1 Department of Medicine, Division of Nephrology, Hypertension and Transplantation, 4 Department of Neuroscience, and 3 Department of Pediatrics, University of Florida, Gainesville, Florida 32610; and 2 Department of Biology, Massachusetts Institute of Technology, Cambridge, Massachusetts 02139
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Heme
oxygenase-1 (HO-1) is a 32-kDa microsomal enzyme that catalyzes the
conversion of heme to biliverdin, releasing iron and carbon monoxide.
Induction of HO-1 occurs as a protective response in cells/tissues
exposed to a wide variety of oxidant stimuli. The chemotherapeutic
effects of cis-diamminedichloroplatinum(II) (cisplatin), a
commonly used anticancer drug, are limited by significant nephrotoxicity, which is characterized by varying degrees of renal tubular apoptosis and necrosis. The purpose of this study was to
evaluate the functional significance of HO-1 expression in cisplatin-induced renal injury. Our studies demonstrate that transgenic mice deficient in HO-1 (/
), develop more severe renal
failure and have significantly greater renal injury compared with
wild-type (+/+) mice treated with cisplatin. In vitro studies in human
renal proximal tubule cells demonstrate that hemin, an inducer of HO-1, significantly attenuated cisplatin-induced apoptosis and necrosis, whereas inhibition of HO-1 enzyme activity reversed the cytoprotective effect. Overexpression of HO-1 resulted in a significant reduction in
cisplatin-induced cytotoxicity. These studies provide a basis for
future studies using targeted gene expression of HO-1 as a therapeutic
and preventive modality in high-risk settings of acute renal failure.
acute renal failure; oxidative stress; cytoprotection; carbon monoxide; cancer chemotherapy
![]() |
INTRODUCTION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
REACTIVE OXYGEN SPECIES PLAY an important role in the pathogenesis of a variety of renal diseases, including acute renal failure (3, 28). The kidney is one of the prominent sites for intense oxidative processes in the body (28) and is, therefore, extremely vulnerable to free radical-mediated injury. Recent studies have reported that in states of heightened oxidant stress, heme oxygenase-1 (HO-1) is induced as a beneficial response in cells exposed to a diverse array of toxic insults (22, 30, 37). HO-1 is a ~32-kDa microsomal enzyme that catalyzes the initial and rate-limiting reaction in heme catabolism. It cleaves heme molecules to yield equimolar quantities of biliverdin, carbon monoxide, and iron. Biliverdin is subsequently converted to bilirubin by biliverdin reductase. Three isoforms of heme oxygenase, HO-1, HO-2, and HO-3, have been described (22, 24). HO-1 is inducible, whereas HO-2 is constitutively expressed in the brain, testes, endothelium, and the medullary thick ascending limb (mTAL) segment of the rat kidney (21, 22). A third isoform, HO-3, with properties similar to HO-2, has recently been described (24). HO-1 is induced by heme products and a variety of nonheme stimuli, including heavy metals, hydrogen peroxide, nitric oxide, endotoxin, cytokines, ultraviolet A irradiation, shear stress, hyperoxia, and oxidized low-density lipoproteins (LDL). Although the cellular processes underlying HO-1 induction are complex and tightly regulated, one denominator common to most of these stimuli is a significant shift in cellular redox (30).
With regard to acute renal injury, previous studies have shown that expression of HO-1 in renal tubules is cytoprotective in both heme- and nonheme-mediated models of renal failure. For example, in heme-mediated renal injury in a rat model of acute renal failure associated with rhabdomyolysis, administration of a specific inhibitor of HO-1, tin protoporphyrin, worsens renal damage. Prior induction of HO-1 with hemoglobin leads to a considerable decrease in mortality (29). Vogt et al. (39) demonstrated a novel phenomenon of acquired resistance to renal tubular injury in acute glomerular inflammation that was dependent on the induction of HO-1 in renal tubules. Induction of HO-1 occurs in immune-mediated renal injury, as demonstrated by the expression of this protein in infiltrating macrophages in acute renal transplant rejection (2). Expression of HO-1 has been linked to prolonged xenograft survival by investigators, showing shortened survival in hearts transplanted from HO-1-knockout mice compared with wild-type or heterozygote mice (36). Previous studies have shown that in cisplatin-induced toxic nephropathy, a model of acute oxidative stress not directly dependent on the delivery of heme proteins to the kidney, HO-1 is induced in renal tubules as early as 6 h after cis-diamminedichloroplatinum(II) [cisplatin (CP)] administration (1). Inhibition of HO-1 by tin protoporphyrin significantly worsens both structural and functional parameters of renal injury, providing indirect evidence for a role of HO-1 in CP nephrotoxicity.
The chemotherapeutic agent CP is widely used for the treatment of several human malignancies, including ovarian, testicular, bladder, head and neck, esophageal, and small-cell lung cancers (17). The beneficial antineoplastic effects of this drug are often mitigated by significant side effects, including nephrotoxicity, bone marrow suppression, peripheral neuropathy, ototoxicity, nausea, vomiting, and anaphylaxis. CP accumulates in the kidney to a greater degree than in other organs and preferentially injures the S3 segment of the proximal tubule and the mTAL, both situated in the outer medullary region of the kidney (33, 34). After a single injection of CP (50-100 mg/m2) for chemotherapy, 28-36% of patients develop dose-dependent nephrotoxicity (17, 33). Urinary levels of CP approach 50-250 mg/ml (~100-500 µM) within 2-6 h of CP infusion (20) and progressively decrease over a few days. Early reductions in glomerular filtration rate are associated with decreases in renal plasma flow and increased renal vascular resistance, making CP nephrotoxicity an ideal model to study the early pathophysiological and biochemical determinants of acute renal failure (34). The pathogenesis of CP-induced acute renal failure is characterized by both necrosis and apoptosis of renal tubular cells (19, 20). High concentrations of CP cause necrosis, whereas lower concentrations induce apoptosis (19). In fact, apoptosis has been recognized as the main mechanism responsible for the antineoplastic action of CP (13). It is, therefore, not surprising that CP-induced apoptosis is important in nephrotoxicity, the major adverse effect of this agent.
The purpose of the present study was to evaluate the specific role of HO-1 in CP-induced renal injury in transgenic mice with targeted deletion of the HO-1 gene. In vitro studies were performed to assess the effects of chemical inducers and inhibitors of HO-1 on CP-mediated necrosis and apoptosis in human renal proximal tubule cells. To further corroborate our findings with chemical inducers and inhibitors of HO-1, we also performed studies in human renal epithelial cells to manipulate the HO-1 gene at the molecular level by selective overexpression.
![]() |
METHODS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Reagents
CP, hemin, fetal bovine serum, 3-(4,5 dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT), and blood urea nitrogen assay kit were obtained from Sigma Chemical (St. Louis, MO). Zinc protoporphyrin (ZnPP) was obtained from Porphyrin Products (Logan, UT). The cytotoxicity detection kit for lactate dehydrogenase (LDH) assay was obtained from Boehringer Mannheim (Indianapolis, IN), anti-HO-1 antibody (SPA-895) from StressGen Biotechnologies (Victoria, BC), lipofectamine from Life Technologies (Grand Island, NY), and the mammalian expression vector pcDNA3.1/Zeo and Zeocin from Invitrogen (Carlsbad, CA).Studies In Vivo
Study protocol.
Homozygous mice, HO-1 (/
), carrying a targeted deletion
of a large portion of the HO-1 gene, were selected from the offsprings of heterozygous/homozygous mating by Southern blotting of tail DNA
(31). Wild-type (+/+) and heterozygous (+/
) littermates were
used as controls. Mice from 12-16 wk of age were studied. CP, 20 mg/kg (1.0 mg/ml solution in sterile normal saline) or vehicle (normal
saline) was administered by a single intraperitoneal injection. Mice
were maintained on standard chow and tap water ad libitum. The
induction of acute renal failure was confirmed by increases in blood
urea nitrogen (BUN) and histological parameters of acute renal injury
(tubular necrosis, casts, loss of brush border, red blood cell
extravastion). The study protocol was approved by the Institutional
Animal Care and Use Committee at the University of Florida, Gainesville.
Apoptosis detection and quantification. Wax sections of 5-6 µm were placed on gelatin-coated slides and processed for the terminal deoxynucleotidyl transferase (TdT)-mediated dUTP nick end-labeling (TUNEL) assay by a modification of previously described methods (12). Briefly, after dewaxing and inactivation of endogenous peroxidases with hydrogen peroxide, sections were incubated for 1 h in 60°C dH2O. Subsequently, sections were incubated with the TdT reaction as described, and labeling was detected by using the avidin-biotin method. Counterstaining with methyl green for 3-5 min facilitated visualization of the sections. To localize the site of apoptotic nuclei, dual staining was performed by using an antibody to Na+-K+-ATPase alpha-1 subunit (Upstate Biotechnology, Waltham, MA). Analyses were performed in a blinded fashion. Random areas were viewed at a magnification of ×40 and scored for the number of apoptotic nuclei present in renal tubules in at least three to four slices of the kidney. An average of 266 ± 55 fields were examined, and the total number of apoptotic nuclei per square millimeter was calculated for each animal. For additional confirmation of apoptosis, transmission electron microscopy was performed from glutaraldehyde-fixed kidney tissue by previously described methods (15).
Studies In Vitro
Cell culture. Human renal proximal tubule cells (HPTC; Clonetics, Walkersville, MD) were maintained in renal epithelial basal medium supplemented with fetal bovine serum (5%), gentamicin (50 µg/ml), amphotericin B (50 µg/ml), insulin (5 µg/ml), transferrin (10 µg/ml), triiodothyronine (6.5 ng/ml), hydrocortisone (0.5 µg/ml), epinephrine (0.5 µg/ml), and human epidermal growth factor (10 ng/ml) at 37°C in 95% air-5% carbon dioxide. Studies were performed on cultures of up to four to five passages. Human embryonic kidney (HEK)-293 cells (ATCC, Rockville, MD) were maintained in Dulbecco's minimum essential medium supplemented with calf serum (10%) and HEPES (25 mM) at 37°C in 90% air-10% carbon dioxide.
Plasmid construction for HO-1 overexpression. A plasmid, pcDNA3.1/HHO-1, was constructed by using the mammalian expression vector pcDNA 3.1/Zeo and a 1.0-kb EcoR I/Xba I fragment containing the entire protein coding region of the HO-1 gene. The expression plasmid was designed to include the HO-1 cDNA encoding an expressed transgene mRNA ~300 bp smaller than the endogenous message. The vector alone was used as control. Transfections were done by using lipofectamine. Transfected cells were selected with Zeocin (100 µg/ml) for a week and incubated for 3 days without Zeocin before experiments. Successful overexpression was confirmed by Northern and Western analyses.
Northern analysis.
Total cellular RNA was extracted from transfected cells in 10-cm plates
by using the method of Chomczynski and Sacchi (7). The RNA was
electrophoresed on 1% agarose-formaldehyde gels, blotted onto nylon
membranes, and hybridized to a 32P-labeled cDNA probe for
human HO-1. To control for loading and transfer of RNA, the blots were
reprobed with a cDNA probe for human G3PDH. Autoradiography was
performed by using intensifying screens at 80°C.
Densitometric analysis was performed on a Power Macintosh computer by
using the National Institutes of Health Image 1.60 software (Bethesda, MD).
Western analysis. For HO-1 immunoblots, cells were washed twice with ice-cold PBS and lysed in Triton lysis buffer. Protein concentration of lysates was determined by bicinchoninic acid assay (Pierce). Samples (40 µg) were separated in a 10% SDS-polyacrylamide gel and then transferred onto a polyvinylidene difluoride membrane. The membranes were incubated for 1.5 h with the anti-HO-1 antibody (1:500 dilution) followed by incubation with peroxidase-conjugated goat anti-rabbit IgG antibody (1:10,000 dilution) for 1 h. Labeled protein bands were examined by using a chemiluminescence method according to the manufacturer's recommendation.
Cytotoxicity assays. Cytotoxicity was assessed by morphology on phase-contrast microscopy, specific LDH release, and the MTT assay for mitochondrial viability (27). For the LDH assay, HPTC were plated in 24-well plates and incubated at 37°C for 24 h. For HO-1 induction, cells were incubated with hemin (5 µM) for 12 h. The cells were washed and replaced with complete media (phenol red free) containing CP (100 µM) and/or ZnPP (5 µM) for 16 h. One hundred microliters of supernatant were transferred into 96-well plates and 200 µl of working solution containing diaphorase, NAD+, iodo-tetrazolium chloride, and sodium lactate were added. The reaction was stopped by the addition of 50 µl of 1 N hydrochloric acid. The absorbance of samples at 492 and 620 nm was measured. Specific LDH release was calculated by the absorbance of treated samples over control and Triton-X-treated samples. Transfected HEK-293 cells were plated in 24-well plates and incubated at 37°C for 24 h and then exposed to CP (100 µM) for 16 h, after which the LDH assay was performed as described above. To assess mitochondrial viability, transfected cells were plated into 96-well plates and incubated for 3 days, followed by exposure to media containing PBS (control, untreated) or CP (100 µM) for 16 h. Ten microliters of 5 mg/ml MTT stock solution were added, and cells were incubated for 4 h. One hundred microliters of acidic isopropanol (0.04 N hydrochloric acid in isopropanol) were added. The absorbance of samples at 550 and 690 nm was measured. Results are expressed as percent change in absorbance of CP-treated cells over control (untreated) cells in each group (vector alone and HO-1 transfected cells, respectively).
Quantification of apoptosis by using the annexin V binding assay. Binding of annexin V to the phosphatidylserine residues on the cell membrane in the presence of an intact cell membrane has been used as a marker for early apoptosis (9, 35). In these experiments, HPTC were pretreated with 5 µM hemin for 12 h, washed with Hanks' balanced salt solution, and exposed to CP (10 µM) in complete media for 5 days. To inhibit HO-1 activity, ZnPP (5 µM) was added, to cells pretreated with hemin, for 2 h before exposure to CP. Annexin V binding assay was performed by using an apoptosis-detection kit (R&D systems, Minneapolis, MN) for analysis of adherent cells as per the manufacturer's recommendations. Cells were harvested by gentle trypsinization and stained in suspension for immunofluorescence studies by using annexin V-FITC. The concentration of trypsin-EDTA used (0.025%) does not interfere with the annexin V assay, as shown by negative staining of control cells. Apoptotic cells were identified as cells staining positive for annexin V and negative for propidium iodide. Necrotic cells stained positive for both annexin V and propidium iodide. The number of apoptotic cells is expressed as a percentage of the total number of cells counted. An average of 700-800 cells was counted from multiple fields under both phase-contrast and fluorescence microscopy in each experimental group from at least three independent experiments.
Statistical analysis. Data are represented as means ± SE. For comparisons involving two groups, the t-test was used. For comparisons involving more than two groups, ANOVA and the Student-Newman-Keuls test were applied. All results are considered significant at P < 0.05.
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Assessment of Renal Function
All animals (+/+, +/
|
|
Morphological Evaluation and Quantification of Apoptosis In Vivo
Renal tissue sections in HO-1 (
|
|
|
Induction of HO-1 by Hemin Attenuates CP-Mediated Cytotoxicity in HPTC In Vitro
To evaluate the cytoprotective role of HO-1 in cytotoxicity induced by CP, HPTC were pretreated with a low dose of hemin (5 µM), a potent inducer of HO-1, 12 h before exposure to CP (100 µM). The maximal time point of HO-1 induction by hemin was determined in HPTC. As shown in Fig. 5A, maximal induction (~6-fold) was observed at 12 h after hemin treatment. Cytotoxicity was determined by phase-contrast microscopy and the LDH assay in these experiments. Exposure of HPTC to CP resulted in morphological evidence of cytotoxicity, as demonstrated by cell rounding, swelling, and detachment (Fig. 5B). Prior induction of HO-1, by hemin pretreatment, resulted in resistance of HPTC to CP-induced cytotoxicity (Fig. 5B). These data were corroborated by measurement of LDH activity, which demonstrated that hemin pretreatment significantly attenuated CP-induced cytotoxicity by 60% (CP alone: 16.1 ± 2.7%; hemin+CP: 6.35 ± 1.6%; P < 0.05; n = 4) at 12 h (Fig. 5C). The specificity of the cytoprotective effect of HO-1 was confirmed by cotreatment of cells with ZnPP (5 µM), an inhibitor of HO-1. Inhibition of HO-1 with ZnPP resulted in reversal of the cytoprotective effect conferred by hemin pretreatment (hemin+ CP+ZnPP: 15.4 ± 1.8%). Hemin and ZnPP alone demonstrated no significant cytotoxicity. The protective effect of hemin pretreatment was also seen after 24 h of exposure to CP (100 µM) (CP: 36.5%; hemin+CP: 17.6%; P < 0.05; n = 2).
|
Induction of HO-1 by Hemin Decreases CP-Mediated Apoptosis In Vitro
Previous studies by Lieberthal et al. (20) demonstrated that low concentrations of CP (~8-10 µM) cause apoptosis, whereas higher concentrations cause necrosis in mouse proximal tubule cells. In view of these observations, we explored the effects of HO-1 expression on CP-induced apoptosis in HPTC. Apoptosis was quantified by the annexin V binding assay. As shown in Fig. 6, exposure of HPTC to CP (10 µM) for 5 days induced apoptosis (C and D). Induction of HO-1 by hemin pretreatment significantly attenuated apoptosis (E and F). Inhibition of HO-1 by ZnPP, after hemin pretreatment, reversed the protective effect, resulting in more severe cellular injury with extensive necrosis (G and H). A quantitative representation of these results is summarized in Fig. 7. Incubation of cells with hemin or ZnPP (5 µM) alone caused no cell injury (apoptosis or necrosis).
|
|
Overexpression of HO-1 Decreases CP-Mediated Cell Injury
To further corroborate the results with chemical inducers and inhibitors, HO-1 was overexpressed in our cellular model of CP-induced cytotoxicity. HEK-293 cells were transfected with a vector containing the entire protein coding region of the human HO-1 gene (HHO-1) under the promoter/enhancer of cytomegalovirus (pcDNA3.1/HHO-1). The vector alone was used as a transfection control. Northern analysis confirmed the presence of successful overexpression (~40-fold) in the form of a ~1.5-kb message under basal, unstimulated conditions (Fig. 8A). Immunoblot analysis in cells transfected with pcDNA3.1/HHO-1 revealed an approximately eightfold increase in HO-1 protein over control cells transfected with vector alone (Fig. 8B).
|
Control and transfected cells were exposed to CP (100 µM) for 16 h,
and cytotoxicity was assessed by phase-contrast microscopy, LDH, and
MTT assays. Morphological indexes of injury, such as cell rounding,
vacuolization, and detachment, were significantly reduced in cells with
elevated levels of HO-1 compared with cells transfected with the vector
alone. These changes were accompanied by a 68% decrease in CP-induced
cytotoxicity as determined by the LDH release assay (vector alone: 20 ± 6.8%; HO-1 overexpression: 6.6 ± 2.82%; P < 0.02; n = 4) (Fig. 9A).
Treatment of cells with a competitive inhibitor, ZnPP, reversed the
cytoprotective effects of HO-1 (data not shown). Mitochondrial
viability, assessed by the MTT assay, revealed significantly decreased
CP-induced injury in cells transfected with HO-1. Cells transfected
with HO-1 and treated with CP had more viable mitochondria, represented
by a smaller change in absorbance, over control (untreated), HO-1
transfected cells (untreated, 100% vs. CP-treated, 94.7 ± 3.5%).
Cells transfected with vector alone had significantly greater
mitochondrial injury (untreated, 100% vs. CP-treated, 83.7 ± 4.9%).
The percent change in absorbance in vector alone vs. HO-1-transfected
cells is shown in Fig. 9B.
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Acute renal failure due to ischemia and nephrotoxic drugs
results in varying degrees of morphological damage, especially in the
S3 segment of the proximal tubule (5). Nephrotoxicity due to the
chemotherapeutic agent, CP, is characterized by similar morphological
changes in this segment of the nephron. In addition to injury in the S3
segment of the proximal tubule, morphological changes also occur in the
distal tubule, specifically the mTAL (5, 25). The results of our study
provide the first direct evidence that the expression of HO-1
determines CP-induced acute renal tubular injury in vivo and in
vitro. Homozygote (/
) mice that lack HO-1 develop
more severe renal failure and have a higher degree of renal tubular
apoptosis compared with wild-type (+/+) mice treated with CP.
Although the exact mechanisms of CP-induced renal toxicity have not
been defined, the toxicity is mediated, at least in part, via oxidant
mechanisms. CP decreases kidney glutathione content, increases lipid
peroxidation, interacts with DNA in a cell-free system, and generates
superoxide anion (23). Oxidant-scavenging enzymes and assorted
antioxidants protect against CP-induced renal injury (3). In addition,
induction of HO-1 occurs in the rat kidney in response to CP-induced
oxidant stress (1). It has been shown that treatment with iron
chelators (e.g., deferoxamine) prevents CP-induced lipid peroxidation
in the kidney (14). More recently, studies have demonstrated a role for
cytochrome P-450 and release of its catalytic iron as a
potential mediator of CP nephrotoxicity (4). The propensity of iron to
participate in the propagation of free radical generation via the
Haber-Weiss reaction and the recent demonstration that HO-1 is required
for the maintenance of iron homeostasis (31) implicate iron as a potential candidate molecule responsible for the injurious effects of
CP in the HO-1 (/
) mice.
Apoptosis of renal tubular cells plays an important pathogenic role in
CP nephrotoxicity as well as in other models of acute renal failure
(19, 20). Studies in mouse proximal tubular cells provided evidence
that CP-induced apoptosis is mediated via the generation of oxygen free
radicals, on the basis of the cytoprotective effect of antioxidants
such as catalase, superoxide dismutase, deferoxamine, and probucol
(20). In contrast, antioxidants did not protect against CP-induced
necrosis. Our in vitro studies showed significant attenuation
of CP-induced apoptosis as well as necrosis in human renal epithelial
cells by chemical inducers and overexpression of HO-1. Increased
apoptosis and necrosis were also observed in the HO-1 (/
)
mice. In this regard, the protection afforded by HO-1 expression is
unique compared with other antioxidants.
In our model of CP nephrotoxicity, a higher degree of apoptosis was
seen in both proximal and distal tubules of HO-1 (/
) mice
kidneys. These results are similar to observations in the p21-knockout
mouse model of CP-induced renal injury (25). Mice deficient in p21
(WAF1, CIP-1, or Sid-1), a cell-cycle inhibitory protein, develop a
rapid onset of renal failure and have a higher mortality after CP
administration compared with wild-type littermates (25). The mice also
have a higher degree of CP-induced apoptosis in proximal and distal
tubules, implicating the importance of cell-cycle regulation in the
pathogenesis of CP-induced renal failure. Renal epithelial cells
transfected with HO-1 have slower growth rates compared with cells
transfected with vector alone (Shiraishi F and Agarwal A,
unpublished observations), similar to findings in pulmonary
epithelial cells after overexpression of HO-1 (18). The cellular
mechanisms for this phenomenon are being evaluated in our laboratory.
It is tempting to extrapolate the effects of nitric oxide-induced cell
growth arrest mediated via activation of cGMP to carbon monoxide
released from the HO-1-catalyzed reaction, because carbon monoxide has
been shown to decrease proliferation of vascular smooth muscle cells
(26).
Our findings of severe CP-induced renal dysfunction in the HO-1
(/
) mice cannot be explained by increased renal tubular apoptosis alone. Most likely, the renal dysfunction involves a combination of increased apoptosis, necrosis, and possible vascular effects because of the absence of HO-1 expression (1, 11). Previous
studies have demonstrated that inhibition of HO-1 exacerbates changes
in renal hemodynamics in CP-induced nephrotoxicity (1). Foremost,
inhibition of HO-1 with tin protoporphyrin increases renal vascular
resistance and decreases renal blood flow in CP nephrotoxicity before
detectable changes in renal function. These effects could be
attributable to decreased generation of carbon monoxide, a metabolite
with vasodilatory effects mediated via cGMP (11). Second, increased
intracellular heme associated with CP nephrotoxicity may quench nitric
oxide, further exacerbating the vascular effects that occur during the
early stages in this model. Enhanced HO-1 activity would reverse these
deleterious vascular effects by generating carbon monoxide and
degrading intracellular heme, thereby providing a cytoprotective effect.
CP-induced mitochondrial injury (6) was significantly reduced by overexpression of HO-1 in our study. Recent studies suggest that mitochondria are the principle sensors and, thus, critical in the initiation of the apoptotic cascade (16, 32, 38). Mitochondrial dysfunction with liberation of cytochrome C, a mitochondrial heme protein, occurs as an early event in CP-induced apoptosis and results in activation of further downstream pathways (10). The processes involved in the antiapoptotic effects of HO-1 expression are not entirely clear. Although tubular injury in CP nephrotoxicity is not directly dependent on the delivery of heme proteins to the kidney, increased amounts of endogenous heme are observed in the kidney, specifically in the cytosolic fraction. This occurs as early as 6 h after CP administration and contributes, at least in part, to the induction of HO-1 and the injury that ensues (1). We speculate that CP-induced oxidant stress destabilizes heme proteins, resulting in the liberation of heme from its binding protein. The heme moiety, either directly or via the effects of its released iron, can damage a number of cellular targets including the lipid bilayer, the cytoskeleton, mitochondria, and the nucleus. Elevated levels of HO-1 will degrade the potentially prooxidant heme groups, released as a consequence of CP-induced oxidant stress and would limit the increase in heme that would otherwise occur. In addition to the induction of HO-1, administration of CP also leads to a concomitant increase in ferritin in the kidney. The increase in ferritin is an additional protective response, because ferritin safely sequesters the iron released from the heme ring (30). Other mechanisms by which the induction of HO-1 may confer protection include increased generation of bilirubin, a metabolite with potent antioxidant activity (8), and improved renal hemodynamics via generation of carbon monoxide (1). Studies to explore the cellular basis of the cytoprotective effects of HO-1 overexpression are presently underway in our laboratory.
Higher concentrations of heme (>50 µM) are cytotoxic whereas lower concentrations (5 µM) are cytoprotective, possibly via induction of HO-1. We do not think that hemin or a heme metabolite is inactivating CP for several reasons. First, in all our experiments where we used hemin pretreatment, the cell monolayer was washed and fresh media were added before CP exposure. Second, we have shown reversal of the cytoprotective effect with a competitive inhibitor of HO-1. Third, to obviate the concerns over chemical inducers and inhibitors, we have performed studies to selectively overexpress HO-1 in our cell culture model as well as used knockout mice to confirm our findings.
In summary, our study demonstrates that the expression of HO-1 protects renal epithelial cells against CP-induced cytotoxicity. CP nephrotoxicity, as assessed by structural and functional changes, was significantly greater in HO-1-knockout mice compared with wild-type mice. In contrast, induction of HO-1 by chemical inducers or by overexpression in renal epithelial cells in vitro protected cells against CP-induced cytotoxicity. Our data provide the impetus to design possible strategies for targeted gene expression of HO-1 as well as the development of novel, physiologically relevant inducers of the endogenous HO-1 gene as a therapeutic and preventive modality in high-risk settings of acute renal failure.
![]() |
ACKNOWLEDGEMENTS |
---|
We are grateful to Dr. Susumu Tonegawa for his contribution in generating the heme oxygenase-1-knockout mice. We thank Li Zhang and Feiyan Liu for excellent technical assistance.
![]() |
FOOTNOTES |
---|
This work was supported by National Institutes of Health Grants K08 DK-02446 (to A. Agarwal), HL-39593 (to H. S. Nick), DK-28330 (K. Madsen) and a pilot project grant from the Howard Hughes Medical Institute Research Resources Program, University of Florida (to A. Agarwal). F. Shiraishi was supported by a research fellowship from the American Heart Association of Florida.
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. §1734 solely to indicate this fact.
Address for reprint requests and other correspondence: A. Agarwal, Div. of Nephrology, Hypertension and Transplantation, Box 100224 JHMHC, 1600 SW Archer Rd., Univ. of Florida, Gainesville FL 32610 (E-mail: agarwal{at}nersp.nerdc.ufl.edu).
Received 7 July 1999; accepted in final form 9 December 1999.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1.
Agarwal, A,
Balla J,
Alam J,
Croatt AJ,
and
Nath KA.
Induction of heme oxygenase in toxic renal injury: a protective role in cisplatin nephrotoxicity in the rat.
Kidney Int
48:
1298-1307,
1995[ISI][Medline].
2.
Agarwal, A,
Kim Y,
Matas AJ,
Alam J,
and
Nath KA.
Gas-generating systems in acute renal allograft rejection in the rat: co-induction of heme oxygenase and nitric oxide synthase.
Transplantation
61:
93-98,
1996[ISI][Medline].
3.
Baliga, R,
Ueda N,
Walker PD,
and
Shah SV.
Oxidant mechanisms in toxic acute renal failure.
Am J Kidney Dis
29:
465-477,
1997[ISI][Medline].
4.
Baliga, R,
Zhang Z,
Baliga M,
Ueda N,
and
Shah SV.
Role of cytochrome P-450 as a source of catalytic iron in cisplatin-induced nephrotoxicity.
Kidney Int
54:
1562-1569,
1998[ISI][Medline].
5.
Bonventre, JV,
Brezis M,
Siegel N,
Rosen S,
Portilla D,
and
Venkatachalam M.
Acute renal failure. I. Relative importance of proximal vs. distal tubular injury.
Am J Physiol Renal Physiol
275:
F623-F632,
1998
6.
Brady, HR,
Kone BC,
Stromski ME,
Zeidel ML,
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
7.
Chomczynski, P,
and
Sacchi N.
Single step method of RNA isolation by acid guanidinium thiocyanate-phenol-chloroform extraction.
Anal Biochem
162:
156-159,
1987[ISI][Medline].
8.
Dore, S,
Takahashi M,
Ferris CD,
Hester LD,
Guastella D,
and
Snyder SH.
Bilirubin, formed by activation of heme oxygenase-2, protects neurons against oxidative stress injury.
Proc Natl Acad Sci USA
96:
2445-2450,
1999
9.
Engeland, MV,
Nieland LJW,
Ramaekers FCS,
Schutte B,
and
Reutelingsperger CPM
Annexin V-affinity assay: a review on an apoptosis detection system based on phosphatidylserine exposure.
Cytometry
31:
1-9,
1998[ISI][Medline].
10.
Fukuoka, K,
Takeda M,
Kobayashi M,
Osaki T,
Shirato I,
Soejima A,
Nagasawa T,
and
Endou H.
Distinct interleukin-1-converting enzyme family of proteases mediate cisplatin- and staurosporine-induced apoptosis of mouse proximal tubule cells.
Life Sci
62:
1125-1138,
1998[ISI][Medline].
11.
Furchgott, RF,
and
Jothianandam D.
Endothelial-dependent and independent vasodilatation involving cGMP: relaxation induced by nitric oxide, carbon monoxide and light.
Blood Vessels
28:
52-61,
1991[ISI][Medline].
12.
Gavrieli, Y,
Sherman Y,
and
Ben-Sasson SA.
Identification of programmed cell death in situ via specific labeling of nuclear DNA fragmentation.
J Cell Biol
119:
493-501,
1992[Abstract].
13.
Jiang, S,
Song MJ,
Shin EC,
Lee MO,
Kim SJ,
and
Park JH.
Apoptosis in human hepatoma cell lines by chemotherapeutic drugs via Fas-dependent and Fas-independent pathways.
Hepatology
29:
101-110,
1999[ISI][Medline].
14.
Kameyama, Y,
and
Gemba M.
The iron chelator, deferoxamine, prevents cisplatin-induced lipid peroxidation in rat kidney cortical slices.
Jpn J Pharmacol
57:
259-262,
1991[ISI][Medline].
15.
Kim, J,
Lee GS,
Tisher CC,
and
Madsen KM.
Role of apoptosis in development of the ascending thin limb of the loop of Henle in rat kidney.
Am J Physiol Renal Fluid Electrolyte Physiol
271:
F831-F845,
1996
16.
Kluck, RM,
Bossy-Wetzel E,
Green DR,
and
Newmeyer DD.
The release of cytochrome c from mitochondria: a primary site for Bcl-2 regulation of apoptosis.
Science
275:
1132-1136,
1997
17.
Lebwohl, D,
and
Canetta R.
Clinical development of platinum complexes in cancer therapy: an historical perspective and an update.
Eur J Cancer
34:
1522-1534,
1998[ISI][Medline].
18.
Lee, PJ,
Alam J,
Wiegand GW,
and
Choi AMK
Overexpression of heme oxygenase-1 in human pulmonary epithelial cells results in cell growth arrest and increased resistance to hyperoxia.
Proc Natl Acad Sci USA
93:
10393-10398,
1996
19.
Lieberthal, W,
Koh JS,
and
Levine JS.
Necrosis and apoptosis in acute renal failure.
Semin Nephrol
18:
505-518,
1998[ISI][Medline].
20.
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
21.
Liu, H,
Mount DB,
Nasjletti A,
and
Wang W.
Carbon monoxide stimulates the apical 70-pS K+ channel of the rat thick ascending limb.
J Clin Invest
103:
963-970,
1999
22.
Maines, MD.
The heme oxygenase system: a regulator of second messenger gases.
Annu Rev Pharmacol Toxicol
37:
517-554,
1997[ISI][Medline].
23.
Masuda, H,
Tanaka T,
and
Takahama U.
Cisplatin generates superoxide anion by interaction with DNA in a cell-free system.
Biochem Biophys Res Commun
203:
1175-1180,
1994[ISI][Medline].
24.
McCoubrey, WK,
Huang TJ,
and
Maines MD.
Isolation and characterization of a cDNA from the rat brain that encodes hemoprotein heme oxygenase-3.
Eur J Biochem
247:
725-732,
1997[Abstract].
25.
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
26.
Morita, T,
Mitsialis SA,
Koike H,
Liu Y,
and
Kourembanas S.
Carbon monoxide controls the proliferation of hypoxic vascular smooth muscle cells.
J Biol Chem
272:
32804-32809,
1997
27.
Mosmann, T.
Rapid colorimetric assay for cellular growth and survival: application to proliferation and cytotoxicity assays.
J Immunol Methods
65:
55-63,
1983[ISI][Medline].
28.
Nath, KA.
Tubulointerstitial changes as a major determinant in the progression of renal damage.
Am J Kidney Dis
20:
1-17,
1992[ISI][Medline].
29.
Nath, KA,
Balla G,
Vercellotti GM,
Balla J,
Jacob HS,
Levitt MD,
and
Rosenberg ME.
Induction of heme oxygenase is a rapid, protective response in rhabdomyolysis in the rat.
J Clin Invest
90:
267-270,
1992[ISI][Medline].
30.
Platt, JL,
and
Nath KA.
Heme oxygenase: protective gene or Trojan horse.
Nat Med
4:
1364-1365,
1998[ISI][Medline].
31.
Poss, KD,
and
Tonegawa S.
Heme oxygenase 1 is required for mammalian iron reutilization.
Proc Natl Acad Sci USA
94:
10919-10924,
1997
32.
Reed, JC.
Cytochrome C: can't live with it-can't live without it.
Cell
91:
559-562,
1997[ISI][Medline].
33.
Ries, F,
and
Klastersky J.
Nephrotoxicity induced by cancer chemotherapy with special emphasis on cisplatin toxicity.
Am J Kidney Dis
8:
368-379,
1986[ISI][Medline].
34.
Safirstein, R,
Winston J,
Goldstein M,
Model D,
Dikman S,
and
Guttenplan J.
Cisplatin nephrotoxicity.
Am J Kidney Dis
8:
356-367,
1986[ISI][Medline].
35.
Savill, J.
Recognition and phagocytosis of cells undergoing apoptosis.
Br Med Bull
53:
491-508,
1997[Abstract].
36.
Soares, MP,
Lin Y,
Anrather J,
Csizmadia E,
Takigami K,
Sato K,
Grey ST,
Colvin RB,
Choi AM,
Poss KD,
and
Bach FH.
Expression of heme oxygenase-1 can determine cardiac xenograft survival.
Nat Med
4:
1073-1077,
1998[ISI][Medline].
37.
Stocker, R.
Induction of haem oxygenase as a defence against oxidative stress.
Free Radic Res Commun
9:
101-112,
1990[ISI][Medline].
38.
Takeda, M,
Kobayashi M,
Shirato I,
Osaki T,
and
Endou H.
Cisplatin-induced apoptosis of immortalized mouse proximal tubule cells is mediated by interleukin-1 converting enzyme (ICE) family of proteases but inhibited by overexpression of Bcl-2.
Arch Toxicol
71:
612-621,
1997[ISI][Medline].
39.
Vogt, BA,
Shanley TP,
Croatt A,
Alam J,
Johnson KJ,
and
Nath KA.
Glomerular inflammation induces resistance to tubular injury in the rat. A novel form of acquired, heme oxygenase-dependent resistance to renal injury.
J Clin Invest
98:
2139-2145,
1996