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【摘要】 Cisplatin induces renal cell injury and death, resulting in nephrotoxicity that limits its use in cancer therapy. Using cell culture models, recent work has suggested the involvement of p53 in renal cell apoptosis during cisplatin treatment. However, the signals upstream of p53 remain elusive. ATM and ATR are critical regulators of p53 under various conditions of DNA damage. Here, we show that ATM, and not ATR, was proteolytically cleaved into specific fragments of 210 and 150 kDa during cisplatin-induced tubular cell apoptosis. ATM cleavage was paralleled by the development of apoptosis. VAD, a broad-spectrum inhibitor of caspases, attenuated the cleavage of ATM, whereas the inhibitors of specific caspases were less effective. In caspase-3-deficient MCF-7 cells, ATM was cleaved, releasing the 210- but not the 150-kDa fragment. Recombinant caspase-3 was much more effective than caspase-7 in cleaving ATM that was immunoprecipitated from cell lysates. During cisplatin incubation, VAD protected ATM and enhanced p53 phosphorylation. In vitro assay of protein kinase activity further showed that ATM immunoprecipitated from cisplatin-treated cells had significantly lower kinase activity toward p53 than that from control cells. Importantly, the protein kinase activity was restored in ATM that was protected by VAD during cisplatin incubation. ATM deficiency sensitized the cells to cisplatin-induced apoptosis, suggesting a cytoprotective role of ATM in this experimental model. Thus proteolysis of ATM by caspases may inactivate this regulatory molecule to facilitate the progression of apoptosis.
【关键词】 ATR proteolysis DNA damage kidney nephrotoxicity
CISPLATIN AND RELATED COMPOUNDS are widely used for cancer therapy. Major side effects of the therapeutics include renal cell injury and tissue damage, resulting in nephrotoxic acute renal failure ( 2, 34 ). Chemoprotective strategies may be derived from the understanding of the critical molecules or molecular steps that regulate renal cell injury and death during cisplatin treatment ( 1, 3, 5, 7, 12, 19 - 22, 24 - 26, 28, 29, 31, 36, 42 ).
After entering the cell, cisplatin becomes highly reactive. Depending on its concentrations, cisplatin may conjugate with molecules that contain nucleophilic groups including glutathione, proteins, RNA, and most noticeably, genomic DNA ( 18, 32 ). Intra- and interstrand cross-linking of DNA by cisplatin not only blocks DNA replication and gene transcription, it also induces single- as well as double-strand DNA breaks. As a result, DNA damage by cisplatin has been recognized as a critical determinant of the cytotoxicity of this chemotherapy drug ( 18, 32 ). Consistently, in renal tubular cells, we and others showed that cisplatin-induced apoptosis involves p53, a critical mediator of DNA damage signaling ( 8, 16, 17, 28 ). However, the upstream events that regulate p53 activation under these conditions remain elusive.
ATM (ataxia telangiectasia-mutated) and ATR (ATM-related) are two apical protein kinases that are activated early in response to DNA damage ( 23, 30 ). Upon activation, these two proteins kinases can further phosphorylate and activate downstream protein kinases such as CHK1 and CHK2, leading to p53 phosphorylation and activation. Alternatively, ATM and ATR may directly phosphorylate and activate p53 and other effector molecules ( 23, 30 ). Despite the general understanding, little is known about the regulation of ATM and ATR during cisplatin-induced renal cell injury and nephrotoxicity.
During the course of examining ATM and ATR, we detected ATM cleavage during cisplatin treatment of renal tubular cells. Our subsequent experiments suggest the involvement of caspases in ATM cleavage in the experimental setting. Importantly, cleavage of ATM appears to inactivate its protein kinase activity toward p53. We further show that cisplatin induces higher apoptosis in ATM-deficient cells. Together, the results suggest a cytoportective role for ATM. Proteolysis of ATM by caspases may inactivate this regulatory protein and facilitate the progression of apoptosis during cisplatin treatment.
MATERIALS AND METHODS
Cells. The rat kidney proximal tubular cell line (RPTC) was originally from Dr. U. Hopfer (Case Western Reserve University, Cleveland, OH) and maintained for experiments as described ( 40 ). HeLa cells were from American Type Culture Collection and maintained as previously ( 41 ). MCF-7 wild-type cells that were deficient of caspase-3 were purchased from American Type Culture Collection. MCF-7 cell lines that were stably transfected with caspase-3 were described in previous work ( 14 ). ATM-mutant (ataxia telangiectasia) and control fibroblasts were originally obtained from Coriell Institute (Camden, NJ) and cultured as described ( 38 ).
Reagents. Mouse monoclonal anti-ATM (2C1), rabbit polyclonal anti-ATM (H248), and goat polyclonal anti-ATR (N19) antibodies were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). Rabbit polyclonal anti-p53 and anti-phospho-p53 (Ser15) antibodies were purchased from Cell Signaling Technology (Beverly, MA). Mouse monoclonal antibody to caspase-3 was from Transduction Laboratories (Lexington, KY). Monoclonal anti- -actin antibody was purchased from Sigma (St. Louis, MO). Cell-permeable peptide inhibitors of caspases including VAD, YVAD, IETD, LEHD, VDVAD, DEVD, and DQMD in the form of fluoromethyl ketone were purchased from Enzyme Systems Products (Dublin, CA). Recombinant p53 was purchased from Santa Cruz Biotechnology. Recombinant caspase-3 and -7 were purchased from BioVision (Mountain View, CA). Other reagents including cisplatin and pifithrin- were from Sigma.
Cell treatment. Cisplatin incubation of cells was described in our recent study ( 16 ). Briefly, cells were plated to reach 90% confluence by the next day for incubation with cisplatin. The concentrations of cisplatin that were used in this study were 20 µM for RPTC and MCF-7 cells, and 30 µM for HeLa cells. These concentrations were titrated in pilot experiments to induce significant amounts of apoptosis within 24 h, without contamination of necrosis (not shown). For inhibitory experiments, inhibitors were added along with cisplatin. To induce apoptosis by ATP depletion, RPTC cells were incubated in glucose-free buffer for 3 h with 10 mM azide, a cellular respiration inhibitor, followed by recovery in full culture medium for 2 h ( 39 ). Apoptosis was also induced in RPTC cells by using 1 µM staurosporine, a general protein kinase inhibitor, as described in our previous work ( 10 ).
Examination of apoptosis. Apoptosis was monitored by morphological methods as described previously ( 16, 37 ). Following staining with 10 µg/ml of Hoechst 33342, cellular and nuclear morphology was examined by phase contrast and fluorescence microscopy (excitation 340 nm/emission 450 nm). Apoptosis was indicated by cellular shrinkage, nuclear condensation and fragmentation, and the formation of apoptotic bodies. Four fields with 200 cells were checked in each dish to estimate the percentage of apoptosis. Representative fields were recorded by phase contrast and fluorescence microscopy to show cellular and nuclear morphology.
Immunoblot analysis. Whole cell lysates were collected using a buffer containing 2% SDS. Protein concentration was measured with the bicinchoninic acid (BCA) reagent (Pierce Chemical, Rockford, IL). Same amounts (usually 25 µg) of protein were loaded from each sample for electrophoresis under reducing condition. ATM was a large protein with an apparent molecular size of 350 kDa. To separate ATM from its cleaved fragments, 6% SDS-PAGE gels were used. For other proteins, 10% gels were used. After electrophoresis, the resolved proteins were electroblotted onto PVDF membranes. The membranes were then incubated with a blocking solution containing 1% (wt/vol) BSA and 2% (wt/vol) fat-free milk and exposed to the primary antibodies overnight at 4°C. After extensive washing and blocking in 5% (wt/vol) milk, the blot membranes were incubated with the horseradish peroxidase-conjugated secondary antibody. Antigens on the blots were revealed using the enhanced chemiluminescence (ECL) kit from Pierce.
Immunoprecipitation of ATM kinase. Cells were extracted with a buffer containing 150 mM NaCl, 1 mM EDTA, 1 mM EGTA, 1% (vol/vol) Triton X-100, 2.5 mM sodium pyrophosphate, 1 mM -glycerol phosphate, 1 mM Na 3 VO 4, 10 µg/ml leupeptin, 10 µg/ml aprotinin, 1 mM phenylmethylsulfonyl fluoride, 50 mM NaF, 0.2% (wt/vol) dodecyl - D -maltoside, and 20 mM Tris (pH 7.5). The soluble extracts were subjected to immunoprecipitation according to our previous work ( 10, 37, 41 ). A rabbit polyclonal anti-ATM antibody (H248) was chosen for immunoprecipitation in this experiment, due to its capacity of precipitating both intact and cleaved ATM (see Fig. 7 A ). Briefly, cell lysates of 400 µg were incubated with 1 µg normal mouse serum and 30 µl agarose protein A/G (Santa Cruz Biotechnology) for preclearing to remove nonspecific binding. The precleared samples were then incubated with 2 µg of rabbit polyclonal anti-ATM antibody and 30 µl of agarose protein A/G. Immunoprecipitates were washed and collected by centrifugation. A portion of the precipitates was subjected to gel electrophoresis and immunoblot analysis to confirm the successful immunoprecipitation of ATM and its cleaved fragments. Another portion was used for p53 kinase activity analysis.
Fig. 1. ATM cleavage during cisplatin-induced tubular cell apoptosis. RPTC cells were incubated with 20 µM cisplatin for 0-24 h. Whole cell lysates were collected in 2% SDS buffer for immunoblot analysis. Apoptosis was estimated by morphological methods. A : ATM blot. B : ATM-related (ATR) blot. C : percentage of apoptosis. The immunoblot results are representatives of at least 2 separate experiments. The data in C are expressed as means ± SD ( n = 4). The results show that ATM, and not ATR, is cleaved during cisplatin-induced apoptosis of renal tubular cells.
Fig. 2. ATM cleavage in other cells and apoptotic models. A : ATM cleavage in HeLa cells during cisplatin treatment. HeLa cells were incubated with 30 µM cisplatin for 0-24 h. Whole cell lysates were collected for immunoblot analysis of ATM. B : ATM cleavage during apoptosis induced by ATP depletion and staurosporine in RPTC cells. RPTC cells were incubated for 3 h with 10 mM azide in a glucose-free buffer followed by 2 h of recovery in cell culture medium (Azide) or incubated for 5 h with 1 µM staurosporine in culture medium (STS). Whole cell lysates were collected for immunoblot analysis of ATM. The immunoblots are representatives of at least 2 separate experiments.
Fig. 3. Inhibition of ATM cleavage during cisplatin incubation by caspase inhibitors. RTPC cells were incubated with 20 µM cisplatin for 24 h in the absence (NA) or presence of 10, 20, or 50 µM caspase inhibitors. At the end of incubation, whole cell lysates were collected for immunoblot analysis of ATM. The immunoblots are representatives of 3 separate experiments.
Fig. 4. Effects of caspase inhibitors on cisplatin-induced apoptosis. RTPC cells were incubated with 20 µM cisplatin for 24 h in the absence or presence of 50 µM caspase inhbitors. At the end of incubation, cells were stained with Hoechst 33342. A : apoptosis was estimated by morphological methods. Data are expressed as means ± SD ( n = 4). *Statistically significantly different from the control group. #Statistically significantly different from the cisplatin-treated group without caspase inhibitors. B : cellular and nuclear morphology were recorded by phase contrast and fluorescence microscopy.
Fig. 5. Involvement of caspase-3 and -7 in ATM cleavage during cisplatin incubation. A : effects of caspase-3 inhibitor DQMD on ATM cleavage during cisplatin incubation. RTPC cells were incubated with 20 µM cisplatin for 24 h in the absence (NA) or presence of 10 or 50 µM caspase inhibitors. B : immunoblot analysis of caspase-3 expression in wild-type MCF-7 cells and MCF-7 cells stably transfected with caspase-3. C : ATM cleavage during cisplatin incubation of caspase-3-deficient and caspase-3-expressing MCF-7 cells. The cells were incubated with 20 µM cisplatin for 0-24 h to collect whole cell lysates for immunoblot analysis of ATM. D : ATM cleavage by recombinant caspase-3 and -7 in vitro. ATM was immunoprcipitated from RPTC cell lysates and then incubated with 1 unit of recombinant caspase-3 or -7 for 1 h. The incubation mixtures were then analyzed for ATM by immunoblotting.
Fig. 6. Effects of VAD inhibition of ATM cleavage on p53 phosphorylation during cisplatin treatment. RPTC and HeLa cells were incubated with 20 µM cisplatin for 24 h in the absence (-) or presence (+) of 50 µM VAD. Whole cell lysates were collected for immunoblot analysis of phospho-p53. The blots were reprobed for -actin to monitor protein loading and transferring. The immunoblots are representative of 3 separate experiments. The results show that VAD increases p53 phosphorylation during cisplatin incubation of RPTC and HeLa cells.
Fig. 7. In vitro assay of ATM kinase activity in immunprecipitates from control, cisplatin-treated, and cispaltin-treated VAD-protected cells. ATM was immunoprecipitated from control, cisplatin-treated, and cispaltin-treated VAD-protected cells. A : immunoblot analysis to confirm immunoprecipitation of ATM and its cleaved fragments. B : ATM kinase activity shown as p53 phosphorylation in in vitro assays. The same amounts of immunoprecipitates were added to a protein kinase assay buffer containing recombinant p53. At the end of reaction, the reaction mixtures were subjected to electrophoresis and immunoblot analysis of phophorylated p53 using an anti-phospho-p53 (ser15) antibody. The blots are representatives of 3 separate experiments. C : semiquantification of phospho-p53 in protein kinase assays by densitometry. The phospho-p53 signal obtained from the immunoprecipitate of control cells was arbitrarily set as 100%, and the signals from the other 2 lanes on the same blots were normalized.
In vitro assay of ATM kinase activity toward p53. The ATM kinase assay was modified from Zhang et al. ( 43 ). ATM and its cleaved fragments were immunoprecipitated from control and cisplatin-treated cells as described above. The same amounts of immunoprecipitate were added to a reaction buffer containing 150 mM NaCl, 4 mM MnCl 2, 6 mM MgCl 2, 10% (vol/vol) glycerol, 1 mM dithiothreitol, 100 µM Na 3 VO 4, 50 mM HEPES (pH 7.5), supplemented with 20 µM ATP and 12.5 ng/µl recombinant p53. The reaction was incubated at 30°C for 20 min. Under this condition, the reaction was within the linear range as determined by pilot experiments (not shown). At the end of reaction, 2% SDS was added, and the samples were subjected to gel electrophoresis, followed by immunoblot analysis of phosphorylated p53. ATM kinase activity was indicated by p53 phosphorylation during the reaction.
Statistics. Data were expressed as means ± SD ( n 3). Statistical analysis was conducted using the GraphPad Prism software. Statistical differences between two groups were determined by Student's t -test. P < 0.05 was considered significantly different.
RESULTS
ATM cleavage during cisplatin-induced tubular cell apoptosis. Depending on its concentrations, cisplatin induces necrosis as well as apoptosis in vivo in the kidneys ( 27, 31, 36 ). To study the apoptotic mechanism of cisplatin nephrotoxicity, our recent work established an in vitro model using cultured RPTC ( 16 ). In this model, 20 µM cisplatin induces apoptosis in a time-dependent manner, with minimal contamination of necrotic cell death. To analyze changes of ATM and ATR, whole cell lysates were collected at various time points of cisplatin incubation and subjected to immunoblot analysis. As shown in Fig. 1 A, a noticeable cleavage of ATM was detected. ATM cleavage started after 8 h of cisplatin treatment, showing a distinguished fragment of 210 kDa. When the incubation was extended to 16 h, the 210-kDa band intensified, accompanied by the appearance of a lower band of 150 kDa. By the end of 24 h of cisplatin incubation, the degradation of full-length ATM was obvious and the cleaved fragments became more noticeable. In contrast, in the same cells, ATR was not cleaved. The time course of ATM cleavage during cisplatin incubation correlated well with the development of apoptosis, which started from 8 h and reached a maximal level by the end of 24 h ( Fig. 1 C ).
ATM cleavage in other cells and apoptotic models. To test whether the observed ATM cleavage was cell type specific, we analyzed HeLa cells. As shown in Fig. 2 A, following 30 µM cisplatin treatment for 16-24 h, ATM was cleaved in HeLa cells, and as in renal tubular cells, two fragments were detected ( lanes 4 and 5 ). ATM cleavage in HeLa cells also correlated well with the development of apoptosis, which was 20-30 and 40-50% at 16 and 24 h of cisplatin incubation, respectively. To test whether ATM cleavage was restricted to cisplatin-induced apoptosis, we analyzed ATM during apoptosis of renal tubular cells following ATP depletion or staurosporine treatment ( Fig. 2 B ). Clearly, ATM was also cleaved in these apoptotic models ( lanes 2 and 3 ), releasing similar fragments as that of cisplatin-treated cells ( lane 4 ). Thus ATM cleavage is not cell type or experimental model specific; rather, it appeares to be a common observation associated with apoptosis.
Inhibition of ATM cleavage by caspase inhibitors. The cleavage of ATM into specific fragments suggested the involvement of a specific protease(s). Because the cleavage correlated with the development of apoptosis, it was reasoned that caspases, a family of cysteine proteases involved in cisplatin-induced apoptosis ( 8, 19, 21, 28 ), might be responsible for ATM cleavage. To test the possibility, we first examined the effects of the broad-spectrum caspase inhibitor VAD. VAD, when added during cisplatin incubation, partially suppressed ATM cleavage at 10-20 µM (shown in Fig. 5 A ) and completely blocked ATM cleavage at 50 µM ( Fig. 3, lane 3 ). To gain further information on the involvement of specific caspases, we tested the effects of cell-permeable peptide inhibitors with documented specificity. The results are shown in Fig. 3. The caspase-1 inhibitor YVAD had little effects on ATM cleavage at 10-50 µM ( lanes 4 - 6 ). IETD and LEHD, specific inhibitors of caspase-8 and -9, were not effective at 10-20 µM and slightly inhibited ATM cleavage at 50 µM ( lanes 8 and 12 ). Similar results were shown for VDVAD, a specific inhibitor of caspase-2. DEVD, a peptide inhibitor of executioner caspases including caspase-3, -6, and -7, appeared to have some inhibitory effects at 50 µM, releasing less 210-kDa ATM fragment and no 150-kDa fragment ( lane 18 ). We also monitored the effects of these caspase inhibitors on apoptosis. As shown in Fig. 4 A, over 70% cells underwent apoptosis after 24 h of cisplatin treatment. VAD at 50 µM suppressed apoptosis to 30%, while YVAD showed a marginal and statistically insignificant inhibition. The inhibitory effects of IETD, LEHD, and VDVAD were comparable, reducing apoptosis to 50-60%. In the same experiment, DEVD suppressed apoptosis to 50% ( Fig. 4 A ). Shown in Fig. 4 B are images of cellular and nuclear morphology of representative fields. Of note, neither cisplatin-induced apoptosis nor ATM cleavage was suppressed by FA, a negative control compound of the caspase inhibitors (not shown). Apparently, the inhibitory efficacies of the caspase inhibitors on apoptosis and ATM cleavage showed a positive correlation. Together, the results suggest the involvement of caspases in the proteolysis of ATM during cisplatin treatment, but the cleavage does not seem to result from the activity of a specific caspase.
Involvement of caspase-3 in ATM cleavage during cisplatin treatment. In other experimental models, previous studies reported ATM cleavage during apoptosis ( 15, 33, 35 ). Interestingly, although these studies showed different cleavage patterns, releasing distinct ATM fragments, they postulated the involvement of caspase-3. However, our inhibitory experiments showed that DEVD, an inhibitor of caspase-3, -6, and -7, was not as effective as VAD, a broad-spectrum caspase inhibitor, in inhibiting ATM proteolysis during cisplatin treatment. To further resolve this issue, we compared the inhibitory effects of VAD and DQMD, a specific peptide inhibitor of caspase-3 ( 11 ). As shown in Fig. 5 A, following 24 h of cisplatin incubation, ATM was cleaved, releasing two fragments ( lane 2 ). VAD showed inhibitory effects on ATM cleavage at 10 µM ( lane 3 ). With 50 µM VAD, ATM cleavage was completely blocked, as evidenced by the preservation of full-length ATM and the absence of cleaved fragments ( lane 4 ). On the contrary, DQMD had no effects on ATM cleavage at 10 µM ( lane 5 ), and at 50 µM, only marginal inhibitory effects were shown ( lane 6 ). To further determine the role of caspase-3 in ATM proteolysis, we analyzed a MCF-7 cell line that was deficient of caspase-3. As a control, the same experiment also analyzed MCF-7 cells that were stably transfected with caspase-3. The absence and presence of caspase-3 in these two cell lines were confirmed by immunoblot analysis ( Fig. 5 B ). When the cells were incubated with cisplatin, significant apoptosis developed after 16-24 h (data not shown). At these time points, ATM cleavage was detected in caspase-3+ cells, releasing the 210- and 150-kDa fragments ( Fig. 5 C : lanes 9 and 10 ). In caspase-3-deficient cells, ATM was still cleaved, showing the loss of intact ATM and the appearance of the 210-kDa fragment; nevertheless, the release of the 150-kDa fragment was suppressed ( Fig. 5 C : lanes 9 and 10 ). In addition to caspase-3, caspase-7 has been shown to cleave ATM ( 15 ). We examined ATM cleavage by recombinant caspase-3 and -7 ( Fig. 5 D ). ATM was immunoprecipitated from RPTC cell lysates and then incubated with recombinant caspases, followed by immunoblot analysis of ATM. As shown in Fig. 5 D, both caspases cleaved ATM, but caspase-3 was much more effective in this function.
Effects of ATM preservation by VAD on p53 phosphorylation during cisplatin treatment. What was the functional consequence of ATM cleavage? To address this question, we examined p53 phosphorylation during cisplatin incubation in the absence or presence of VAD. It was reasoned that, if ATM cleavage regulated its kinase activity, the level of p53 phosphorylation would change under conditions where ATM was protected by VAD. In this experiment, we specifically analyzed p53 phosphorylation at serine 15, a site shown to be phosphorylated by ATM in response to DNA damage ( 4, 6 ). The results are shown in Fig. 6. In both renal tubular cells and HeLa cells, cisplatin treatment induced drastic increases in p53 phosphorylation at serine-15 (p-p53; lanes 2 and 5 compared with lanes 1 and 4 ). In the presence of VAD, which prevented ATM cleavage, p53 phosphorylation was increased drastically in HeLa cells ( lane 6 ) and, to a lesser extent, in renal tubular cells ( lane 3 ).
Cleavage of ATM inactivates its kinase activity toward p53 in in vitro assays. To further determine the functional consequences of ATM cleavage, we immunoprecipitated ATM and its fragments and analyzed their kinase activity toward p53. As shown in Fig. 7 A, intact ATM was precipitated from the control cell lysate ( lane 1 ), while both ATM and its fragments were precipitated from cisplatin-treated cell lysate ( lane 2 ). In cells that were treated with cisplatin but protected with VAD, mainly intact ATM was precipitated ( lane 3 ). When these three types of immunoprecipitates were added to a reaction containing recombinant p53, the immunoprecipitate of control cells showed the highest protein kinase activity, leading to rapid p53 phosphorylation ( Fig. 7 B : lane 1 ). The immunoprecipitate of cisplatin-treated cells, which contained cleaved ATM, had obviously lower p53 kinase activity ( lane 2 ). Importantly, VAD prevented ATM cleavage during cispaltin incubation ( Fig. 7 A : lane 3 ) and partially restored the ATM kinase activity toward p53 ( Fig. 7 B : lane 3 ). The conclusion was further substantiated by densitometry of the phospho-p53 signals from separate experiments ( Fig. 7 C ). Together, the results suggested that cleavage of ATM during cispaltin-induced apoptosis inactivated its kinase activity.
ATM deficiency sensitizes cells to cisplatin-induced apoptosis. Molecular signaling originated from ATM is very complex. Depending on the experimental conditions, it may lead to apoptosis, cell cycle arrest, or DNA repair and cell recovery ( 23, 30 ). Previous work showed that overexpression of ATM protects Rat-1 cells against c- myc -induced apoptosis ( 15 ). To determine the involvement of ATM in cisplatin-induced apoptosis, we compared wild-type and ATM-deficient fibroblasts. ATM expression in the wild-type but not ATM-deficient cells was confirmed by immunoblot analysis (data not shown). Basal levels of apoptosis in these two cell lines were below 5%. Incubation with 50 µM cisplatin for 24 h induced 47.5% apoptosis in wild-type cells. The same treatment induced 73.8% apoptosis in ATM-deficient cells, significantly higher than the wild-type ( Fig. 8 ). Similarly, 16 h of cisplatin treatment induced 5-10% apoptosis in wild-type cells but 20-30% in ATM-deficient cells (not shown). The worsened apoptosis in ATM-deficient cells suggests that ATM might have cytoprotective effects during cisplatin injury.
Fig. 8. Cisplatin-induced apoptosis in wild-type and ATM-deficient cells. Wild-type and ATM-deficient human fibroblast cells were incubated with 50 µM cisplatin for 24 h. Apoptosis was assessed by morphological criteria. Data are expressed as means ± SD ( n = 4). *Statistically significantly different from the control. #Statistically significantly different from the cisplatin-treated wild-type group.
DISCUSSION
ATM and ATR belong to the PI3K-related protein kinase family, which are critical regulators of cellular response to DNA damage ( 23, 30 ). Depending on the nature of DNA damage, ATM and ATR can be activated together or separately. Upon activation, these two kinases phosphorylate various substrate proteins to initiate DNA repair, cell cycle arrest, or cell death by apoptosis ( 23, 30 ). Despite intensive research in cancerous cells, the information on ATM/ATR regulation in normal tissues including the kidneys is very limited. Interestingly, recent work suggested ATM activation in response to high salt-induced DNA damage and a role of ATM in osmotic stress in renal medullary cells ( 9 ).
The current study showed the first evidence of proteolytic regulation of ATM during cisplatin-induced apoptosis of renal tubular cells. The regulation appears to be specific for ATM, since ATR is not proteolysed or cleaved in the same cells. Experiments using peptide inhibitors further suggest the involvement of caspases in ATM cleavage. Importantly, cleavage of ATM seems to inactivate the protein kinase activity toward p53. ATM and ATR have sequence homology but the homology is not very high. Rat ATM and ATR have 19.1% identity and 34.4% similarity in sequence (database analyzed by EMBOSS-Align of EMBL-EBI). The potential caspase cleavage sites in ATM including the DLCD site at 1372-1375 are not conserved in ATR.
While ATM cleavage is blocked by VAD, a broad-spectrum caspase inhibitor, peptide inhibitors with specificity toward individual caspases are less effective. Of note, all of the peptide inhibitors used in this study are cell permeable due to side-chain modification via methylation. Among the inhibitors tested in this study, YVAD, IETD, and LEHD are well-documented inhibitors for caspase-1, -8, and -9, respectively ( 13 ). VDVAD is a peptide inhibitor specific for caspase-2. All these caspases are considered upstream initiator caspases in various apoptotic models and have been implicated in the development of tubular cell apoptosis and nephrotoxicity during cisplatin treatment ( 19, 28, 36 ). In the experimental model of the current study, we also show that the various caspase inhibitors suppress apoptosis to some extents, but none of them is more effective than VAD ( Fig. 4 ). Interestingly, the efficacies of the caspase inhibitors on apoptosis and ATM cleavage correlate ( Figs. 3 and 4 ). Recent study further suggested the involvement of endoplasmic reticulum stress and caspase-12 in cispaltin-induced apoptosis in renal tubular cells ( 21 ). Thus it is likely that cisplatin treatment activates multiple pathways of caspase activation, culminating in the final outcome of apoptosis. The fact that inhibitors of specific caspases are not as effective as VAD in blocking ATM cleavage is in line with the complexity of apoptotic signaling under this pathological condition.
ATM cleavage during apoptosis has been documented in other experimental models ( 15, 33, 35 ). However, the patterns of ATM cleavage in these studies are inconsistent. Smith et al. ( 33 ) showed ATM fragments of 100, 150, and 240 kDa, while Hotti et al. ( 15 ) detected four ATM fragments of 50, 60, 150, and 230 kDa. On the other hand, Tong et al. ( 35 ) showed a much smaller fragment of 20 kDa. Interestingly, despite the differences in ATM cleavage patterns, these studies suggested the involvement of caspase-3. Particularly, partially purified ATM could be proteolytically cleaved by recombinant caspase-3 in vitro ( 33, 35 ). In our experiments, we detected two ATM fragments following its cleavage, with apparent sizes of 210 and 150 kDa. The cleavage pattern was not cell type or species specific, because the cleavage fragments were shown in both human (HeLa) and rat (RPTC) cells. Noticeably, these two ATM fragments were not generated at the same time. The 210-kDa fragment was detected at 8 h of cisplatin treatment and reached a maximal level at 16 h. Subsequently, this fragment was degraded, accompanied by the generation of the 150-kDa fragment ( Fig. 1 A ). The results suggest a sequential proteolysis of ATM. The first cleavage releases the 210-kDa fragment, and further cleavage of 210 kDa leads to the generation of the 150-kDa fragment. Interestingly and somewhat to our surprise, caspase-3 does not seem to play a critical role in the first step of ATM cleavage during cisplatin-induced apoptosis. Peptide inhibitors of caspase-3 were not as effective as the general caspase inhibitor VAD in blocking ATM cleavage. Furthermore, ATM was cleaved into the 210-kDa fragment in caspase-3-deficient MCF-7 cells following cisplatin incubation, although the generation of the 150-kDa fragment was suppressed ( Fig. 5 ). The discrepancy between this and earlier studies may derive from the differences in the experimental models. As discussed above, cisplatin may activate multiple pathways of apoptosis, resulting in the activation of various caspases. Recombinant caspase-3 was much more effective than caspase-7 in cleaving ATM immunoprecipated from the cells ( Fig. 5 D ). The results suggest that, in renal tubular cells where both caspases are present, caspase-3 is mainly responsible for ATM cleavage, while in caspase-3-deficient MCF, caspase-7 may cleave ATM. Considering the functional redundancy of caspases, particularly the downstream executioner caspases, it is not surprising that proteolysis of a substrate protein (e.g., ATM) persists under conditions of inhibition or deficiency of specific caspases.
In this study, ATM cleavage occurred at late stage of cisplatin treatment ( Fig. 1 ), whereas p53 phosphorylation and protein induction started earlier (not shown, Ref. 17 ), suggesting that ATM cleavage is not involved in early p53 regulation. This however does not suggest that ATM per se is not involved in early p53 regulation during cisplatin treatment. The role of ATM in cisplatin nephrotoxicity is unknown, which requires a systematic analysis to determine probably using transgenic and gene knockout models. For the functional consequence of ATM cleavage, we showed that the cleavage inactivated ATM's kinase activity toward p53 ( Fig. 7 ). Moreover, prevention of ATM cleavage by VAD increased p53 phosphorylation in HeLa and RPTC cells ( Fig. 6 ), suggesting that ATM participates in p53 regulation at late stage of cisplatin treatment. Our results further showed that ATM-deficient cells were more sensitive to cisplatin-induced apoptosis ( Fig. 8 ), suggesting that ATM is cytoprotective in this experimental model. We postulate that functionally ATM cleavage by caspases may diminish the cytoprotective effects of ATM to facilitate the progression of apoptosis.
GRANTS
This study was supported in part by grants from National Institutes of Health and Department of Veterans Affairs.
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作者单位：1 Department of Cellular Biology and Anatomy, 2 Center for Genomic Medicine, Medical College of Georgia, Augusta; 4 Medical Research Service, Department of Veterans Affairs Medical Center, Augusta, Georgia; and 3 Mary Babb Randolph Cancer Center, West Virginia University, Morgantown, West Virginia