ToxSci Advance Access originally published online on June 12, 2007
Toxicological Sciences 2007 99(1):346-353; doi:10.1093/toxsci/kfm152
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Cisplatin, Gentamicin, and p-Aminophenol Induce Markers of Endoplasmic Reticulum Stress in the Rat Kidneys
* Biomedical Sciences
Pathology and Microbiology, University of Prince Edward Island, Charlottetown, PE, C1A 4P3 Canada
1 To whom correspondence should be addressed at Biomedical Sciences, University of Prince Edward Island, 550 University Av., Charlottetown, PE C1A 4P3 Canada. Fax: (902) 566-0832. E-mail: mpeyrou{at}upei.ca.
Received December 6, 2006; accepted June 6, 2007
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ABSTRACT |
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In vitro evidence of the involvement of the endoplasmic reticulum (ER) during drug-induced renal toxicity is accumulating. ER stress and ER-mediated cell death markers have been reported after exposure of renal cells to model toxicants and nephrotoxic drugs in various in vitro models, but in vivo experiments with clinically relevant nephrotoxic compounds are lacking. In order to determine the relevance of the in vitro findings, markers of ER stress (XBP1 messenger RNA processing and protein expression; GRP78 and GRP94 upregulation) and ER-mediated cell death (caspase-12 and calpain activation) were examined in kidney tissue of rats exposed to nephrotoxic doses of cisplatin (CIS), gentamicin (GEN), and p-aminophenol (PAP), a nephrotoxic metabolite of acetaminophen. XBP1 signaling was observed with all three drugs and was associated with increased expression of GRP94 and GRP78 in GEN- and PAP-treated animals, but surprisingly not after CIS exposure. m-Calpain expression was increased after 7 days of CIS treatment, whereas it was decreased in PAP-treated rats. Caspase-12 cleavage products were increased after CIS, GEN, and PAP administration. The results of this study demonstrate that three clinically relevant nephrotoxic drugs are all associated with changes in markers of ER stress and ER-mediated cell death in vivo. Further investigation is warranted to determine the role of the ER, the calpain system, and caspase-12 in drug-induced renal cell death.
Key Words: endoplasmic reticulum stress; nephrotoxicity; nephrotoxic drugs; caspase; calpain.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONFUNDINGREFERENCES |
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Drug-induced renal toxicity is a frequent adverse drug reaction with potentially devastating consequences on patients' health. Approximately 10–20% of acute renal failures are thought to be drug related (Brivet et al., 1996
) and one quarter of the 100 most used drugs in Intensive Care Units have nephrotoxic potential (Taber and Mueller, 2006). Mechanisms of renal damage are numerous, resulting in ischemic or chemically induced injury with endothelial, glomerular, tubular, papillary, or interstitial lesions (Schnellmann, 2001). At the cellular level, renal damage has been linked to covalent binding of reactive molecules to cellular macromolecules and oxidative stress (Schnellmann, 2001).
Evidence is accumulating that the endoplasmic reticulum (ER) is one of the subcellular target of toxic compounds and may play an important role in xenobiotic-induced nephrotoxicity (Cribb et al., 2005). Initial support was provided by in vitro experiments performed primarily with the porcine renal proximal tubular cell line LLC-PK1. Increased expression of ER stress proteins, a hallmark of ER stress, provided cytoprotection against model toxicants (tert-butylhydroperoxide, iodoacetamide, menadione, and others) causing oxidative stress and protein alkylation in LLC-PK1 cells (Bedard et al., 2004; Halleck et al., 1997; Hung et al., 2003; Liu et al., 1997, 1998; Peyrou and Cribb, 2007; Ryan et al., 2005; van De Water et al., 1999). The protective effect of prior ER stress was also substantiated in vivo in kidneys of rats exposed to the nephrotoxicant S-(1,1,2,2,-tetrafluoroethyl)-L-cysteine (Asmellash et al., 2005). Model cytotoxins and some clinically relevant nephrotoxins have also been associated with disruption of the ER and subsequent induction of ER stress proteins in in vitro systems. Cisplatin (CIS), cyclosporine A, acetaminophen, iodoacetamine (IDAM), menadione (MEN), and tert-butylhydroperoxide (TBHP) have been associated with induction of ER stress proteins and activation of ER-mediated cell death markers such as caspase-12, calpain, and GADD153 (Huang et al., 2001; Justo et al., 2003; Lame et al., 2003; Liu and Baliga, 2005; Lorz et al., 2004; Mandic et al., 2003; Muruganandan and Cribb, 2006; Paslaru et al., 1994; Ryan et al., 2005; Szegezdi et al., 2006). Inhibition of calpain, a calcium-dependent protease implicated in ER-mediated renal cell death, decreased cell death after IDAM, TBHP, and MEN exposure in LLC-PK1 cells (Muruganandan and Cribb, 2006) or after model toxicants and nephrotoxic compounds exposure in renal proximal tubule suspensions (Harriman et al., 2000; Schnellmann and Williams, 1998). Calpains have been shown to trigger plasma membrane degradation and loss of integrity of renal cells (Liu and Schnellmann, 2003). Finally, ER proteins can be direct targets of nephrotoxic compounds, such as aminoglycosides which can alter the activity of ER chaperone proteins (Horibe et al., 2002, 2004).
Although these in vitro data strongly suggest the involvement of the ER in cell death caused by nephrotoxicants, there is little in vivo evidence to support the involvement of the ER in nephrotoxicity. We hypothesized that, if the ER is involved in nephrotoxicity in vivo, changes in markers of ER stress and ER-mediated cell death would be observed in kidneys of rats exposed to nephrotoxicants. We elected to use three clinically relevant nephrotoxins to test this hypothesis: CIS, gentamicin (GEN), and p-aminophenol (PAP), a metabolite of acetaminophen. This study provides in vivo evidence of the involvement of the ER in drug-induced renal toxicity.
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MATERIALS AND METHODS |
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Chemicals and materials.
CIS, GEN, and PAP were purchased from Sigma-Aldrich Canada, Ltd (Oakville, ON, Canada). Sterile drug solutions were prepared in PBS and pH was adjusted to 7.4. Sterile materials were purchased from VWR International (Mississauga, ON, Canada). Routine chemicals were obtained from Sigma or Fischer Scientific (Nepean, ON, Canada).
Animal, treatment and sample collection.
The protocol of this study was approved by the Atlantic Veterinary College Animal Care Committee and followed the regulations of the Canadian Council on Animal Care.
Male Sprague–Dawley rats of 175–250 g body weight were purchased from Charles River Laboratories, Inc. (Wilmington, MA) and allowed 1 week of acclimatization upon arrival. All animals were housed in group of four or five animals, at a temperature of 22 ± 2°C with a relative humidity of 45 ± 10%, and submitted to a light cycle from 6 A.M. to 6 P.M. Animals had free access to food (Purina Rodent Chow diet #5001) and water at all time during the experiments. Doses of clinically relevant nephrotoxic compounds were selected from the current literature to result in significant toxicity based on histological examination and biochemical markers (Amin et al., 2004; Newton et al., 1982). CIS and GEN were injected ip once a day for 2 (CIS D2, GEN D2) and 7 days (CIS D7, GEN D7) at daily doses of 5 and 160 mg/kg, respectively. Twenty-four hours after the last injection, rats were anesthetized with pentobarbital for sample collection. PAP was given by a single ip injection for a dose of 225 mg/kg, and samples were collected 6 h (PAP H6) and 24 h (PAP H24) after administration.
Intracardiac blood samples and urine samples were obtained via laparo-thoracotomy. One kidney was removed and placed in formalin after longitudinal cut for histopathological examination. The other kidney was removed and cut to obtain two kidney poles and two transverse kidney slices. One kidney slice was snap-frozen in liquid nitrogen and kept at – 80°C until messenger RNA (mRNA) extraction (see below). The other slice was placed on ice in a calpain extraction buffer for calpain activity assay (see below). One kidney pole was kept on ice in Tris–KCl buffer (pH = 7.4) and homogenized with a tissue tearer (Polytron PT3000, Kinematica, Inc., Newark, NJ) within 15 min. Differential centrifugation was used to obtain the S9 fraction (9000 x g, 20 min, 4°C), cytosolic, and microsomal fractions (100,000 x g, 1 h, 4°C). The other kidney pole was kept on ice in caspase lysis buffer for caspase-12 activity assay (see below). Protein concentrations were determined with a modified Lowry protocol (DC Protein Assay, Bio-Rad Laboratories, Mississauga, ON, Canada).
Biochemistry and histopathological examination.
Serum creatinine (CREAT) and blood urea nitrogen (BUN) concentrations were assayed by spectrophotometry in a Roche Hitachi 917 analyzer (Roche Diagnostics, Indianapolis, IN; Cat. No. 11875418 and 11729691). Formalin-fixed tissues were embedded in paraffin, cut and stained with hematoxylin–eosin prior to light microscopic evaluation of the extent of tissue damage.
Immunoblotting.
Cell fraction proteins (30–100 µg per lane) were resolved by sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE) with 10% polyacrylamide gels and transferred onto a nitrocellulose membrane for Western blotting. The following antibodies were used: anti-m-calpain (RP1-calpain-2; 1/1000; Triple Point Biologics, Forest Grove, OR), anti-XBP1 (sc-7160; 1/500; Santa Cruz Biotechnology, CA), anti-caspase-12 (C7611; 1/500; Sigma-Aldrich Canada), anti-GRP78 (#610979; 1/1000; BD Biosciences, Mississauga, ON, Canada), anti-GRP94 (SPA-850; 1/5000; Sigma-Aldrich Canada). All secondary antibodies were purchased from Sigma-Aldrich Canada. ß-Actin was used as an internal loading standard.
Immunohistochemistry.
Kidney sections were deparaffinized and submitted to heat-induced antigen retrieval in citrate buffer (tribasic sodium citrate 10mM, 0.05% Tween 20, pH = 6) for 20 min at boiling temperature. A rabbit anti-XBP1 antibody (1/100; Santa Cruz Biotechnology) was used as primary antibody with an overnight incubation, followed by incubation with a fluorescent secondary antibody Alexa Fluor 594 goat anti-rabbit (1/200; 1 h; Invitrogen, Burlington, ON, Canada).
Activity assay.
Calpain and caspase-12 fluorometric activity assay kits were purchased from Biovision (Cat. No. K240 and K139, Mountain View, CA). Calpain and caspase-12 assays are based on detection of cleavage of a specific substrate, ac-LLY-AFC (7-amino-4-trifluoromethyl coumarin) and amino acids sequence-AFC, respectively. When the substrate is cleaved, free AFC emits a yellow-green fluorescence which can be quantified by a fluorometer. For the calpain assay, 1-mm-wide kidney slices were incubated 30 min at 4°C with mild shaking in the calpain extraction buffer provided in the assay kit and centrifuged (9000 x g, 10 min, 4°C). The supernatants were processed according to the manufacturer protocol, with 100 µg of extracted protein employed per reaction well. For the caspase-12 assay, S9 fractions of total kidney samples in lysis buffer were processed according to the manufacturer's protocol, with 200 µg of protein employed per reaction well.
RNA isolation.
Total RNA was extracted from approximately 50–100 mg of tissue with TRIzol reagent according to the manufacturer's directions (Invitrogen). RNA was quantified using ultraviolet (UV) spectroscopy on a Shimadzu UV-1601PC spectrophotometer (Mandel Scientific, Guelph, ON, Canada) on samples diluted in Na2PO4 buffer (10mM).
Reverse transcription and real-time polymerase chain reaction.
Total RNA (0.5 µg) was used along with Thermoscript RT-PCR (reverse transcription–PCR) Systems kit following the manufacturer's directions (Invitrogen) to produce complementary DNA (cDNA) for PCR reactions, using a GeneAmp PCR System 9700 thermocycler (Applied Biosystems, Toronto, ON, Canada). Platinum Sybergreen qPCR Supermix-UDG with ROX (10 µl) (Invitrogen) was used to amplify 4 µl of sample cDNA in a 20-µl reaction containing 0.4 µl of 10µM sense (5'-AAACAGAGTAGCA GCGCAGACTGC-3') and antisense (3'-GATCTCTAAAACTAGAGGCTTGGTG-5') rat XBP1 primers (BioCorp, Montreal, PQ, Canada) and 5.2 µl of sterile DD-H2O. Real-time amplification of XBP1 was performed with a Rotor Gene RG-3000 (Corbett Research, Montreal Biotech Inc., Kirkland, PQ, Canada) under the following conditions: 50°C for 2 min, 95°C for 2 min; 45 cycles of 95°C for 15 s, 55°C for 30 s, 72°C for 30 s, 83 for 15 s; then 72°C for 5 min. 18S ribosomal RNA (Ambion, Austin, TX) was used as an internal control and followed the same amplification cycling as XBP1. Relative quantification was determined by cycles to threshold, adjusted for 18S ribosomal RNA expression.
GRP78 levels were assessed using the same reverse transcription process. Real-time PCR primers were purchased from SuperArray (RT2 PCR primer set for rat GRP78; Cat. No. PRR45224A; Cerdarlane, Hornby, ON, Canada). RT2 Real-Time SYBR Green PCR Master Mix (10 µl) (SuperArray, Cerdarlane) was used in a 20-µl reaction, along with 4 µl of sample cDNA, 0.8 µl of 10µM GRP78 rat primer pair and 5.2 µl of sterile DD-H2O. Amplification cycling for GRP78 was 95°C for 10 min; 50 cycles of 95 for 15 s, 55°C for 30 s, 72°C for 30 s, then 72°C for 2 min.
Statistical analysis.
A minimum of three rats per treatment group was used for each experimental procedure. The results are presented as mean ± standard error of the mean. All statistical analyses were performed using GraphPad Prism version 3.03 (GraphPad Software, San Diego, CA). Data were analyzed by one-way ANOVA and Dunnett's post hoc tests. A p value lower than 0.05 was considered to reflect a statistically significant difference.
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RESULTS |
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In order to confirm the kidney damage resulting from the exposure to the three nephrotoxic drugs (CIS, GEN, and PAP), we assessed serum CREAT and BUN concentrations 24 h after the last injection. Increased CREAT concentrations were observed after CIS and PAP administration (Fig. 1). The rise of BUN concentration was robust after 7 days of CIS administration, but failed to achieve statistical significance for the other treatments. GEN administration resulted in a mild and nonstatistically significant increase of BUN or CREAT (Fig. 1), although it was associated with tubular necrosis in individual rats after 7 days of administration (Fig. 2B). Histological examination did not reveal any alterations after administration CIS and GEN for 2 days (data not shown). CIS-treated rats had extensive acute tubular necrosis within a narrow zone along the corticomedullary junction after 7 days of treatment. Most tubules in this region were dilated with tubular lumina containing sloughed necrotic or degenerating epithelial cells (Fig. 2C). Some scattered tubules contained proteinaceous casts. The PAP-treated rats had extensive coagulative necrosis of tubular epithelium in the inner cortical region at 6- and 24-h postadministration. Additionally, the lumina of many tubules in other areas, were filled with proteinaceous casts (Fig. 2D). CIS administration was associated with significant adverse clinical signs starting at day 4 (altered behavior, anorexia, dehydration). PAP and GEN were not associated with any noticeable clinical signs.
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XBP1 mRNA splicing and protein expression were monitored by real-time PCR and immunoblotting, respectively. XBP1 spliced mRNA was increased in kidney tissue of rats treated for 7 days with GEN and CIS, but not in rat kidneys 24 h after PAP treatment (Fig. 3A). XBP1 protein expression was not significantly altered after 2 days of CIS or GEN treatment, but was increased two to threefold in kidney S9 fraction of rats treated for 7 days (Fig. 3B). Immunohistochemistry revealed increased XBP1 antigenicity in the cortex of rat treated for 7 days with GEN (Fig. 4), but no changes were observed after 2 days of treatment (data not shown). CIS treatment was not associated with a specific pattern of increased antigenicity (Fig. 4). XBP1 protein expression was significantly increased 24 h after PAP administration, whereas it was only slightly increased 6-h postadministration (Fig. 3B). Increased XBP1 antigenicity was observed at 6 and 24 h but with a marked change in its localization with time. Six hours after PAP administration, XBP1 antigenicity matched perfectly the area of acute tubular necrosis, whereas it was found in tubular cells of dilated tubules by 24 h after PAP administration (Fig. 4) and was no longer seen in areas of significant cellular necrosis.
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GRP94 protein expression was significantly induced following 7 days of GEN and PAP treatment, but not after CIS administration (Fig. 5). GRP78 was significantly induced in GEN-treated rats. GRP78 was also induced in individual animals following PAP treatment, but there was greater variability between animals and the changes failed to achieve statistical significance despite the same trend as for GRP94 (Fig. 5). GRP78 mRNA expression was significantly increased in kidney of rats treated for 7 days with GEN, but was unchanged after CIS and PAP administration (Fig. 5).
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To explore the possible involvement of m-calpain in ER stress during nephrotoxic drug administration, m-calpain activation was monitored indirectly by immunoblotting of kidney S9 fractions and directly with a fluorometric assay performed on kidney tissue extracts. On immunoblots, the amount of 80-kDa calpain (uncleaved large subunit) was greatly decreased 6 and 24 h after PAP administration. CIS treatment resulted in a decreased expression of calpain after 2 days and in a mild but significant increase of calpain expression after 7 days of treatment (Fig. 6A). GEN was associated with decreased calpain expression after 2 days of treatment but no change was observed after 7 days (Fig. 6A). With the activity assay, both CIS and PAP administration resulted in significant reduction of calpain activity, but GEN decrease failed to achieve statistical significance (Fig. 6B).
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All three drugs were associated with increased caspase-12 cleavage products on immunoblots (Figs. 7A and 7B). CIS exposure also significantly increased the amount of uncleaved caspase-12 (Fig. 7B). No drugs achieved statistically significant alteration of caspase-12 activity, although CIS and PAP samples mean activities were slightly decreased (data not shown).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONFUNDINGREFERENCES |
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The role of the ER in nephrotoxicity has been under increasing scrutiny, as discussed in the introduction. ER stress and ER-mediated cell death markers were observed in various in vitro models of toxicity and ER stress preconditioning was shown to be protective against renal cell death in vitro (Cribb et al., 2005
). Despite the in vitro evidence implicating the involvement of the ER in renal cell death, very little in vivo data exist. The purpose of this study was to determine if markers of ER stress or ER-mediated cell death are altered in the kidney following in vivo exposure of Sprague–Dawley rats to the clinically relevant nephrotoxicants CIS, GEN, and PAP.
To uncover the presence of ER stress in rat kidneys after administration of these nephrotoxic drugs, we assessed the expression of the ER stress associated transcription factor XBP1 (Back et al., 2005; Calfon et al., 2002) by real-time PCR and by immunohistochemistry. The XBP1 transcription factor binds to two consensus sequences that regulate Unfolded Protein Response (UPR) gene expression: the ER stress response element and the unfolded protein response element. Upon ER stress, a 26-bp intron of the XBP1 pre-mRNA is spliced out by increased Inositol Requiring Enzyme 1 (IRE1) endoribonuclease activity, resulting in synthesis of the potent XBP1 transcription factor (Yoshida et al., 2001). CIS and GEN administration for 7 days resulted in 1.9- and 2.6-fold increases of spliced XBP1 mRNA, respectively. PAP administration was not associated with increased XBP1 mRNA splicing at 24 h, but XBP1 protein was found in greater amount than in control in kidney S9 fractions, as for GEN and CIS samples (Fig. 3). Failure to observe increased XBP1 spliced mRNA 24 h after PAP administration may have been related to the time point examined (the spliced mRNA may have already returned to baseline, with protein expression still increased) or to the use of total renal mRNA in which changes in a small injured area may be masked by normal tissue. Nevertheless, the observed changes in XBP1 mRNA and/or protein are consistent with activation of the ER stress response pathways in response to the toxicants.
Immunohistochemistry data further support these observations in GEN- and PAP-treated rats. There was a correlation between cellular damage and XBP1 expression. GEN administration was associated with cortical damage (Fig. 2; Mingeot-Leclercq and Tulkens, 1999) and increased XBP1 antigenicity observed in the cortex (Fig. 4). With PAP administration, XBP1 expression was first increased in the area of acute tubular necrosis and later in tubular cells of dilated tubules (Fig. 4). A clear pattern of expression was not observed with CIS.
To confirm that the activation of the IRE1–XBP1 signaling pathway was associated with upregulation of UPR-related genes, we assessed the expression of GRP94 and GRP78 proteins and upregulation of GRP78 mRNA. GRP94 and GRP78 protein and GRP78 mRNA expression were increased in kidneys of rats treated with GEN for 7 days. PAP administration led to increased GRP94 and GRP78 protein expression, although GRP78 mRNA was not increased. This is similar to the response observed for XBP1; GRP78 mRNA expression may have returned to baseline at 24-h postadministration despite an earlier induction. CIS did not alter the expression of GRP78 or GRP94 protein, nor GRP78 mRNA. The absence of GRP78 mRNA or protein upregulation with CIS treatment was unexpected as exposure of human melanoma cells to CIS has been associated with increased GRP78 mRNA and protein expression and increased XBP1 mRNA splicing (Mandic et al., 2003). Further studies would be required to determine if there was a true uncoupling between XBP1 and GRP78/GRP94 upregulation in rat kidney cells, or if there was only a modest increase that was not detected with the quantitative approaches used.
m-Calpain and µ-calpain are calcium-dependent proteases that have been implicated in cell death following renal ischemia/reperfusion injury and exposure to toxins or reactive chemicals (Harriman et al., 2002; Liu et al., 2001). Recently, calpain activation has been shown to be an important event in vitro for LLC-PK1 cells undergoing toxin-induced ER stress and ER-related cell death (Muruganandan and Cribb, 2006). To investigate a potential role of calpains in vivo in CIS, GEN, and PAP-induced nephrotoxicity, immunoblotting of kidney S9 fraction for m-calpain was performed. CIS and GEN administration for 2 days, as well as PAP treatment, resulted in a significant decrease of the parent immunoreactive band (around 80 kDa). After 7 days of treatment with CIS and GEN, m-calpain expression was, respectively, slightly increased or unchanged. No cleavage products were observed with any drugs. We have previously observed that tunicamycin and thapsigargin, known inducers of ER stress, can lead to an activation of calpain associated with a decrease in calpain expression without the appearance of immunoreactive bands (Muruganandan and Cribb, 2006). Moreover, we have observed that the appearance of calpain cleavage products and increases or decreases in parent calpain expression are very time and concentration dependent after exposure of renal cells to cytotoxins in vitro (Muruganandan and Cribb, 2006). Others have observed this variation when calpain is activated and the absence of autolyzed m-calpain does not indicate the absence of activity as full length calpain may display catalytic activity without cleavage (Goll et al., 2003). Therefore, while loss of parent calpain does not prove calpain activation, it can be consistent with calpain activation as both losses or increases in the parent calpain can be associated with activation of the calpain system depending on the specific time point examined and the conditions.
The immunoreactive band associated with PAP administration (Fig. 6) appears to have a slightly lower molecular weight, which may reflect the appearance of the active (autolyzed) 78-kDa calpain. Validation of this hypothesis would require protein sequencing as SDS-PAGE may not be sensitive enough to differentiate with certainty between the 80- and the 78-kDa form.
Another method for assessing calpain activation is to measure calpain activity. We measured calpain activity at a single time point and observed an apparent loss of calpain activity following CIS and PAP. This could be interpreted to reflect either a downregulation of calpain or it may be indicative of the consumption of calpain earlier in the time course of toxicity. The loss of calpain activity would be consistent with the loss of calpain protein observed and neither proves nor disproves the activation of calpain.
Other methods to illustrate the activation of the calpain system would be the use of calpain inhibitors and the demonstration of the appearance of calpain-dependent cleavage of other proteins. We have demonstrated in vitro that calpain inhibition blocks downstream calpain-associated events even when there is a loss of the parent calpain and no appearance of calpain cleavage products (Muruganandan and Cribb, 2006). In a related set of experiments (Peyrou and Cribb, 2007), we were able to demonstrate that calpain inhibition protected against PAP nephrotoxicity in vivo, confirming that the loss of parent calpain was in fact associated with calpain-mediated events. These data support the contention that the decrease in parent calpain observed following PAP, CIS, and GEN administration does reflect calpain activation.
Caspase-12 cleavage and activation is one of the hallmarks of ER-mediated cell death that has been observed in vitro following exposure of LLC-PK1 cells to cytotoxins (Muruganandan and Cribb, 2006; Ryan et al., 2005) and it has been shown that activated calpain can mediate this cleavage. Caspase-12 cleavage products were increased after PAP and CIS administration and CIS was also associated with an increased amount of procaspase-12. GEN also appeared to increase caspase-12 cleavage, but the increase was not statistically significant (p = 0.07). The lack of statistical significance with GEN administration might be explained by the combination of the lower kidney toxicity observed and the relatively small number of animals (four rats per group) used in this experiment. The appearance of caspase-12 cleavage products is consistent with activation of ER-related cell death pathways and the calpain system. Indeed, in related experiments, we observed that inhibition of calpain blocked the cleavage of caspase-12 following PAP administration (Peyrou and Cribb, 2007). Our observations with CIS are in concordance with in vitro studies suggesting a pivotal role of caspase-12 for CIS renal toxicity (Liu and Baliga, 2005); no in vitro studies with caspase-12 have been performed for PAP or GEN.
In an attempt to directly assess caspase-12 activity, S9 fractions of kidney tissues were assayed with a commercial capsase-12 fluorometric assay kit. No significant changes were observed, but we also had no positive control for activation of caspase-12 pathways in the rat.
The results of this study indicate that three clinically relevant nephrotoxicants (CIS, GEN, and PAP) are associated with changes in markers of ER stress and ER-mediated cell death signaling pathways. ER stress markers and ER-mediated cell death signaling pathways were not always altered to the same extent, but this is consistent with in vitro results and may in part reflect the time points and dose chosen to assess changes in these pathways (Muruganandan and Cribb, 2006; Ryan et al., 2005). This is the first description of the involvement of XBP1 splicing, GRP protein upregulation, calpain and caspase-12 activation during in vivo toxicity of clinically relevant drugs. These results warrant further investigation to determine the roles and relationship of the ER, the calpain system, and caspase-12 in renal cell death following exposure to nephrotoxic drugs. In a companion article, published in this journal, the consequences of ER stress preconditioning and calpain inhibition on the in vivo nephrotoxicity of PAP are presented (Peyrou and Cribb, 2007).
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FUNDING |
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Canadian Institutes of Health Research; CIHR-RPP fellowship to M.P.; and Canada Research Chair to A.C.
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ACKNOWLEDGMENTS |
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Work was performed with equipment provided by the Canada Foundation for Innovation.
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