions as described previously for rabbit renal proximal tubular cells (34). Culture dishes were constantly swirled on an orbital shaker to improve media oxygenation. Human transferrin (5 µg/ml), selenium (5 ng/ml), hydrocortisone (50 nM), bovine insulin (5 µg/ml), and L-ascorbic acid-2-phosphate (50 µM) were added to the medium immediately before daily media change (2 ml/dish).
O2 consumption. Confluent MPTC monolayers were gently detached from the dishes with a rubber policeman, suspended in 37°C culture medium, and transferred to the O2 consumption (QO2) measurement chamber. QO2 was measured polarographically using a Clark-type electrode as described previously (33, 34). Uncoupled QO2 was used as a marker of the activity of electron transfer through the respiratory chain and was measured after addition of carbonyl cyanide p-(trifluoromethoxy)phenylhydrazone (2 µM).
Active Na+ transport. Ouabain-sensitive QO2 was used as a marker of active Na+ transport as described previously (33). Ouabain-sensitive QO2 was measured in the presence of 1 mM ouabain and calculated as a difference between basal and ouabain-insensitive QO2.
Na+-dependent glucose uptake. Glucose uptake was assessed using a nonmetabolizable glucose analog, MGP, as described previously (34). In brief, confluent monolayers of MPTC were washed with 37°C glucose-free culture medium and incubated (with shaking) in glucose-free medium containing 1 mM MGP and 0.2 µCi/ml of [14C]MGP for 30 min at 37°C in a 5% CO2-95% air atmosphere. Following washing, the amount of [14C]MGP associated with the renal proximal tubular cell monolayer was determined by liquid scintillation spectrometry.
Cisplatin treatment of MPTC. MPTC monolayers reached confluence within 6 days and were treated with cisplatin on day 7 of culture. Samples of MPTC were taken at various time points (between 4 and 18 h) of cisplatin exposure. Caspase inhibitors DEVD-fmk and Z-VAD-fmk were added 0.5 h before cisplatin treatment. Control MPTC were treated with diluent (DMSO; 0.1% final concentration).
Measurement of caspase activities. Caspase-3-, caspase-8-, and caspase-9-like activities were quantified by fluorometric detection of AFC after cleavage from DEVD-AFC, IETDAFC, and LEHD-AFC, respectively. In brief, media were aspirated from culture dishes, MPTC monolayers were washed twice with ice-cold PBS, scraped from the culture dishes, resuspended and lysed in cell lysis buffer (10 min on ice, BioVision, Palo Alto, CA), and centrifuged at 15,000 g for 10 min at 4°C. The pellet was discarded, and the supernatant was used for caspase assays. The supernatant samples were incubated for 1 h at 37°C in the presence of the reaction buffer optimized for caspase activity assays (BioVision), 10 mM dithiothreitol, and 50 µM DEVD-AFC (for measurements of caspase-3-like activity), 50 µM IETD-AFC (for measurement of caspase-8-like activity), or 50 µM LEHD-AFC (for measurement of caspase-9-like activity). The samples were read in a fluorometer at 380/500 nm (excitation/emission), and the amount of product cleaved under linear conditions was determined from the AFC standard curve.
Immunoblotting. Immunoblot analysis was used for the measurement
of protein levels of p21 and cleaved (active) caspase-3, caspase-8, and
caspase-9 in renal proximal tubular cell homogenates. Renal proximal tubular
cell homogenates were lysed and boiled for 10 min in Laemmli sample buffer (60
mM Tris·HCl, pH 6.8, containing 2% SDS, 10% glycerol, 100 mM
-mercaptoethanol, and 0.01% bromophenol blue)
(18), and proteins were
separated using SDS-PAGE. Following electroblotting of the proteins to a
nitrocellulose membrane, blots were blocked for 1 h in 50 mM Tris-buffered
saline (pH 7.5) containing 0.5% casein and 0.1% Tween 20 (blocking buffer) and
incubated overnight at 4°C in the presence of primary antibodies diluted
in the blocking buffer. Following washing with 50 mM Tris-buffered saline
containing 0.05% Tween 20, the membranes were incubated for 1 h with
anti-rabbit or anti-mouse IgG coupled to horseradish peroxidase and washed
again. The supersignal chemiluminescent system (Pierce, Rockford, IL) was used
for protein detection, and scanning densitometry was utilized for the
quantification of results.
Assessment of oncosis. The release of LDH into the culture medium was measured as a marker of oncosis, as described previously (31). Oncosis was also assessed by measuring the permeability of the MPTC plasma membrane to ethidium homodimer. As plasma membrane permeability increases, ethidium homodimer enters the cell and binds to DNA, producing a bright red fluorescence. After 24 h of cisplatin exposure, MPTC monolayers were incubated with 25 µM ethidium homodimer for 15 min on ice, and cells were processed for fluorescence analysis. In brief, media were aspirated and MPTC were scraped from the culture dishes, washed twice with ice-cold PBS, and resuspended in PBS. Cell-associated fluorescence was analyzed by flow cytometry (BD FACSCalibur) using excitation at 488 nm and emission at 590 nm. For each sample, 10,000 events were counted.
Assessment of apoptosis. Apoptosis was assessed using two dissimilar markers: plasma membrane phosphatidylserine externalization measured by annexin V/propidium iodide binding assay and chromatin condensation and nuclear fragmentation measured by immunocytochemistry. MPTC were washed twice with a binding buffer consisting of (in mM) 10 HEPES (pH 7.4), 140 NaCl, 5 KCl, 1 MgCl2, and 1.8 CaCl2. Following resuspension in the binding buffer, MPTC were incubated in the presence of propidium iodide (2 µg/ml) for 15 min on ice, washed three times with the binding buffer, and incubated in the presence of annexin V-FITC (125 ng/ml) for 10 min at room temperature. MPTC were washed three times with the binding buffer and processed for flow cytometry. Propidium iodide and annexin V-FITC fluorescence was quantified by flow cytometry using excitation at 488 nm and emission at 590 and 530 nm for propidium iodide and annexin V-FITC, respectively. For each sample, 10,000 events were counted. Cells positive for annexin V and negative for propidium iodide were considered apoptotic.
MPTC nuclei were visualized by DAPI staining. The monolayers were fixed in 3.7% formaldehyde for 15 min, rinsed with PBS, and incubated with 8 µM DAPI for 2 h at room temperature. Following staining, MPTC monolayers were washed with PBS, coverslips were applied, and nuclei were evaluated using a Zeiss fluorescent microscope (Axioscope). Total and condensed or fragmented nuclei were counted in six to eight different areas of every monolayer using two plates per each experimental group from three independent MPTC isolations.
Biochemical assays. Protein concentration was determined using bicinchoninic acid assay and bovine serum albumin as the standard. Cellular DNA content was determined using a PicoGreen dsDNA Quantitation Kit (Molecular Probes). Monolayers were solubilized in 0.1 M Tris·HCl (pH 7.4) containing 0.15 M NaCl and 0.05% Triton X-100, and DNA was determined in cell lysates according to the manufacturer's protocol.
Statistical analysis. Data are presented as means ± SE and were analyzed for significance using ANOVA. Multiple means were compared using the Student-Newman-Keuls test. Statements of significance were based on P < 0.05. Murine renal proximal tubules isolated on a given day represent a separate experiment (n = 1) consisting of data obtained from two plates.
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RESULTS |
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Protein and DNA contents of confluent MPTC monolayers grown in the improved culture conditions are illustrated in Table 1. Basal and uncoupled QO2 were used as markers of oxidative metabolism and electron transport through the respiratory chain, respectively (Table 1). Both basal and uncoupled QO2 were similar to QO2 in freshly isolated mouse renal proximal tubules (basal QO2: 32 ± 2 nmol O2·min-1·mg protein-1; uncoupled QO2:48 ± 6 nmol O2·min-1·mg protein-1) and rabbit renal proximal tubular cells grown under optimized conditions (34). Ouabain-sensitive QO2 was used as a marker of active Na+ transport in MPTC. Ouabain-sensitive QO2 in confluent cultures of MPTC grown in the improved culture conditions (Table 1) was equivalent to ouabain-sensitive QO2 in freshly isolated mouse renal proximal tubules (14 ± 1 nmol O2·min-1·mg protein-1) and rabbit renal proximal tubular cells grown under optimized conditions (30). Na+-dependent glucose uptake was measured using a nonmetabolizable glucose analog MGP. MGP uptake in confluent quiescent MPTC (Table 1) was similar to MGP uptake in primary cultures of rabbit renal proximal tubular cells (34). Phlorizin (1 mM) inhibited MGP uptake in MPTC by 97 ± 1%. Cellular density, respiration, active Na+ transport, and Na+-dependent glucose uptake in MPTC isolated from mice lacking p21WAF1/CIP1 were equivalent to those in MPTC from p21(+/+) mice (Table 1). These data show that primary cultures of MPTC grown in optimized conditions maintain in vivo-like respiration and active Na+ transport of renal proximal tubules in the presence and absence of p21WAF1/CIP1.
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Cisplatin exposure and p21WAF1/CIP1 protein levels. Protein levels of p21WAF1/CIP1 in MPTC from p21(+/+) mice increased twofold at 4 h of exposure to 15 µM cisplatin and remained increased until the end of the treatment (18 h) (Fig. 1A). Caspase inhibitors had no effect on cisplatin-induced increases in p21 levels (Fig. 1A). p21WAF1/CIP1 protein was absent in control and cisplatin-treated MPTC from p21(-/-) mice (Fig. 1B).
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Cisplatin-induced caspase activation in p21(+/+) MPTC. Caspase activation in MPTC was assessed by measuring caspase activities and evaluating the formation of caspase cleavage products during cisplatin exposure.
Caspase-3, caspase-8, and caspase-9 activities were assessed in MPTC after cisplatin (5, 10, and 15 µM) exposure and in MPTC treated with the vehicle (0.1% DMSO, controls). No significant changes in caspase-3 activity were observed in MPTC treated with DMSO. Caspase-3 activity was not altered during the first 8 h of cisplatin treatment in MPTC (Fig. 2A). At 12 h, caspase-3 activity was unaffected in MPTC treated with 5 and 10 µM cisplatin but increased fivefold in MPTC incubated with 15 µM cisplatin (Fig. 2A). At 18 h, caspase-3 activity increased 11-fold in MPTC incubated with 10 µM cisplatin and was 22-fold higher in MPTC treated with 15 µM cisplatin (Fig. 2A). Immunoblot analysis showed the presence of significant amounts of the cleaved (active) caspase-3 (17-20 kDa) in MPTC treated with 15 µM cisplatin for 18 h (Fig. 2C). The caspase inhibitors DEVD-fmk and ZVAD-fmk decreased caspase-3 processing during 15 µM cisplatin exposure (Fig. 2C).
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To elucidate whether caspase-8 and caspase-9 play a role in cisplatin-induced caspase-3 activation in MPTC, we examined the time course of the activation of caspase-8 and caspase-9 during cisplatin exposure compared with the activation of caspase-3. There was no significant change in caspase-8 activity during 12 h of exposure at any concentration of cisplatin, whereas caspase-3 activity was markedly increased (Figs. 2A and 3A). Cisplatin at 10 and 15 µM increased caspase-8 activity two- and threefold, respectively, after 18 h of treatment (Fig. 3A). Nevertheless, immunoblot analysis showed no cleavage products of caspase-8 (10 kDa) in MPTC exposed to 15 µM cisplatin for 18 h (Fig. 3C). These results show that caspase-8 activation either does not occur in cisplatin-treated MPTC or caspase-8 activation by cisplatin is minor and follows caspase-3 activation.
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No activation of caspase-9 occurred during an 18-h exposure of MPTC to 5 and 10 µM cisplatin (Fig. 4A). An 18-h treatment with 15 µM cisplatin increased MPTC caspase-9 activity 2.3-fold (Fig. 4A). However, immunoblot analysis demonstrated only the presence of procaspase-9 (47 kDa) and no cleaved caspase-9 (35 and 37 kDa) in MPTC exposed to 15 µM cisplatin for 18 h (Fig. 4C). Based on the caspase activity and immunblot assays, we conclude that cisplatin activates caspase-3 without activation of caspase-9 and caspase-8 in p21(+/+) MPTC.
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Cisplatin-induced caspase activation in p21(-/-) MPTC. Exposure to 15 µM cisplatin for 8 h resulted in a threefold increase in caspase-3 activity in p21(-/-) MPTC, whereas no activation of caspase-3 occurred at this time point in p21(+/+) MPTC (Fig. 2, A and B). At 12 h of treatment, caspase-3 activity increased fourfold and sixfold in the presence of 10 and 15 µM cisplatin, respectively (Fig. 2B). Caspase-3 activity continued to increase and, at 18 h of exposure, was 14-fold higher in cells treated with 15 µM cisplatin than in controls and was threefold higher than in p21(+/+) MPTC (Fig. 2, A and B). Immunoblot analysis demonstrated the presence of the cleaved form of caspase-3 at 12 h of 15 µM cisplatin exposure in p21(-/-) MPTC and a further increase in protein levels of the active caspase-3 at 18 h of cisplatin treatment (Fig. 2D). Similar to p21(+/+) MPTC, the caspase inhibitors DEVD-fmk and ZVAD-fmk decreased caspase-3 cleavage during cisplatin exposure in p21(-/-) MPTC (Fig. 2D).
No increase in caspase-8 activity was observed in p21(-/-) MPTC during the first 8 h of cisplatin treatment (Fig. 3B). After 12 h of exposure to 15 µM cisplatin, caspase-8 activity increased threefold in p21(-/-) MPTC, whereas no activation of caspase-8 occurred in p21(+/+) MPTC at this time point (Fig. 3, A and B). At 18 h of treatment, caspase-8 activity in p21(-/-) MPTC treated with 15 µM cisplatin was fourfold higher than in respective controls and threefold higher than in p21(+/+) MPTC undergoing the same treatment (Fig. 3, A and B). However, immunoblot analysis showed no cleaved (active) form of caspase-8 in p21(-/-) MPTC exposed to 15 µM cisplatin for 18 h (Fig. 3D).
No increase in caspase-9 activity occurred during the first 8 h of cisplatin treatment in p21(-/-) MPTC (Fig. 4B). After 12 h of cisplatin (15 µM) exposure, caspase-9 activity increased sevenfold in p21(-/-) MPTC, whereas no increase in caspase-9 activity occurred in p21(+/+) MPTC (Fig. 4, A and B). At 18 h of treatment, caspase-9 activity in p21(-/-) MPTC treated with 15 µM cisplatin was eightfold higher than in controls and twofold higher than in p21(+/+) MPTC undergoing the same treatment (Fig. 4, A and B). In contrast to p21(+/+) MPTC, immunoblot analysis demonstrated the presence of cleaved forms of caspase-9 (35 and 37 kDA) in p21 (-/-) MPTC starting at 4 h of the cisplatin exposure (Fig. 4D). The caspase inhibitors DEVD-fmk and ZVAD-fmk did not block cisplatin-induced caspase-9 processing in p21(-/-) MPTC (Fig. 4D). These results demonstrate that the lack of p21WAF1/CIP1 results in the acceleration and greater extent of activation of caspase-3 and the activation caspase-9 by cisplatin.
Assessment of cisplatin-induced MPTC apoptosis by annexin V/propidium iodide binding. Cisplatin-induced MPTC apoptosis was assessed using annexin V binding as a marker of phosphatidylserine externalization. Cisplatin (15 µM) induced 18 ± 6 and 25 ± 9% apoptosis in p21(+/+) MPTC at 12 and 18 h of exposure, respectively (Fig. 5A). The caspase inhibitors DEVD-fmk and ZVAD-fmk had no effect on the number of annexin V-positive cells at 12 or 18 h of cisplatin exposure (Fig. 5A). These results suggest that caspase inhibition does not block cisplatin-induced apoptosis in MPTC.
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Annexin V binding in p21(-/-) MPTC was 26 ± 9 and 30 ± 9% at 12 and 18 h of cisplatin treatment, respectively, and was caspase inhibitor independent (Fig. 5B). Furthermore, the apoptosis in control p21(-/-) MPTC (i.e., spontaneous apoptosis) was increased 1.4-fold at both 12- and 18-h time points compared with control p21(+/+) MPTC (Fig. 5). These results show that the lack of p21 accelerates cisplatin-induced caspase-independent apoptosis in MPTC.
LDH release was 4 ± 2% in p21(+/+) MPTC and 4 ± 1% in p21(-/-) MPTC after 24 h of cisplatin (15 µM) exposure and was not different from LDH release in controls (1 ± 1%). Uptake of ethidium homodimer in MPTC treated with 15 µM cisplatin for 24 h (4 ± 1%) was not different from that in controls (3 ± 1%). These results show that the plasma membrane was not compromised by cisplatin exposure and support the conclusion that cisplatin induced apoptosis in MPTC in the absence of oncosis.
Assessment of cisplatin-induced MPTC chromatin condensation and nuclear fragmentation. In p21(+/+) MPTC, nuclear changes occurred in 32 ± 1% of the cells treated with 15 µM cisplatin for 18 h (Figs. 6B and 7A), an amount significantly greater (45 ± 2%) than in p21(-/-) MPTC (Figs. 6F and 7B). Monolayers pretreated with DEVD-fmk or 50 µM ZVAD-fmk before cisplatin (15 µM) treatment and caspase activities and chromatin condensation were assessed after 18 h of exposure. DEVD-fmk and ZVAD-fmk prevented the increase in caspase-3 activity and procaspase-3 cleavage in cisplatin-treated MPTC regardless of the presence or absence of p21WAF1/CIP1 (Fig. 2, A and B). Similarly, ZVAD-fmk and DEVD-fmk abolished cisplatin-induced increases in LEHD-ase (Fig. 4, A and B) and IETD-ase (Fig. 3, A and B) activities in both p21(+/+) and p21(-/-) MPTC. However, neither DEVD-fmk nor ZVAD-fmk blocked procaspase-9 processing in p21(-/-) MPTC treated with cisplatin (Fig. 4D). These data suggest that blocking caspase-3 activation in cisplatin-treated MPTC prevents increases in caspase-8 and caspase-9 activities and support our conclusion that increases in caspase-8-like and caspase-9-like activities are secondary to caspase-3 activation.
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Treatment of MPTC with DEVD-fmk and ZVAD-fmk for 18 h had no effect on nuclear morphology in control MPTC (Figs. 5, A and B, and 7, A and B). Pretreatment of p21(+/+) MPTC with DEVD-fmk or ZVAD-fmk before cisplatin exposure had no effect on the number of cells with chromatin condensation and nuclear fragmentation (Figs. 6, C and D, and 7A). Similarly, pretreatment with both inhibitors did not prevent cisplatin-induced MPTC shrinkage and detachment from the monolayer (data not shown). These data suggest that cisplatin-induced apoptosis in p21(+/+) MPTC is caspase independent.
Pretreatment of p21(-/-) MPTC with DEVD-fmk had no significant effect on
cisplatin-induced chromatin condensation and nuclear fragmentation despite the
decrease in caspase-3 processing and complete inhibition of caspase-3 activity
(Figs. 2, B and
D,
6G, and
7B). However,
pretreatment with ZVAD-fmk before cisplatin exposure decreased nuclear
condensation in p21(-/-) MPTC 51% at 18 h (Figs.
6H and
7B). These results
suggest that, in addition to caspase-independent apoptosis, another
mechanism(s) is responsible for the increased cisplatin sensitivity of
p21(-/-) MPTC. The other mechanism is caspase dependent, but the pathway is
independent of caspase-3, -8, and -9 activation.
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DISCUSSION |
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The lack of p21WAF1/CIP1 is associated with the acceleration of caspase activation and apoptosis. Caspase proteases play an important role in apoptosis by degrading specific structural, regulatory, and DNA repair proteins within the cell. Caspase-3 is one of the key executioners of apoptosis and is responsible for the proteolytic cleavage of proteins essential for cell survival. The best known pathways of caspase-3 activation are those through proteolytic cleavage of the caspase-3-inactive zymogen by initiator caspases, caspase-8 and caspase-9. Caspase-8 is recruited after activation of receptors belonging to a family of "death receptors." Caspase-9 is activated as a result of mitochondrial dysfunction and release of cytochrome c from the mitochondrial intermembrane space. Activated caspase-8 and caspase-9 initiate the proteolytic activity of other downstream caspases, including caspase-3 and caspase-6. Therefore, the activation of caspase-8 or caspase-9 precedes activation of caspase-3 during caspase-8- or caspase-9-initiated apoptosis. While cisplatin-induced apoptosis in MPTC is associated with activation of caspase-3, caspase-8 is not cleaved to the enzymatically active form despite increases in IETD-ase activity that accompany apoptosis in MPTC in the presence or absence of p21WAF1/CIP1. Therefore, we conclude that 1) the increases in IETD-ase activity were not due to caspase-8, 2) caspase-8 is not activated by cisplatin in MPTC, and 3) caspase-3 is not activated by caspase-8.
Similarly, our data show that caspase-9 was not processed to the active form in p21(+/+) MPTC treated with cisplatin. Therefore, the increases in LEHD-ase activity at 18 h of cisplatin exposure were not due to caspase-9 activation and caspase-3 activation occurred without caspase-9. In contrast, caspase-9 was cleaved in cisplatin-treated p21(-/-) MPTC and caspase-9 processing preceded caspase-3 activation. These results suggest the sequential activation of caspase-9 and caspase-3 in p21(-/-) MPTC but not in p21(+/+) MPTC after cisplatin exposure. These data are somewhat in contrast to the results of Kaushal and colleagues (14), who demonstrated the activation of caspase-8, caspase-9, and caspase-3 by cisplatin in LLC-PK1 cells. This discrepancy may be due to differences between immortalized cell lines and primary cultures, including differences in cellular metabolism. It is also possible that caspase-8 and caspase-9 in wild-type MPTC are activated late during cisplatin-induced apoptosis (>18 h of exposure). However, similar to our results, Kaushal et al. reported that the activation of caspase-3 preceded that of caspase-8 and caspase-9 and concluded that caspase-3 activation may be initiated by a caspase-independent pathway or caspases other than caspase-8 and caspase-9.
Our results suggest that caspase-3 activation by cisplatin in MPTC may be partially regulated through a p21WAF1/CIP1-dependent mechanism because the lack of functional p21WAF1/CIP1 protein accelerates caspase-3 activation. This is in agreement with reports that p21WAF1/CIP1 overexpression suppresses caspase activation and apoptosis (47, 48). However, despite caspase-3 activation during cisplatin-induced apoptosis, caspase-3 does not appear to be involved in the execution of apoptosis in either p21(+/+) MPTC or p21(-/-) MPTC. This conclusion is based on the observation that a caspase-3 inhibitor, DEVD-fmk, blocked caspase-3 activation but did not prevent MPTC phosphatidylserine externalization, chromatin condensation, and cell shrinkage (Figs. 5 and 7; data not shown). Therefore, we conclude that caspase-3 is activated by cisplatin but is not required for cisplatin-induced apoptosis in MPTC. Caspase-3-independent apoptosis has been reported in other models, including X-irradiation-induced apoptosis in rat embryonic fibroblasts (16) and cisplatin-induced apoptosis in rabbit primary cultures of renal proximal tubular cells (4, 32). Furthermore, genetic deletion of caspase-3 and caspase-9 does not alter apoptosis during embryogenesis and fetal development in mice, further supporting the existence of caspase-3- and caspase-9-independent mechanisms of apoptosis (35). Apoptosis induced by a variety of stimuli in human breast cancer MCF-7 cells bypasses the need for caspase-3 activation and proceeds through the activation of caspase-7 and caspase-6 (24).
We determined whether other caspases mediate cisplatin-induced apoptosis in MPTC and whether p21WAF1/CIP1 plays a role in these events. Our results demonstrate that the broad spectrum caspase inhibitor ZVAD-fmk had no protective effects against cisplatin-induced phosphatidylserine externalization, chromatin condensation, and cell shrinkage in MPTC expressing functional p21WAF1/CIP1 despite completely inhibiting caspase-3 activation. These observations suggest that caspases do not play a role in apoptosis in MPTC expressing p21WAF1/CIP1. In contrast, in cisplatin-treated MPTC lacking p21WAF1/CIP1, ZVAD-fmk decreased cell shrinkage (data not shown) and nuclear fragmentation by 51% but did not offer any protection against phosphatidylserine externalization. These results suggest that the lack of p21WAF1/CIP1 results in the activation of a ZVAD-fmk-sensitive caspase that accelerates and potentiates cisplatin-induced chromatin and nuclear condensation but not phosphatidylserine externalization. Because caspase-9 is activated by cisplatin in MPTC lacking p21WAF1/CIP1 and inhibited by ZVAD-fmk, mitochondria-dependent activation of caspase-9 may be involved in mediating nuclear changes. Furthermore, these results suggest that phosphatidylserine externalization and nuclear condensation are mediated by different mechanisms. However, ZVAD-fmk provided only 50% protection against cell shrinkage and nuclear fragmentation, which suggests that 50% of cisplatin-induced apoptosis in MPTC lacking p21WAF1/CIP1 is caspase independent. In contrast, cisplatin-induced apoptosis in MPTC expressing functional p21WAF1/CIP1 was entirely caspase independent.
An alternate mechanism for the antiapoptotic effect of p21WAF1/CIP1 may be through the inhibition of the activation of the cyclin A-cyclin-dependent kinase 2 (Cdk2) complex (13). DNA damage by cisplatin and cell cycle progression in injured cells is blocked by the binding of functional p21WAF1/CIP1 to the cyclin A-Cdk2 complex and inhibition of Cdk2 activity. Caspase-3-mediated cleavage of p21WAF1/CIP1 prevents inhibition of the cyclin A-Cdk2 complex and leads to apoptosis (13). In kidney cells, the lack of p21WAF1/CIP1 leads to progression from the G1 to S phase of the cell cycle after cisplatin-induced DNA damage and cell injury in vivo (28). Similarly, in primary culture of MPTC, the lack of p21WAF1/CIP1 increased the number of cells in S phase [17% in p21(-/-) MPTC vs. 3% in p21(+/+) MPTC] and G2/M phase [15% in p21(-/-) MPTC vs. 2% in p21(+/+) MPTC] of the cell cycle at 18 h of cisplatin exposure (data not shown). In contrast, overexpression of p21WAF1/CIP1 or transfection of cells with a p21WAF1/CIP1 mutant resistant to caspase-3 cleavage suppresses cyclin A-Cdk2 activity and subsequent apoptosis (13). Caspase inhibitors DEVD-fmk and ZVAD-fmk did not block cisplatin-induced cell cycle progression to G2/M phase in MPTC lacking p21WAF1/CIP1 (data not shown).
In conclusion, this study shows that cisplatin treatment in wild-type MPTC activates caspase-3 through a caspase-8- and caspase-9-independent pathway and induces apoptosis. However, caspase activation is not the mechanism by which cisplatin induces apoptosis in MPTC expressing functional p21WAF1/CIP1. Lack of p21WAF1/CIP1 stimulates caspase-9 activation and accelerates caspase-3 activation in cisplatin-induced MPTC apoptosis, which suggests that p21WAF1/CIP1 functions, in part, through decreasing activation of caspase(s). Cisplatin-induced apoptosis in the absence of p21WAF1/CIP1 is caspase-3 independent but dependent, in part, on other caspase(s). Our study also suggests that some pharmacological agents activate caspases and induce apoptosis but produce nephrotoxicity through mechanisms other than caspase-mediated proteolysis.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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