Lack of a functional p21WAF1/CIP1 gene accelerates caspase-independent apoptosis induced by cisplatin in renal cells

Grazyna Nowak,1 Peter M. Price,2 and Rick G. Schnellmann3

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


    ABSTRACT
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The lack of cyclin-dependent kinase inhibitor p21WAF1/CIP1 (p21) in mice increases renal proximal tubular cell death and enhances sensitivity to acute renal failure produced by the chemotherapeutic agent cisplatin. We used primary cultures of mouse renal proximal tubular cells (MPTC) grown in optimized culture conditions to investigate the cellular basis for increased apoptosis in p21 knockout mice. Cisplatin (15 µM) activated caspase-3 but not caspase-8 or caspase-9 and produced phosphatidylserine externalization, chromatin condensation, and nuclear fragmentation in wild-type [p21(+/+)] MPTC. Caspase-3 activation and apoptosis were accelerated in cisplatin-treated MPTC lacking p21 [p21(-/-) MPTC]. In contrast to p21(+/+) MPTC, cisplatin activated caspase-9 but not caspase-8 in p21(-/-) MPTC before caspase-3 activation. The caspase-3 inhibitor Asp-Glu-Val-Asp-fluoromethylketone (DEVD-fmk) inhibited caspase-3 activity but did not abolish apoptosis in p21(+/+) and p21(-/-) MPTC. General caspase inhibitor Z-Val-Ala-Asp(OCH3)-fluoromethylketone (ZVAD-fmk) inhibited caspase activity and decreased chromatin condensation by 51% in p21(-/-) but not in p21(+/+) MPTC. However, cisplatin-induced phosphatidylserine externalization was not inhibited by ZVAD-fmk in p21(-/-) MPTC. We conclude that 1) in the presence of p21, cisplatin activates caspase-3 through a mechanism independent of caspase-8 or caspase-9; 2) in the absence of p21, caspase-9 activation precedes caspase-3 activation; 3) the lack of p21 accelerates caspase-3 activation and cisplatin-induced MPTC apoptosis; and 4) MPTC apoptosis is caspase independent in the presence of p21 but partially dependent on caspases in the absence of p21.

renal proximal tubular cells; caspase activation; DNA damage; knockout mice


CISPLATIN IS A WIDELY USED anticancer agent in the treatment of many solid tumors and metastatic cancers (6, 19). Cisplatin administration causes cell-cycle arrest at different phases of the cell cycle and induces apoptosis in a variety of cancer cells (6). The major disadvantage of this antineoplastic agent is a dose-dependent and cumulative nephrotoxicity, which manifests primarily as proximal tubule morphological damage (oncosis and apoptosis) and physiological dysfunction (1, 3, 8, 10, 20, 21, 30, 38, 45). Multiple mechanisms have been implicated in cisplatin-induced nephrotoxicity, including inhibition of protein synthesis, DNA damage, oxidative stress, mitochondrial dysfunction, and alterations in signal transduction pathways involved in apoptosis (12, 17, 36, 39, 41, 45, 48). Lower concentrations of cisplatin induce apoptosis, whereas concentrations >50 µM cause oncosis in renal proximal tubular cells (7, 25). Cisplatin-induced apoptosis in this cell type is preceded by activation of caspase-3, but not caspase-1 (3, 4, 8, 32, 44, 48), and is prevented by bcl-2 (48). Activation of caspase-8 and caspase-9 by cisplatin has been reported in the LLC-PK1 cell line (14).

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.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Materials. Cell isolation media, cell culture media, horse serum, and soybean trypsin inhibitor were supplied by Life Technologies (Grand Island, NY). Collagenase type 4 and L-ascorbic acid-2-phosphate magnesium salt were provided by Worthington Biochemical (Freehold, NJ) and Wako Bio-Products (Richmond, VA), respectively. Cisplatin [cis-diamminedichloroplatinum(II)], Percoll, methyl {alpha}-D-glucopyranoside, propidium iodide, and cell culture hormones were purchased from Sigma (St. Louis, MO). Ethidium homodimer and 4',6-diamidino-2-phenylindole dihydrochloride (DAPI) were obtained from Molecular Probes (Eugene, OR). Caspase-3 [Asp-Glu-Val-Asp (DEVD)-7-amino-4-trifluoromethylcoumarin (AFC)] and caspase-8 [Ile-Glu-Thr-Asp (IETD)-AFC] fluorometric substrates, AFC, and buffers for caspase assays were purchased from BioVision (Palo Alto, CA). Caspase-9 substrate [Leu-Glu-His-Asp (LEHD)-AFC] was obtained from Calbiochem (La Jolla, CA). Caspase-3 [DEVD-fluoromethylketone (fmk)] and pan-caspase [Z-Val-Ala-Asp(OCH3)-fmk (ZVAD-fmk)] inhibitors were purchased from R&D Systems (Minneapolis, MN). Antibodies against cleaved forms of caspase-3, caspase-8, and caspase-9 were supplied by Cell Signaling Technology (Beverly, MA), and an anti-p21 antibody was supplied by Santa Cruz Biotechnology (Santa Cruz, CA). Methyl {alpha}-D-[U-14C]glucopyranoside (MGP; specific activity 230 mCi/mmol) was purchased from Amersham Pharmacia Biotech (Piscataway, NJ). The sources of the other reagents have been described previously (34).

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 {beta}-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.


    RESULTS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Metabolic characteristics of primary cultures of MPTC grown in optimized culture conditions. Previously, we showed that primary cultures of rabbit renal proximal tubular cells maintain many in vivo differentiated functions of renal proximal tubules when grown under optimized culture conditions (33, 34). Improving culture media oxygenation (by shaking the dishes) in conjunction with decreasing glucose concentration and supplementing with lactate and ascorbic acid results in a reduction in glycolysis, an in vivo-like respiration, and an increase in transport functions. Therefore, we modified the original culture conditions (33) used to grow MPTC in primary culture to improve the growth, oxidative metabolism, and transport functions in these cells. The modifications included 1) decreasing glucose concentration in the culture media from 5 to 1 mM, 2) supplementing media with 5 mM lactate (a major substrate for the kidney in vivo) and 50 µM ascorbic acid phosphate, and 3) shaking the dishes to improve media oxygenation.

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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Table 1. Metabolic characteristics of primary cultures of MPTC isolated from wild-type and p21WAF1/CIP1 knockout mice and grown for 7 days in optimized culture conditions

 

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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Fig. 1. Effect of cisplatin on protein levels of p21WAF1/CIP1 in primary cultures of mouse renal proximal tubular cells (MPTC) expressing (A) and lacking (B) p21WAF1/CIP1. MPTC were treated with 15 µM cisplatin, and samples were taken at 4, 8, 12, and 18 h of exposure for immunoblot analysis. The experiment was repeated 3 times with similar results. DEVD, Asp-Glu-Val-Asp; fmk, fluoromethylketone; ZVAD, Z-Val-Ala-Asp(OCH3).

 

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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Fig. 2. Effect of cisplatin on caspase-3-like activity in primary cultures of MPTC expressing (A) and lacking (B) p21WAF1/CIP1. MPTC were treated with 0, 5, 10, and 15 µM cisplatin, and samples were taken at 4, 8, 12, and 18 h of cisplatin exposure for measurements of caspase-3-like activity. Control MPTC were treated with vehicle (DMSO) alone. Values are means ± SE of 4-5 experiments. Values with different letters are significantly different (P < 0.05) from each other. C and D: protein levels of cleaved (active) caspase-3 (17-20 kDa) during cisplatin (15 µM) exposure in primary cultures of MPTC expressing (C) and lacking (D) p21WAF1/CIP1. Presented blots are representative of results obtained from 3 independent experiments.

 

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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Fig. 3. Effect of cisplatin on caspase-8-like activity in primary cultures of MPTC expressing (A) and lacking (B) p21WAF1/CIP1. MPTC were treated with 0, 5, 10, and 15 µM cisplatin, and samples were taken at 4, 8, 12, and 18 h of cisplatin exposure for measurements of caspase-8-like activity. Control MPTC were treated with vehicle (DMSO) alone. Values are means ± SE of 3-4 experiments. Values with different letters are significantly different (P < 0.05) from each other. C and D: absence of cleaved (active) caspase-8 (10 kDa) during cisplatin (15 µM) exposure in primary cultures of MPTC expressing (C) and lacking (D) p21WAF1/CIP1. Presented blots are representative of results obtained from 3 independent experiments. C, untreated Jurkat cells; PC, positive control (Jurkat cells treated with etoposide).

 

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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Fig. 4. Effect of cisplatin on caspase-9-like activity in primary cultures of MPTC expressing (A) and lacking (B) p21WAF1/CIP1. MPTC were treated with 0, 5, 10, and 15 µM cisplatin, and samples were taken at 4, 8, 12, and 18 h of cisplatin exposure for measurements of caspase-9-like activity. Control MPTC were treated with vehicle (DMSO) alone. Values are means ± SE of 6 experiments. Values with different letters are significantly different (P < 0.05) from each other. C and D: protein levels of procaspase-9 (top band) and cleaved (active) caspase-9 (bottom bands; 35-37 kDa) during cisplatin (15 µM) exposure in primary cultures of MPTC expressing (C) and lacking (D) p21WAF1/CIP1. Presented blots are representative of results obtained from 3 independent experiments. C, untreated Jurkat cells; PC, positive control (Jurkat cells treated with cytochrome c).

 

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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Fig. 5. Effects of cisplatin (15 µM), DEVD-fmk (50 µM), and ZVAD-fmk (50 µM) on annexin V-FITC binding in primary cultures of MPTC expressing (A) and lacking (B) p21WAF1/CIP1. The graphs show the percentage of annexin V-FITC-positive and propidium iodide (PI)-negative MPTC as determined by flow cytometry. Values are means ± SE of 3 experiments. Values with different letters are significantly different (P < 0.05) from each other.

 

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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Fig. 6. Effect of caspase inhibitors on cisplatin-induced changes in nuclear morphology after 18 h of cisplatin exposure in primary cultures of MPTC. MPTC were pretreated with caspase inhibitors (50 µM DEVD-fmk and 50 µM ZVAD-fmk) for 30 min before cisplatin (15 µM) treatment. Control MPTC were treated with vehicle (DMSO) alone. MPTC were fixed and stained with 4',6-diamidino-2-phenylindole dihydrochloride, and nuclei were evaluated using a fluorescent microscope. A: control [p21(+/+)]. B: cisplatin [p21(+/+)]. C: DEVD-fmk+cisplatin [p21(+/+)]. D: ZVAD-fmk+cisplatin [p21(+/+)]. E: control [p21(-/-)]. F: cisplatin [p21(-/-)]. G: DEVD-fmk+cisplatin [p21(-/-)]. H: ZVAD-fmk+cisplatin [p21(-/-)]. Microphotographs are representative of results obtained from 3 independent experiments. Magnification: x400.

 


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Fig. 7. Effect of caspase inhibitors on the percentage of cells displaying chromatin condensation and nuclear fragmentation after 18 h of cisplatin exposure in primary cultures of MPTC expressing (A) and lacking (B) p21WAF1/CIP1. MPTC were pretreated with caspase inhibitors (50 µM DEVD-fmk and 50 µM ZVAD-fmk) for 30 min before cisplatin (15 µM) treatment. Control MPTC were treated with vehicle (DMSO) alone. See Fig. 6 for sample processing. Values are means ± SE of 3 experiments. Values with different letters are significantly different (P < 0.05) from each other.

 

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.


    DISCUSSION
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
ARF after ischemia or cisplatin administration is associated with a rapid induction of the cell cycle inhibitory protein p21WAF1/CIP1 gene and overexpression of p21WAF1/CIP1 protein (26, 29). p21WAF1/CIP1 induction is associated with cell-cycle interruption and arrest in the G1 phase of the cell cycle (46). Induction of p21WAF1/CIP1 after cisplatin administration has a protective effect on the murine kidney as mice lacking p21WAF1/CIP1 display a more rapid onset of ARF and have a higher mortality rate than wild-type animals (27). The proximal tubule segment of the nephron is the most sensitive to cisplatin and undergoes severe damage in response to cisplatin-based chemotherapy (10). In vivo studies demonstrated that proximal tubular cells in mice lacking p21WAF1/CIP1 exhibited greater cell death than proximal tubular cells in wild-type mice (28). Our in vitro studies show that low concentrations of cisplatin cause cell death by apoptosis, but not oncosis, and that the lack of p21WAF1/CIP1 increases the extent of apoptosis by 40% in primary cultures of MPTC. Thus our results suggest that p21WAF1/CIP1 plays a protective role by decreasing cisplatin-induced cell death. Furthermore, our data demonstrate that the increase in apoptosis in mice lacking p21WAF1/CIP1 is the result of a direct effect of cisplatin on the renal proximal tubular cells.

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.


    ACKNOWLEDGMENTS
 
We thank Dr. Philip Leder (Harvard Medical School, Boston, MA) for providing several heterozygous mice carrying the p21WAF1/CIP1 gene deletion and for providing a probe for screening and Dr. Bert Vogelstein (Johns Hopkins Oncology Center, Baltimore, MD) for cloned mouse p21WAF1/CIP1 cDNA. We also thank Ashley B. Whitlow (UAMS, Little Rock, AR) for assistance with flow cytometry.


    FOOTNOTES
 

Address for reprint requests and other correspondence: G. Nowak, Dept. of Pharmaceutical Sciences, Univ. of Arkansas for Medical Sciences, 4301 West Markham St., Little Rock, AR 72205 (E-mail: gnowak{at}uams.edu).

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.


    REFERENCES
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Brady HR, Kone BC, Stromski ME, Zeidel M, Giebisch G, and Gullans SR. Mitochondrial injury: an early event in cisplatin toxicity to renal proximal tubules. Am J Physiol Renal Fluid Electrolyte Physiol 258: F1181-F1187, 1990.[Abstract/Free Full Text]
  2. Brugarolas J, Chandrasekaran C, Gordon JI, Beach D, Jacks T, and Hannon GJ. Radiation-induced cell cycle arrest compromised by p21 deficiency. Nature 377: 552-557, 1995.[ISI][Medline]
  3. Chopra S, Kauffman JS, Jones TW, Hong WK, Gehr MK, Hamburger RJ, Flamenbaum W, and Trump GF. Cis-diamminedichlorplatinum-induced acute renal failure in the rat. Kidney Int 21: 54-64, 1982.[ISI][Medline]
  4. Cummings BS and Schnellmann RG. Cisplatin-induced renal cell apoptosis: caspase 3-dependent and -independent pathways. J Pharmacol Exp Ther 302: 8-17, 2002.[Abstract/Free Full Text]
  5. Deng C, Zhang P, Harper JW, Elledge SJ, and Leder P. Mice lacking p21WAF1/CIP1 undergo normal development, but are defective in G1 checkpoint control. Cell 82: 675-684, 1995.[ISI][Medline]
  6. Eastman A. Activation of programmed cell death by anticancer agents: cisplatin as a model system. Cancer Cells 2: 275-280, 1990.[ISI][Medline]
  7. Fan S, Chang JK, Smith ML, Duba D, Fornace AJ Jr, and O'Connor PM. Cells lacking CIP1/WAF1 genes exhibit preferential sensitivity to cisplatin and nitrogen mustard. Oncogene 14: 2127-2136, 1997.[ISI][Medline]
  8. Fan S, Smith ML, Rivet DJ, Duba D, Zhan Q, Kohn KW, Fornace AJ Jr, and O'Connor PM. Disruption of p53 function sensitizes breast cancer MCF-7 cells to cisplatin and pentoxifylline. Cancer Res 55: 1649-1654, 1995.[Abstract]
  9. Glaser T, Wagenknecht B, and Weller M. Identification of p21 as a target of cycloheximide-mediated facilitation of CD95-mediated apoptosis in human malignant glioma cells. Oncogene 20: 4757-4767, 2001.[ISI][Medline]
  10. Goldstein RS and Mayor GH. Minireview. The nephrotoxicity of cisplatin. Life Sci 32: 685-690, 1983.[ISI][Medline]
  11. Gorospe M, Wang X, and Holbrook NJ. Functional role of p21 during the cellular response to stress. Gene Express 7: 377-385, 1999.[ISI]
  12. Hanigan MH, Gallagher BC, Taylor PT Jr, and Large MK. Inhibition of gamma-glutamyl transpeptidase activity by acivicin in vivo protects the kidney from cisplatin-induced toxicity. Cancer Res 54: 5925-5929, 1994.[Abstract]
  13. Jin YH, Yoo KJ, Lee SK, and Lee SK. Caspase-3 mediated cleavage of p21WAF1/CIP1 associated with the cyclin A-cyclin-dependent kinase 2 complex is a prerequisite for apoptosis in SK-HEP-1 cells. J Biol Chem 275: 3025-3063, 2000.[Abstract/Free Full Text]
  14. Kaushal GP, Kaushal V, Hong X, and Shah SV. Role and regulation of activation of caspases in cisplatin-induced injury to renal tubular epithelial cells. Kidney Int 60: 1726-1736, 2002.[ISI]
  15. Kim SG, Kim SN, Jong HS, Kim NK, Hong SH, Kim SJ, and Bang YJ. Caspase-mediated Cdk2 activation is a critical step to execute transforming growth factor-{beta}1-induced apoptosis in human gastric cancer cells. Oncogene 20: 1254-1265, 2001.[ISI][Medline]
  16. Kobayashi D, Tokino T, and Watanabe N. Contribution of caspase-3 differs by p53 status in apoptosis induced by X-irradiation. Jpn J Cancer Res 92: 475-481, 2001.[ISI][Medline]
  17. Kruidering M, van de Water B, Zhan Y, Baelde JJ, de Heer E, Mulder GJ, Stevens JL, and Nagelkerke JF. Cisplatin effects on F-actin and matrix proteins precede renal tubular cell detachment and apoptosis in vitro. Cell Death Diff 5: 601-614, 1998.[ISI][Medline]
  18. Laemmli UK. Cleavage of structural proteins during the assembly of the head bacteriophage T4. Nature 227: 680-685, 1970.[ISI][Medline]
  19. Lau AH. Apoptosis induced by cisplatin nephrotoxic injury. Kidney Int 56: 1295-1298, 1999.[ISI][Medline]
  20. Leibrandt MEI, Wolfgang GHI, Metz AL, Ozobia AA, and Haskins JR. Critical subcellular targets of cisplatin and related platinum analogs in rat renal proximal tubule cells. Kidney Int 48: 761-770, 1995.[ISI][Medline]
  21. Levi J, Jacobs C, Kalman SB, Mctigue M, and Weiner MW. Mechanism of cis-platinum nephrotoxicity. I. Effects of sulfhydryl groups in rat kidneys. J Pharmacol Exp Ther 213: 545-550, 1980.[Abstract]
  22. Li WW, Fan J, Hochhauser D, and Bertino JR. Overexpression of p21waf1 leads to increased inhibition of E2F-1 phosphorylation and sensitivity to anticancer drugs in retinoblastoma-negative human sarcoma cells. Cancer Res 57: 2193-2199, 1997.[Abstract]
  23. Li Y, Jenkins CW, Nichols MA, and Xiong Y. Cell cycle expression and p53 regulation of the cyclin-dependent kinase inhibitor p21. Oncogene 9: 2261-2268, 1994.[ISI][Medline]
  24. Liang Y, Yan C, and Schor NF. Apoptosis in the absence of caspase 3. Oncogene 20: 6570-6578, 2001.[ISI][Medline]
  25. Lieberthal W, Triaca V, and Levine J. Mechanisms of death induced by cisplatin in proximal tubular epithelial cells: apoptosis vs. necrosis. Am J Physiol Renal Fluid Electrolyte Physiol 270: F700-F708, 1996.[Abstract/Free Full Text]
  26. Megyesi J, Andrade L, Vieira JM Jr, Safirstein RL, and Price PM. Positive effect of the induction of p21WAF1/CIP1 on the course of ischemic acute renal failure. Kidney Int 60: 2164-2172, 2001.[ISI][Medline]
  27. Megyesi J, Price PM, Tamayo E, and Safirstein RL. The lack of p21WAF1/CIP1 gene ameliorates progression to chronic renal failure. Proc Natl Acad Sci USA 96: 10830-10835, 1999.[Abstract/Free Full Text]
  28. Megyesi J, Safirstein RL, and Price PM. Induction of p21WAF1/CIP1/SDI1 in kidney tubule cells affects the course of cisplatin-induced acute renal failure. J Clin Invest 101: 777-782, 1998.[Abstract/Free Full Text]
  29. Megyesi J, Udvarhelyi N, Safirstein RL, and Price PM. The p53-independent activation of transcription of p21WAF1/CIP1/SDI1 after acute renal failure. Am J Physiol Renal Fluid Electrolyte Physiol 271: F1211-F1216, 1996.[Abstract/Free Full Text]
  30. Miura K, Goldstein RS, Pasino DA, and Hook JB. Cisplatin nephrotoxicity: role of filtration and tubular transport of cisplatin in isolated perfused kidneys. Toxicology 44: 147-158, 1987.[ISI][Medline]
  31. Moran JH and Schnellman RG. A rapid beta-NADH-linked fluorescence assay for lactate dehydrogenase in cellular death. J Pharm Toxicol Methods 36: 41-44, 1996.[ISI][Medline]
  32. Nowak G. PKC-{alpha} and ERK1/2 mediate mitochondrial dysfunction, decreases in active Na+ transport, and cisplatin-induced apoptosis in renal cells. J Biol Chem 277: 43377-43388, 2002.[Abstract/Free Full Text]
  33. Nowak G and Schnellmann RG. Improved culture conditions stimulate gluconeogenesis in primary cultures of renal proximal tubule cells. Am J Physiol Cell Physiol 268: C1053-C1061, 1995.[Abstract/Free Full Text]
  34. Nowak G and Schnellmann RG. L-Ascorbic acid regulates growth and metabolism of renal cells: improvements in cell culture. Am J Physiol Cell Physiol 271: C2072-C2080, 1996.[Abstract/Free Full Text]
  35. Oppenheim RW, Flavell RA, Vinsant S, Prevette D, Kuan CY, and Rakic P. Programmed cell death of developing mammalian neurons after genetic deletion of caspases. J Neurosci 21: 4752-4760, 2001.[Abstract/Free Full Text]
  36. Osman AM, El-Sayed EM, El-Demerdash E, Al-Hyder A, El-Didi M, Attia AS, and Hamada FM. Prevention of cisplatin-induced nephrotoxicity by methimazole. Pharm Res 41: 115-121, 2000.
  37. Qin LF and Ng IOL. Exogenous expression of p21WAF1/CIP1 exerts cell growth inhibition and enhances sensitivity to cisplatin in hepatoma cells. Cancer Lett 172: 7-15, 2001.[ISI][Medline]
  38. Safirstein RL, Winston J, Moel D, Dikman S, and Guttenplan J. Cisplatin nephrotoxicity: insights into mechanism. Int J Androl 10: 325-346, 1987.[ISI][Medline]
  39. Sener G, Satiroglu H, Kabasakal L, Arbak S, Oner S, Ercan F, and Keyer-Uysal M. The protective effect of melatonin on cisplatin nephrotoxicity. Fund Clin Pharmacol 14: 553-560, 2000.[ISI][Medline]
  40. Sheridan AM, Schwartz JH, Kroshian VM, Tercyak AM, Laraia J, Masino S, and Lieberthal W. Renal mouse proximal tubular cells are more susceptible than MDCK cells to chemical anoxia. Am J Physiol Renal Fluid Electrolyte Physiol 265: F342-F350, 1993.[Abstract/Free Full Text]
  41. Somani SM, Husain K, Whitworth C, Trammell GL, Malafa M, and Rybak LP. Dose-dependent protection by lipoic acid against cisplatin-induced nephrotoxicity in rats: antioxidant defense system. Pharmacol Toxicol 86: 234-241, 2000.[ISI][Medline]
  42. Suzuki A, Tsutomi Y, Akahane K, Araki T, and Miura M. Resistance to Fas-mediated apoptosis: activation of caspase-3 is regulated by cell cycle regulator p21WAF1 and IAP gene family ILP. Oncogene 17: 931-939, 1998.[ISI][Medline]
  43. Suzuki A, Tsutomi Y, Miura M, and Akahane K. Mitochondrial regulation of cell death: mitochondria are essential for procaspase 3-p21 complex formation to resist Fas-mediated cell death. Oncogene 18: 1239-1244, 1999.[ISI][Medline]
  44. Van de Water B, Nagelkerke JF, and Stevens JL. Dephosphorylation of focal adhesion kinase (FAK) and loss of focal contacts precede caspase-mediated cleavage of FAK during apoptosis in renal epithelial cells. J Biol Chem 274: 13328-13337, 1999.[Abstract/Free Full Text]
  45. Van de Water B, Tijdens IB, Verbrugge A, Huisloot M, Dihal AA, Stevens JL, Jaken S, and Mulder GJ. Cleavage of the actin-capping protein a-adducin at Asp-Asp-Ser-Asp633-Ala by caspase-3 is preceded by its phosphorylation on serine 726 in cisplatin-induced apoptosis of renal epithelial cells. J Biol Chem 275: 25805-25813, 2000.[Abstract/Free Full Text]
  46. Waldman T, Kinzler KW, and Vogelstein B. p21 Is necessary for the p53-mediated G1 arrest in human cancer cells. Cancer Res 55: 5187-5190, 1995.[Abstract]
  47. Xu SQ and El-Diery WS. p21(WAF1/CIP1) inhibits initiator caspase cleavage by TRAIL death receptor DR4. Biochem Biophys Res Commun 269: 179-190, 2000.[ISI][Medline]
  48. Zhan Y, van der Water B, Wang Y, and Stevens JL. The roles of caspase-3 and bcl-2 in chemically induced apoptosis but not necrosis of renal epithelial cells. Oncogene 18: 6505-6512, 1999.[ISI][Medline]
  49. Zhang H, Hannon GJ, and Beach D. p21-Containing cyclin kinases exist in both active and inactive states. Genes Dev 8: 1750-1758, 1994.[Abstract]