High glucose-induced oxidative stress causes apoptosis in

The FASEB Journal express article10.1096/fj.02-0130fje. Published online March 28, 2003.
High glucose-induced oxidative stress causes apoptosis in
proximal tubular epithelial cells and is mediated by multiple
caspases
David A. Allen, Steven M. Harwood, Mira Varagunam, Martin J. Raftery, and Muhammad M.
Yaqoob
Department of Experimental Medicine & Nephrology, William Harvey Research Institute, St.
Bartholomew’s & Royal London School of Medicine and Dentistry, Queen Mary, University of
London, UK
Corresponding author: David A. Allen, Department of Experimental Medicine and Nephrology,
Suite 22, Dominion House, 59 Bartholomew Close, West Smithfield, London EC1A 7BE UK. Email: [email protected]
ABSTRACT
Diabetic nephropathy is the leading cause of end-stage renal disease in the Western world. Poor
glycemic control contributes to the development of diabetic nephropathy, but the mechanisms
underlying high glucose-induced tissue injury are not fully understood. In the present study, the
effect of high glucose on a proximal tubular epithelial cell (PTEC) line was investigated.
Reactive oxygen species (ROS) were detected using the fluorescent probes dichlorofluorescein
diacetate, dihydrorhodamine 123, and 2,3-diaminonapthalene. Peroxynitrite (ONOO–) generation
and nitrite concentrations were increased after 24 h of high glucose treatment (P<0.05). LLCPK1 cells exposed to high D-glucose (25 mM) for up to 48 h had increased DNA fragmentation
(P<0.01), caspase-3 activity (P<0.001), and annexin-V staining (P<0.05) as well as decreased
expression of XIAP when compared with controls (5 mM D-glucose). The ONOO– scavenger
ebselen reduced DNA fragmentation and caspase-3 activity as well as the high glucose-induced
nitrite production and DCF fluorescence. High glucose-induced DNA fragmentation was
completely prevented by an inhibitor of caspase-3 (P<0.01) and a pan-caspase inhibitor
(P<0.001). Caspase inhibition did not affect ROS generation. This study, in a PTEC line,
demonstrates that high glucose causes the generation of ONOO–, leading to caspase-mediated
apoptosis. Ebselen and a caspase-3 inhibitor provided significant protection against high glucosemediated apoptosis, implicating ONOO– as a proapoptotic ROS in early diabetic nephropathy.
Key words: diabetic nephropathy • peroxynitrite • cell death • ebselen
D
iabetic nephropathy is a serious complication for patients with diabetes mellitus (DM).
Approximately 30–40% of patients with type I and 15% with type II DM develop endstage renal disease (1, 2). Tubular dysfunction has been reported in DM in the absence
of microabuminuria, which suggests a pivotal role of tubulo-interstitium in the development of
diabetic nephropathy (3, 4). However, the exact mechanism underlying tubular dysfunction in
DM remains unknown. High glucose concentrations have been implicated as a causal factor in
the initiation and progression of diabetic nephropathy (5–7), and there is evidence to suggest that
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hyperglycemia increases the production of free radicals and oxidant stress (8–10). In addition,
glucose can undergo autooxidation, leading to the production of intermediates that result in
reactive oxygen species (ROS) generation and may form adducts in proteins to form advanced
glycation end products (AGEs) or Amadori products (10, 11). Overproduction of superoxide (O2–
) and nitric oxide (NO!) has been reported to occur in response to high glucose concentrations
both in vivo (12, 13) and in vitro (14–16). Coproduction of NO! and O2– in the same cellular
compartment would create favorable conditions for a reaction between them. The product of the
reaction is peroxynitrite (ONOO–), a powerful oxidant that causes nitration of tyrosine residues
in proteins and adversely affects their function (17).
ONOO– can undergo protonation, yielding peroxynitrous acid and ultimately hydroxyl radicals or
oxidation by its reaction with CO2-generating nitrosoperoxycarbonate. Damaging effects of
ONOO– include oxidative DNA damage such as point mutation and double-strand breaks as well
as lipid peroxidation (18–20). ONOO– activated several caspases in the HL-60 leukemic cell line,
although only caspase-3 was essential for apoptosis (21, 22). These studies indicate the
mechanisms by which ONOO–, added directly to cell culture, can promote apoptosis in vitro.
Numerous studies have provided evidence of a role for ONOO– in diabetic complications in vivo
and in vitro. The presence of 3-nitrotyrosine in tissue sections is a reliable indicator of ONOO–induced oxidative stress and cellular injury in vivo. A recent study showed increased
nitrotyrosine staining in the proximal tubules of patients with diabetic renal disease that was not
evident in those with renal disease originating from other causes, suggesting ONOO–-induced
injury in diabetic nephropathy (23). Similarly, nitrotyrosine was detected in epithelial cells
following endotoxin-induced kidney injury in the rat (24) and in the renal cortex of rats with DM
(13). In a study by Ceriello et al. (25), nitrotyrosine was detected in the plasma of diabetic
patients but not in the plasma of healthy controls. In addition, there is evidence for ONOO–
generation in islet β-cells in diabetic NOD mice (26), suggestive of oxidative stress. Treatment
of NOD mice with a ONOO– scavenger was able to prevent the development of DM (12). The
role of glucose in oxidative stress-induced kidney injury has been previously documented (27–
29). Glucose-induced oxidative stress has been reported to inhibit proximal tubular epithelial cell
(PTEC) proliferation via activation of protein kinase C (PKC) and the transforming growth
factor-β1 (TGFβ) signaling pathway (30). Furthermore, ROS-induced TGFβ promotes fibrotic
injury in mesangial cells (31) and glomerular endothelial cells (32). In the present study, we
focused on the effect of high ambient glucose concentrations, and the resultant oxidative stress,
on apoptotic pathways in a PTEC line. Furthermore, we have attempted to define a role for nitric
oxide-derived ROS in high glucose-induced cell injury. Our data support the view that a
proapoptotic effect of high glucose is primarily mediated via increased ONOO–generation and
involves multiple caspases.
MATERIALS AND METHODS
Cell culture and chemicals
The porcine PTEC line LLC-PK1 was purchased from the European Collection of Animal Cell
Cultures (ECACC, Porton Down, UK). LLC-PK1 cells were grown on 75 cm3 tissue culture
flasks in Dulbecco’s modified Eagle’s medium containing 10% v/v fetal calf serum plus
antibiotics (100 U/ml penicillin, 100 µg/ml streptomycin, and 0.25 µg/ml amphotericin B).
Subconfluent LLC-PK1 cells were harvested and seeded into a six-well tissue culture plate in 1
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ml of growth medium. The cells were allowed to adhere for 18 h in a humidified incubator at
37°C with 5% CO2 in air. Immediately before treatment, the medium was removed and replaced
with fresh medium containing 5 mM D-glucose plus 20 mM L-glucose or 25 mM D-glucose.
Inhibitors or free-radical scavengers were added simultaneously with low or high D-glucose
media. Ebselen (2-phenyl-1,2-benzisoselenazol-3[2H]-one) (CN Biosciences, Nottingham, UK)
was used as a ONOO– scavenger. The cell-permeable caspase-3 inhibitor DEVD-CHO, the
caspase-8 inhibitor IETD-CHO (CN Biosciences), and the pan-caspase inhibitor Z-Asp-2,6dichlorobenzoyloxymethylketone (Z-D-DCB, Bachem, Saffron Walden, UK) were used at 1.25
µM and 100 µM, respectively. N-[3-(aminomethyl)benzyl]acetamidine (1400W), catalase
(purified from Aspergillus niger), and 6-hydroxy-2,5,7,8-tetramethylchroman-2-carboxylic acid
(Trolox) were purchased from CN Biosciences. Unless stated, all other chemicals, including Lglucose (mixed anomers) and D-glucose, were from Sigma-Aldrich (Poole, UK).
Analysis of DNA fragmentation
Monolayers of LLC-PK1 cells were disrupted with a Teflon cell scraper and placed in phosphatebuffered saline (PBS). Cells were centrifuged at 2000g, and the pellet was stored at –80°C. For
the DNA fragmentation ELISA, cell pellets (~2×106 cells) were incubated for 30 min on ice in
200 µl of lysis buffer (PBS, pH 7.4, containing 10 µM digitonin). After centrifugation at 16,000g
for 15 min at 4°C, the supernatant was diluted 1:10 with lysis buffer. The diluted lysate (20 µl)
was assayed in duplicate for microsomal DNA fragments by using a commercial ELISA (Cell
Death Detection ELISA Plus, Roche Diagnostics, Lewes, UK). Total protein was measured by
the Bradford microprotein assay (Bio-Rad, Hemel Hempstead, UK). Results of the DNA
fragmentation ELISA are expressed as a multiple of control normalized to the amount of protein
in the lysate.
Measurement of annexin V staining
Subconfluent monolayers of LLC-PK1 cells were exposed to media containing either 5 mM or 25
mM D-glucose for 24 h. Detection of annexin V on the cell surface was performed by flow
cytometry, using a Becton Coulter EPICS flow cytometer. System II software was used for
acquisition and analysis. Annexin V-FITC conjugate (R&D Systems, Abingdon, UK) was used
to stain cells. In brief, cells were harvested, washed, and incubated for 15 min with annexin VFITC and propidium iodide. This combination allows for the differentiation between early
apoptotic cells (annexin V positive), late apoptotic and/or necrotic cells (annexin V and
propidium iodide positive), and viable cells (unstained). Results (±SD) are expressed as fold of
control.
Measurement of caspase activity
The activity of caspase-3 was measured using the fluorimetric substrate Ac-DEVD-AMC. In
brief, ~50 µg of cellular protein was incubated with 2.5 mM substrate in caspase assay buffer
(312.5 mM HEPES, pH 7.5, 31.25% sucrose, and 0.3125% CHAPS) for 1 h, and fluorescence
was measured on a microplate reader (Fluostar Galaxy, BMG Laboratory Technologies,
Aylesbury, Bucks, UK), with excitation at 380 nm and emission at 460 nm. For each sample, a
negative control is assayed in parallel containing 2.5 mM of the appropriate caspase inhibitor
(Ac-DEVD-CHO). The activities of caspases-8 and -9 were measured in the same way, using
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Ac-IETD-AMC and Ac-LEHD-AMC as substrates with Ac-IETD-CHO and Ac-LEHD-CHO as
inhibitors, respectively. Fluorescence readings from wells containing inhibitor were subtracted
from total fluorescence, and the results were calculated as nmol AMC/mg/min and expressed as
a multiple of control (LLC-PK1 cells grown in 5 mM D-glucose).
Western blotting and detection of 3-nitrotyrosine and XIAP
For Western blot analysis, 10 µg of whole-cell lysate was electrophoresed on a 10% acrylamide
gel in the presence of sodium dodecyl sulfate and electroblotted onto a polyvinylidene fluoride
membrane. After they were blocked overnight at 4°C with 5% w/v bovine serum albumin,
membranes were probed with a monoclonal anti-nitrotyrosine antibody (Clone 1A6, Upstate
Biotech, Milton Keynes, Bucks, UK) or anti-XIAP (1:500, R&D Systems). The secondary
antibody was horseradish peroxidase-labeled IgG (1:2000, Santa Cruz Biotechnology, Santa
Cruz, CA), and detection was performed using enhanced chemiluminescence (ECL, Amersham
Biosciences, Little Chalfont, UK).
Detection of ONOO–
The cell-permeable fluorogenic probe 2'7'-dichlordihydrofluorescin diacetate (DCF-DA) was
used to detect oxidative stress in LLC-PK1 cells. DCF-DA diffuses across cell membranes and is
hydrolyzed by nonspecific cellular esterases to the nonfluorescent compound dichlorofluorescin
(DCFH), which is predominantly trapped within the cell. In the presence of ROS, DCFH rapidly
undergoes one-electron oxidation to the highly fluorescent compound dichlorofluorescein (DCF).
The assay is a modification of that described by Bestwick and Milne (33). Glucose-exposed cell
monolayers are incubated with 20 µM DCF-DA (CN Biosciences, Nottingham, UK) in PBS, pH
7.4, for 30 min at 37°C. The cells were washed once with PBS and lysed in PBS containing 0.1%
Triton X-100 and 0.5 mM EDTA. Lysates were collected and cleared of cell debris by
centrifugation at 14,000g for 5 min. The formation of DCF was monitored in a fluorescent
microplate reader with emission at 520 nm and excitation at 500 nm. Results were normalized to
total cell protein and expressed as fold of control.
Dihydrorhodamine 123 (DHR; Sigma-Aldrich) was prepared as a 10% stock solution in
dimethylformamide. DHR forms the fluorescent product rhodamine 123 upon oxidation by
peroxides and notably ONOO–. Monolayers of LLC-PK1 cells were exposed to 5 mM or 25 mM
D-glucose for 24 h, washed twice in PBS (pH 7.4), and incubated in PBS containing 5 µM DHR
for 30 min at 37°C. Aliquots of supernatant (duplicate samples of 100 µL) were removed at
regular intervals, and fluorescence was detected with emission at 530 nm and excitation at 485
nm. Formation of rhodamine 123 is calculated as fluorescence units per milligram cell protein
and expressed as a fold of control.
Nitrite was measured using the fluorescent probe 2′,3′-diaminonapthalene (DAN) essentially as
previously described (34). In brief, 100 µl of culture supernatant was incubated in a 96-well
microtiter plate with 30 µL of buffer containing 1 U/ml nitrite reductase, 200 µM NADPH, and
50 µM FAD for 15 min at 37°C. Ten microliters of DAN (0.05 mg/mL) were added, and the
plate was incubated for an additional 10 min in the dark followed by neutralization with 10 µl
1.4 N NaOH. The fluorescence was then measured (excitation at 380 nm and emission at 460
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nm) using sodium nitrite as a standard. Results were calculated as nitrite per milligram total cell
protein per hour and expressed as fold of control.
Statistical analysis
Results of three or more independent experiments are expressed as mean ±SD/SE, and statistical
comparison of two groups was performed using the Student’s t test for paired samples or the
Wilcoxon’s signed ranks test for paired, nonnormally distributed data. Unpaired, nonnormally
distributed data were compared using the Mann Whitney U test. When comparing the means of
two groups, P<0.05 was considered significant.
RESULTS
High glucose causes oxidative stress in LLC-PK1 cells by increasing ONOO– generation
Control cultures containing 5 mM D-glucose were routinely supplemented with 20 mM Lglucose to ensure isosmolar conditions to cultures containing 25 mM D-glucose. LLC-PK1 cells
cultured in monolayers were exposed to 25 mM D-glucose or 5 mM D-glucose as a control for 24
and 48 h. Nitrite concentrations were measured in supernatants taken from cell monolayers after
24 h and normalized to total cellular protein content. Nitrite concentrations were significantly
increased in the supernatants of LLC-PK1 cells exposed to high glucose when compared with
controls (n=4, P<0.05; Fig. 1A). This effect was time- and glucose concentration-dependent
(fold of control: 5 mM 16 h=1.07±0.06, 5 mM 24 h=1.11±0.09, 15 mM 16 h=1.16±0.12, 15 mM
24 h=1.13±0.05, 25 mM 16 h=1.26±0.14*, 25 mM 24 h=1.33±0.06*, n=6; *P<0.05). Ebselen
caused a 40% reduction in nitrite concentration when compared with cells exposed to high
glucose alone (n=6, P<0.05, Fig. 1A). A NOS inhibitor (1400 W, 50 µM) did not significantly
alter the nitrite concentration in high glucose-exposed LLC-PK1 cells (n=6, Fig. 1A). When
compared with the effect of ebselen, this suggests that the majority of the nitrite is either
generated by decomposition of ONOO– or by reduction of nitrate in the medium and not directly
from NOS. This was confirmed by the addition of synthetic ONOO– to LLC-PK1 cells. After 24
h, there was an increase in nitrite in ONOO–-treated cells when compared with controls that had
been incubated with degraded ONOO– (5 mM+ONOO–=1.28±0.02-fold of control, 25
mM+ONOO–=1.61±0.14-fold of control, n=3, both P<0.05).
H2-DCF-DA was used to detect intracellular ONOO– generation. There was a residual amount of
fluorescence in control cultures that showed a small increase over the study period. However,
when the cells were exposed to 25 mM D-glucose, there was a large increase in fluorescence,
indicating a marked increase in ROS generation (n=9, P<0.02; Fig. 1B), measured 24 h after
exposure to high glucose. The fluorescence declined by 48 h, suggesting that ROS are consumed
by reaction with target molecules or antioxidant defences. The identity of the ROS detected by
this method is not exclusively ONOO– because DCFH can also emit fluorescence upon reaction
with hydroperoxides. However, the fact that fluorescence is reduced in the presence of ebselen
provides some evidence that the ROS is ONOO– or a derivative (n=8, P<0.05, Fig. 1B). In
addition, catalase (500 U/mL) did not significantly reduce high glucose-induced fluorescence,
which would exclude the production of hydrogen peroxide as a cause (fold of control: 25 mM Dglucose, 1.44±0.30; 25 mM D-glucose+catalase, 1.34±0.35; n=3, P=not significant when
compared with 25 mM D-glucose alone). We then used another antioxidant to further define the
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nature of the oxidant generated in high glucose. Trolox (100 µM in 50 mM phosphate buffer, pH
7.4), a water-soluble vitamin E analog that can undergo one-electron oxidation by ONOO–,
caused a significant reduction in DCF fluorescence (n=6, P<0.05, Fig. 1C).
DHR was used as an additional marker of ONOO– formation and yielded similar results to those
obtained with H2-DCF-DA. In cultures of LLC-PK1 cells incubated with 25 mM D-glucose,
DHR-associated fluorescence was increased by 28% (25 mM=1.28±0.06-fold of control, 5 mM,
n=6, P<0.05). Taken together, these data demonstrate that increased oxidative and/or nitrosative
stress is evident in LLC-PK1 cells exposed to 25 mM D-glucose for up to 24 h. The accumulation
of nitrotyrosine in cells and tissues is an additional marker of peroxynitrite-induced oxidative
stress. Western blot analysis of whole-cell lysates from LLC-PK1 cells exposed to 5 mM or 25
mM D-glucose showed that there was significant nitrotyrosine staining in cultures containing 25
mM D-glucose after 24 h (Fig. 2, lanes 2 and 3).
Exposure of LLC-PK1 cells to high glucose causes apoptosis
In cultures of LLC-PK1 cells supplemented with 20 mM L-glucose, DNA fragmentation was
similar to that in cultures containing only 5 mM D-glucose, indicating that any increase in DNA
fragmentation was not due to an osmotic effect. LLC-PK1 cells exposed to 25 mM D-glucose for
48 h showed increased DNA fragmentation when compared with cells exposed to 5 mM Dglucose (n=4, P<0.02, Fig. 3A). DNA fragmentation was ~2.5-fold higher in high glucoseexposed cells and was maximal at 48 h. In addition to DNA fragmentation, annexin V staining
was used as an early marker of apoptosis. LLC-PK1 cells cultured for 24 h in media containing
25 mM D-glucose showed significantly greater numbers of annexin V-positive cells than those
exposed to low D-glucose (n=3, P<0.05; Fig. 3B).
The same lysates in which DNA fragmentation was measured were used to assay caspase-3
activity, and the results are shown in Fig. 4A. There was a significant increase in caspase-3
activity in LLC-PK1 cells grown in 25 mM D-glucose when compared with those grown in 5 mM
D-glucose (n=9, P<0.05). The increase in caspase-3 activity was maximal at 48 h but was also
elevated at 24 h. To define a mechanism of caspase-3 activation, we assayed the same lysates for
caspase-8 and caspase-9 activity, using the substrates Ac-IETD-AMC and Ac-LEHD-AMC,
respectively. Caspase-8 activity was increased when compared with controls but did not reach
significance (n=6, Fig. 4B), whereas caspase-9 activity was significantly increased in high
glucose-exposed LLC-PK1 cells when compared with controls (n=3, P<0.01, Fig. 4C). Caspase-9
activity was maximal after 24 h. To define a causal role for ONOO– in caspase-mediated
apoptosis in LLC-PK1 cells, we measured caspase-3 activity in lysates from cells incubated for
24 h with 100 µM ONOO– or degraded ONOO– as a control. ONOO– significantly increased
caspase-3 activity (ONOO–=1.35±0.14-fold of control compared with degraded ONOO–
=1.00±0.04, n=4, P<0.05). In addition, the expression of X-linked inhibitor of apoptosis (XIAP,
an endogenous inhibitor of caspases) was decreased in whole-cell lysates from LLC-PK1 cells
incubated with 25 mM D-glucose at 24 and 48 h when compared with controls (Fig. 5).
Effect of a ONOO– scavenger on proximal tubular cell injury
The seleno-methionine compound ebselen was used as a scavenger of ONOO–. Previous studies
had determined that the optimal concentration of ebselen was between 10 and 20 µM. Ebselen
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(10 µM), when present throughout the experiment, reduced high glucose-induced DNA
fragmentation by ~30% when compared with controls (n=6, P<0.05, Fig. 6A). To define the
mechanism of this effect, we cultured LLC-PK1 cells in 5 mM and 25 mM D-glucose with or
without ebselen and measured caspase-3 activity on whole-cell lysates. High glucose-induced
caspase-3 activity was reduced by ∼25% in the presence of ebselen (48 h, n=3, P<0.05; Fig. 6B),
suggesting that in renal PTECs exposed to high ambient glucose concentrations, peroxynitrite
activates caspase-3. Additional experiments revealed that ebselen also reduced the high glucoseinduced activation of caspase-9 (fold of control [5 mM D-glucose]: 5 mM D-glucose+ebselen,
0.84±0.09; 25 mM D-glucose, 1.7±0.22; 25 mM D-glucose+ebselen, 1.41±0.24; n=3, P<0.05).
Inhibition of caspase activity prevents high glucose-induced DNA fragmentation
The data previously described demonstrate that the activities of caspase-9 and caspase-3 are
increased in LLC-PK1 cells exposed to high ambient glucose concentrations. To further define
the role of caspases in high glucose-mediated apoptosis, we incubated LLC-PK1 cells with the
cell-permeable caspase inhibitors DEVD-CHO (caspase-3 inhibitor), IETD-CHO (caspase-8
inhibitor), and Z-Asp-DCB (pan-caspase inhibitor). The optimum concentration of the caspase-3
inhibitor (DEVD-CHO, 1.25 µM) was confirmed by titration to ensure that it did not cause
inhibition of caspase-9 (fold of control caspase-9 activity: 25 mM, 1.61±0.21; 25 mM+1.25 µM
DEVD-CHO, 1.85±0.13; n=6, P=not significant). High glucose-induced caspase-3 activity was
abolished by all caspase inhibitors (48 h, n=6, P<0.001; Fig. 7A). Similarly, all three caspase
inhibitors reduced high glucose-induced DNA fragmentation (caspase-3 inhibitor, n=8, P<0.05;
caspase-8 inhibitor, n=4, P<0.05), although the caspase-3 and -8 inhibitors were less effective
than the pan-caspase inhibitor, which afforded complete protection against high glucose-induced
DNA fragmentation (n=8, P<0.005; Fig. 7B). To rule out the possibility that the caspase
inhibitors can scavenge ROS, we tested the effect of each of the caspase inhibitors on H2-DCFDA-induced fluorescence. The caspase inhibitors or DMSO had no affect on the level of
fluorescence in LLC-PK1 cells exposed to 25 mM D-glucose. Thus, caspase-3 is activated
predominantly by a mechanism involving caspase-9 but may also involve caspase-8, and a
caspase cascade mediates the apoptotic response to high glucose in LLC-PK1 cells.
DISCUSSION
In this study, we have shown that PTECs rapidly undergo apoptotic cell death following
exposure to high glucose. Detection of annexin V on the surface of LLC-PK1 cells was used as
an early apoptotic marker and at 24 h was significantly increased in high glucose-exposed cells
when compared with controls. In agreement with this finding, DNA fragmentation, a relatively
late event in apoptosis, was maximally increased after 48 h of exposure to high glucose. We have
provided evidence that oxidative stress and subsequent caspase activation are pivotal steps that
lead to an apoptotic cascade in PTECs. High glucose stimulated nitrite production and increased
oxidative stress in PTECs. Significant inhibition of nitrite generation by ebselen implies that the
greater fraction of nitrite is generated by decomposition of ONOO–. High glucose inhibits NOS
activity in mesangial cells by inhibiting the availability (or binding) of tetrahydrobiopterin
(BH4), an important cofactor for NOS enzymes (35). This appears to contrast with our data in
PTECs in which nitrite concentration is increased in high glucose. One possible explanation is
that high glucose causes a relative deficiency of BH4 in PTECs. NOS can carry out uncoupled
electron transfer in the absence of BH4, resulting in O2– generation instead of NO! in a similar
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manner to the reaction that occurs when L-arginine is lacking. The concomitant generation of O2–
by NOS would consume NO! in the generation of ONOO–, leading to reduced bioavailability of
NO!. A similar mechanism has been proposed by Tilton et al. (36) in which glucose-induced O2–
generation, via the sorbitol pathway, leads to an increase in intracellular calcium that activates
NOS. This would lead to coproduction of O2– and NO! and facilitate ONOO– generation. In
another study, high glucose activated nuclear transcription factor (NF)-κB and caused apoptosis
in human umbilical vein endothelial cells that was prevented by an antisense oligonucleotide
directed against NF-κB (14). Hyperglycemia can cause an increase in the production of O2–,
hydrogen peroxide, and hydroxyl radical (37, 38), possibly by reactions involving transition
metals. In addition, autooxidation of glycated proteins may yield hydrogen peroxide and other
ROS (39). Thus, O2– can be produced either by the uncoupled reactions of NOS or by other
unidentified mechanisms in hyperglycemia and, by its reaction with NO!, lead ultimately to the
generation of ONOO–.
In the present study, the observed increase in ROS generation and nitrotyrosine formation
preceded the cell injury, suggesting a causal relationship. Further evidence of a causal role for
ROS, particularly ONOO–, is demonstrated by increased caspase-3 activity following the
addition of synthetic ONOO– to LLC-PK1 cells. Moreover, the ONOO– scavenger ebselen
reduced caspase-3 activity and significantly reduced cell injury. Carbon dioxide, primarily due to
its abundance in living systems (mainly from bicarbonate), reacts readily with ONOO–, forming
nitrosoperoxycarbonate--a highly reactive intermediate that further decomposes to form nitrogen
dioxide (!NO2) and the carbonate anion (CO3!–), free radicals that carry out oxidation and
nitration reactions. The combined effect of these compounds is the nitration and nitrosation of
key regulatory proteins. ONOO– and other ROS can induce apoptosis in several systems (22, 40–
44) largely by increasing mitochondrial membrane permeability and thereby causing the release
of cytochrome c and sequential activation of caspases-9 and -3 (22). In addition, caspase-3 can
be activated by caspase-8 via the Fas/FADD pathway (45). We have shown here that the
expression of XIAP is decreased in high glucose-exposed LLC-PK1 cells. To our knowledge, this
is the first demonstration of XIAP regulation by high glucose concentrations. XIAP inhibits
caspases primarily by preventing cleavage of pro-caspases but may also directly inhibit activated
caspases (46). A mechanism by which high glucose concentrations cause cell death by downregulating endogenous inhibitors of caspases is an attractive one and needs to be explored
further.
In endothelial cells high glucose-induced oxidative stress sequentially activated c-Jun NH2terminal kinase and casapse-3, leading to apoptotic cell death (47). It is conceivable that one of
these mechanisms operates to cause proximal tubular cell death in hyperglycaemia.
Alternatively, activation of either caspase-8 or -9 could independently induce an apoptotic
response involving other effector caspases such as caspases-6 and -7 (45, 48, 49) as well as the
calcium-dependent cysteine protease, calpain (50, 51). More likely, a combination of apoptotic
pathways may contribute to PTEC apoptosis in high glucose. The lack of a significant increase in
caspase-8 activity in LLC-PK1 cells exposed to high glucose is confusing because the cellpermeable caspase-8 inhibitor prevented caspase-3 activation and apoptosis. The reason for this
discrepancy is not clear, but it is possible that the cell-permeable caspase-8 inhibitor affects
caspase-3 directly via nonspecific inhibition. The role of caspase-8 in high glucose-induced
proximal tubular cell death is thus unconfirmed and may be peripheral to the role of caspase-9
and -3. The role of caspase-8 and -9 in high glucose-induced PTEC apoptosis is currently under
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investigation in our laboratory. Significantly, treatment of proximal tubular cells with ebselen
reduced caspase activation and DNA damage, indicating that regardless of the identity of the
caspases involved, oxidative stress precedes activation of cell death pathways.
It is not possible from the experiments we describe here to definitively state that high glucoseinduced ONOO– formation occurs within the mitochondria or in the cytosol. The half-life of
ONOO–, either as anion or protonated acid (ONOOH) is sufficient to allow diffusion across the
mitochondrial membrane. Thus, detection of ONOO– in the mitochondrion does not in itself
prove mitochondrial ONOO– formation. Nevertheless, wherever ONOO– is formed, the effect on
mitochondrial cell death pathways is clear. ONOO– can oxidize several components in the inner
and outer mitochondrial membrane, leading to the opening of mitochondrial pores. The change
in mitochondrial membrane potential and subsequent release of cytochrome c leads to activation
of caspase-9 and initiation of an apoptotic cascade. Taken together, our data show that high
glucose induces the formation of ONOO– either in the cytosol or the mitochondrion and that this
powerful oxidant initiates apoptosis involving caspase-9 activation. The partial reversal of these
processes by ebselen suggests that this probably occurs as a result of a combination of oxidation
and nitration reactions that are commonly associated with the reactivity of ONOO– and its
derivatives.
Metalloporphyrins, which are often used as ONOO– scavengers, react with ONOO– directly in a
second-order rate reaction. The metal (either Mn3+ or Fe3+) catalyzes the isomerization of
ONOO– to nitrate. However, a product of this reaction is !NO2, which can enhance the nitration
reactions of ONOO–. Thus, any protective effect of the scavenger is masked first by its inability
to compete with CO2 and second generating !NO2. In contrast, the data presented here show that
the seleno-organic compound ebselen, which reacts with ONOO– in preference to CO2 (17),
reduced high glucose-induced DNA fragmentation in PTECs. Ebselen undergoes oxidation upon
reaction with ONOO– in a similar manner to glutathione and thus may mimic cellular glutathione
in its antioxidant effects. In a study by Powell et al. (52), in vascular smooth muscle cells,
glutathione levels were reduced and DNA damage increased in response to high glucose.
Similarly, the reduction of glucose to sorbitol consumes NADPH and could exacerbate oxidative
stress because NADPH is required for reduction of oxidized cellular glutathione (53). In our
study, a vitamin E analog (Trolox) reduced ONOO– generation in LLC-PK1 cells exposed to high
glucose. Trolox can undergo one- or two-electron oxidation by ONOO– in a reaction that
generates nitrite, nitrate, and the oxidation product quinone (54). Thus, two chemically distinct
ONOO– scavengers reduce overall oxidative stress and apoptosis in LLC-PK1 cells exposed to
high ambient glucose concentrations. Taken collectively, our data show that ONOO– mediates
high glucose-induced proximal tubular apoptosis by activating a caspase cascade.
Traditionally, hyperglycemic injury has been studied in chronic models. Glomerular basement
membrane thickening and the deposition of extracellular matrix, which result in the development
of interstitial fibrosis, are commonly acknowledged as pivotal steps in the pathogenesis of
diabetic nephropathy (55). Our data, obtained within 48 h of initial exposure, suggest that high
ambient glucose concentrations cause proximal tubular cell apoptosis as a result of oxidative
stress. Moreover, this effect of high glucose involves multiple proteases, including caspases-3
and -9. The use of a pan-caspase inhibitor completely prevented high glucose-induced PTEC
apoptosis, and this may be useful in the design of future therapies for the complications of
diabetic nephropathy.
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ACKNOWLEDGMENTS
We thank Dr. Neil Ashman for helpful discussions.
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Received May 30, 2002; accepted January 10, 2003.
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Fig. 1
Figure 1. High glucose induces oxidative stress in LLC-PK1 cells. A) Subconfluent monolayers of LLC-PK1 cells (in
six-well tissue culture plates) were exposed to 5 mM or 25 mM D-glucose for 24 h, and nitrite production was used as a
measure of nitric oxide synthase (NOS) activity. Nitrite concentrations were measured in supernatants by using 2,3diaminonapthalene (DAN) against a sodium nitrite standard curve. Results (in the range of 15–40 nmol/mg/h) are
expressed as fold of control ±SE (n=4, *P<0.05). B) The oxidation of dichlorofluorescein (DCFH) was used to detect
cellular peroxynitrite generation. LLC-PK1 cells were exposed to 5 mM or 25 mM D-glucose with or without 10 µM
ebselen or 50 µM 1400W for 24 h followed by 20 µM DCF-DA for 30 min. Cells were harvested, and DCF fluorescence
was detected using a microplate reader. Fluorescence, normalized for protein content, is presented as arbitrary units (n=4,
**P<0.02). C) The effect of a vitamin E analogue (Trolox, 100 µM) on high glucose-induced DCF fluorescence was
assessed in the same way as in B. Data are fold of control ±SD (n=6, *P<0.05).
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Fig. 2
Figure 2. Increased nitrotyrosine staining in LLC-PK1 cells exposed to high glucose. Western blots were carried out
on whole-cell lysates from LLC-PK1 cells after 24 or 48 h exposure to 5 mM or 25 mM D-glucose. Protein (~10 µg) was
subjected to electrophoresis on a 10% polyacrylamide gel. Membranes were probed with a monoclonal anti-nitrotyrosine
antibody followed by detection using chemiluminescence. Lane 1, day 0 control sample; lane 2, 5 mM D-glucose (24 h);
lane 3, 25 mM D-glucose (24 h). One of three experiments is shown.
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Fig. 3
Figure 3. High glucose causes apoptosis in LLC-PK1 cells. A) Subconfluent monolayers of LLC-PK1 cells in six-well
tissue culture plates were exposed to 5 mM or 25 mM D-glucose for 48 h. Whole-cell lysates were used for evaluation of
DNA fragmentation by ELISA as described in Materials and Methods. Samples (20 µl) were assayed in duplicate, and the
results were expressed as fold of control ±SE (n=4, *P<0.02). B) LLC-PK1 cells were grown in 25-cm3 tissue culture
flasks and exposed to 5 mM or 25 mM D-glucose for 24 h. Cells were harvested, and annexin V expression was analyzed
by flow cytometry. Duplicate measurements were carried out per sample for three independent experiments. Results are
expressed as the percentage of annexin V-positive cells ±SD (n=3, **P<0.05).
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Fig. 4
Figure 4. Caspase activity in LLC-PK1 cells exposed to high glucose. LLC-PK1 cells were exposed to 5 mM or 25
mM D-glucose for 24 or 48 h, and whole-cell lysates were analyzed for caspase activity. A) Caspase-3 activity was
measured using the fluorescent substrate Ac-DEVD-AMC. Results were calculated as nmol 7-amido-4-methylcoumarin
(AMC) released/mg/h and expressed as fold of control ±SE (n=9, *P<0.05). B) Caspase-8 activity (±SE) was measured
using Ac-IETD-AMC as the fluorescent substrate (48 h, n=5, P=not significant [NS]). C) Caspase-9 activity (±SE) was
measured using Ac-LEHD-AMC as the substrate (n=3, **P<0.01).
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Fig. 5
Figure 5. Expression of XIAP in high glucose-exposed LLC-PK1 cells. Western blot on whole-cell lysates from LLCPK1 cells after 24 or 48 h exposure to 5 mM or 25 mM D-glucose. Protein (~10 µg) was subjected to electrophoresis on a
10% polyacrylamide gel. Membranes were probed with a monoclonal anti-XIAP antibody (1:500) followed by detection
using chemiluminescence. Two experiments are shown for each time point at 5 mM and 25 mM D-glucose.
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Fig. 6
Figure 6. Effect of a peroxynitrite scavenger on high glucose-induced apoptosis in LLC-PK1 cells. Subconfluent
monolayers of LLC-PK1 cells in six-well tissue culture plates were exposed to 5 mM or 25 mM D-glucose for 48 h in the
presence or absence of the peroxynitrite scavenger ebselen (10 µM, present throughout). A) Whole-cell lysates were used
for evaluation of DNA fragmentation (ELISA). Results (±SE) are expressed as fold of control (48 h, n=6, *P<0.05).
B) The same lysates were assayed for caspase-3 activity, and results (±SE) are expressed as fold of control (48 h, n=3,
*P<0.05).
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Fig. 7
Figure 7. Caspase inhibitors prevent high glucose-induced apoptosis in LLC-PK1 cells. Subconfluent monolayers of
LLC-PK1 cells in six-well tissue culture plates were exposed to 5 mM or 25 mM D-glucose for 48 h with or without a
caspase-3 inhibitor (Ac-DEVD-CHO, 1.25 µM), a caspase-8 inhibitor (Ac-IETD-CHO, 1.25 µM), or a pan-caspase
inhibitor (Z-D-DCB, 100 µM). A) Whole-cell lysates were assayed for caspase-3 activity, and results (±SE) are expressed
as fold of control (n=6, *P<0.001). B) The same lysates were assayed for DNA fragmentation (ELISA), and results (±SE)
are expressed as fold of control (48 h, n=8, **P<0.05, ***P<0.01, #n=4).
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