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1
* Departments of Medicine,
Pathology, and
Microbiology and Immunology, State University of New York Health Science Center, College of Medicine, Syracuse, New York 13210, USA
1Correspondence: SUNY HSC, 750 East Adams Street, Syracuse, NY 13210, USA. E-mail:perla{at}upstate.edu
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Key Words: DHA • transaldolase • glucose 6-phosphate dehydrogenase • GSH • apoptosis
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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, 2)
. However, vitamin C also has
pro-oxidant properties and may cause apoptosis of human lymphocytes and
myelogenous leukemia cell lines (3
, 4)
. Under
physiological conditions, vitamin C has a predominantly antioxidant
role (5)
. Pro-oxidant and proapoptotic effects of vitamin
C may be related to hydroxylation (6)
and/or formation of
ascorbyl radicals (7)
. Apoptosis, a form of programmed
cell death (PCD), is indispensable for normal development and
homeostasis within multicellular organisms (8)
. Defects in
PCD may underlie the etiology of neurodegenerative diseases, cancer,
autoimmune diseases, and the acquired immune deficiency syndrome
(9
, 10)
.
Humans and other primates lack gulonolactone oxidase, a key enzyme for
ascorbic acid biosynthesis, and therefore it must be provided from
external sources (11)
. Even in ascorbic acid-synthesizing
species, the majority of cells need ascorbic acid from the outside
(12)
. Vitamin C is absorbed from the gastrointestinal
tract in the form of ascorbic acid and circulates in the blood as
ascorbate at pH of 7.4. Ascorbate and its oxidized form,
dehydroascorbate (DHA), are taken up by transporters and facilitated
diffusion in a cell type-specific manner (12)
. DHA is the
major transport form of ascorbate in blood cells, primarily via the
hexose transporter GLUT-1 (13)
. Within the cell, DHA is
regenerated into ascorbate at the expense of reduced glutathione (GSH)
(2)
. However, ascorbate cannot regenerate GSH from its
oxidized form, oxidized glutathione (GSSG). GSSG is reduced to GSH at
the expense of NADPH, which is produced by the pentose phosphate
pathway (PPP) (14)
. In fact, a fundamental function of PPP
is to maintain GSH in a reduced state, thereby protecting sulfhydryl
groups and cellular integrity from emerging oxygen radicals.
The PPP comprises two separate, oxidative and nonoxidative, phases
(14)
. Reactions in the oxidative phase are irreversible,
whereas all reactions of the nonoxidative phase are fully reversible.
The two phases are functionally connected. The nonoxidative phase
converts ribose 5-phosphate to glucose 6-phosphate for utilization by
the oxidative phase, and thus indirectly contributes to generation of
NADPH. Different enzymes are rate limiting in the two phases. The
oxidative phase primarily depends on glucose 6-phosphate dehydrogenase
(G6PD) (15)
, whereas transaldolase (TAL) is the
rate-limiting enzyme for the nonoxidative phase (16)
. TAL
(17)
and G6PD (18)
regulate NADPH production
and thereby influence GSH levels, mitochondrial transmembrane potential
(
m), and susceptibility to apoptosis
signals (17
, 19
, 20)
. Unless reduced back to ascorbate,
DHA is rapidly hydrolyzed into 2,3-diketo-L-gulonate and decarboxylated
to L-xylonate and L-lyxonate (21)
. In turn, these 5-carbon
sugars can enter the nonoxidative branch of the PPP (22
, 23)
.
This study provides evidence that DHA stimulates the activity of PPP
enzymes TAL, G6PD, and 6-phosphogluconate dehydrogenase (6PGD),
elevates intracellular GSH levels, and increases resistance of Jurkat
human T cells to
H2O2-induced cell death.
Although the antioxidant and pro-oxidant properties of vitamin C have
long been recognized, the existence of DHA at low concentrations in
plant and animal cells has been regarded only as evidence of ascorbate
oxidation (24
, 25)
. The present data show that DHA is not
merely a transport form of vitamin C. DHA stimulates the PPP and GSH
levels and inhibits
H2O2-induced cell death of
cells susceptible to the pro-oxidant properties of vitamin C.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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. Staining with phycoerythrin-conjugated
annexin V (annexin V-PE, R & D Systems) was used to monitor
phosphatidylserine (PS) externalization (FL-2) in parallel with
measurement of reactive oxygen intermediates (ROI) levels and
m, using dihydrorhodamine 123 (DHR),
5,6-carboxy-2',7'-dichlorofluorescein (DCF), or
DiOC6 fluorescence. PS externalization was
monitored with annexin V-FITC (FL-1) in parallel with measurement of
ROI levels using ethidium (FL-2), as described (20)
. Thus,
annexin V-PE or annexin V-FITC was matched with emission spectra of
potentiometric and oxidation-sensitive fluorescent probes. Specific
combinations are described in each figure legend.
Flow cytometric analysis of ROI production and m
The production of ROIs was estimated fluorometrically
using oxidation-sensitive fluorescent probes
5,6-carboxy-2',7'-dichlorofluorescein-diacetate (DCFH-DA), DHR, and
hydroethidine (HE, Molecular Probes, Eugene, Oreg.) as described
(17)
. After apoptosis assay, cells were washed three times
in 5 mM HEPES-buffered saline (HBS), pH 7.4, incubated in HBS with 0.1
µM DHR for 2 min, 1 µM DCFH-DA for 15 min, or 1 µM HE for 15 min,
and samples were analyzed using a Becton Dickinson FACStar Plus flow
cytometer equipped with an argon ion laser delivering 200 mW of power
at 488 nm. Fluorescence emission from DCF (green) or DHR (green) was
detected at a wavelength of 530 ± 30 nm. Fluorescence emission
from oxidized HE, ethidium (red), was detected at a wavelength of 605
nm. Dead cells and debris were excluded from the analysis by electronic
gating of forward and side scatter measurements. Whereas R123, the
fluorescent product of DHR oxidation, binds selectively to the inner
mitochondrial membrane, ethidium and DCF remain in the cytosol of
living cells. Mitochondrial transmembrane potential
(m) was estimated by staining with 40 nm
3,3'-dihexyloxacarbocyanine iodide (DiOC6,
Molecular Probes), a cationic lipophilic dye (26
27
28)
, for
15 min at 37°C in the dark before flow cytometry (excitation: 488 nm,
emission: 525 nm recorded in FL-1). Fluorescence of
DiOC6 is oxidation independent and correlates
with m (27)
.
DiOC6 staining was complete after a 15 min
incubation. DiOC6 fluorescence was diminished 3-
to 4-fold by 5 µM mClCCP and 10-fold or more by 50 µM mClCCP, as
described (20)
. m was also
quantitated using a potential-dependent, J aggregate-forming lipophilic
cation,
5,5',6,6'-tetrachloro-1,1',3,3'-tetraethylbenzimidazolocarbocyanine
iodide (JC-1) (29)
. JC-1 selectively incorporates
into mitochondria, where it forms monomers (fluorescence in green, 527
nm) or aggregates at high transmembrane potentials (fluorescence in
red, 590 nm) (29
, 30)
. Cells were incubated with 1 µM
JC-1 for 15 min at 37°C before flow cytometry. Cotreatment with a
protonophore, 5 µM carbonyl cyanide m-chlorophenylhydrazone (mClCCP,
Sigma) for 15 min at 37°C resulted in decreased DHR,
DiOC6, and JC-1 fluorescence and served as a
positive control for disruption of mitochondrial transmembrane
potential (20)
.
Pentose phosphate pathway enzyme activities
TAL activity was tested in the presence of 3.2 mM D-fructose
6-phosphate, 0.2 mM erythrose 4-phosphate, 0.1 mM NADH, 10 µg
-glycerophosphate dehydrogenase/triosephosphate isomerase at a 1:6
ratio at room temperature by continuous absorbance reading at 340 nm
for 6 min (31)
. The enzyme assays were conducted in the
activity range of 0.001–0.01 U/ml. G6PD was measured in the presence
of 120 mM Tris pH 7.7, 10 mM MgCl2, 2 mM glucose
6-phosphate, 0.9 mM NADP, and 0.1 U/ml 6PGD (32)
. 6PGD
activity was determined in 120 mM Tris pH 7.7, 10 mM
MgCl2, 0.9 mM NADP, 2 mM 6-phosphogluconate
(32)
.
Glutathione levels
Total glutathione content was determined by the enzymatic
recycling procedure essentially, as described by Tietze
(33)
. 106 cells were resuspended in
50 µl of 4.5% 5-sulfosalicylic acid. The acid-precipitated protein
was pelleted by centrifugation at 4°C for 10 min at 15,000
g. The total protein content of each sample was determined
using the Lowry assay (34)
. GSH content of the aliquot
assayed was determined in comparison to reference curves generated with
known amounts of GSH (17)
. GSH and GSSG were measured by
reverse phase ion exchange high-performance liquid chromatography
(HPLC) using UV detection at 365 nm (35)
. Briefly,
106 cells were deproteinized in the presence of
10% perchloric acid and 1 mM bathophenanthroline disulfonic acid.
After repeated freezing and thawing, samples were centrifuged at 15,000
g for 3 min. Fifty microliters of 100 mM mono-iodo-acetic
acid in 0.2 mM m-cresol purple was added to 500 µl supernatant.
Samples were neutralized by addition of 480 µl of 2 M KOH and 2.4 M
KHCO3 and incubated in the dark at room temperature for 10 min. Then 1
ml of 1% fluoro-dinitro-benzene was added and the samples were
incubated in the dark at 4°C overnight. After centrifugation and
filtering, 100 µl of supernatants were injected into the HPLC
equipped with a photodiode array detector (Waters Alliance System,
Milford, Mass.) and a Waters Spherisorb NH2
column (4.6x250 mm; 10 µm) .
Measurement of ascorbate
4 x 106 Jurkat cells were
treated with 400–800 µM DHA for 15 min to 48 h. After
incubation, cells were washed with phosphate-buffered saline and the
pellets were deproteinized in 5% metaphosphoric acid. The lysate were
centrifuged for 10 min at 14,000 g; 25 µl of each
supernatant was injected on a C18 column (Waters
Nova-Pak; 3.9x150 mm, 4 µm). Ascorbate was detected at 232 nm
wavelength using HPLC (36)
.
Western blot analysis
Forty micrograms of total cell lysate in 10 µl per well
was separated by sodium dodecyl sulfate-polyacrylamide gel
electrophoresis and electroblotted to nitrocellulose, as described
(17)
. Nitrocellulose strips were incubated in 100 mM Tris
pH 7.5, 0.9% NaCl, 0.1% Tween 20, and 5% skim milk with the primary
antibodies, anti-G6PD rabbit antibody (37)
(kindly
provided by Rolf Kletzien, Upjohn, Kalamazoo, Mich.) and anti-actin
monoclonal antibody C4 (Boehringer, Indianapolis, Ind.), at room
temperature overnight. After washing, the blots were incubated with
biotinylated secondary antibodies and then with horseradish
peroxidase-conjugated avidin (Jackson Laboratories, West Grove, Pa.).
In between incubations, the strips were washed in 0.1% Tween-20, 100
mM Tris pH 7.5, and 0.9% NaCl. The blots were developed with a
substrate comprised of 1 mg/ml 4-chloronaphthol and 0.003% hydrogen
peroxide. G6PD and actin protein levels of control and DHA-treated
cells were quantified by densitometry (Model GS-700, Bio-Rad, Hercules,
Calif.) of Western blots.
Statistics
Alterations in cell survival, PPP enzyme activities, and GSH
levels were analyzed by Student’s t test. Changes were
considered significant at P<0.05.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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, 4
, 7)
. Accordingly, incubation of Jurkat human T cells with
200–800 µM ascorbic acid dose-dependently diminished cell survival
(Fig. 1A
). Since lymphoid cells preferentially import DHA, the
oxidized form of ascorbic acid, we examined the effect of DHA on cell
survival and intracellular accumulation of ascorbate. Treatment with
DHA, up to 1 mM, did not influence cell viability (Fig. 1B
).
Fifteen minutes after addition of DHA (400 µM), intracellular
ascorbate levels increased from 2.72 ± 0.05 mM to 4.54 ±
0.01 mM (mean±SD of four experiments;
P<0.001). Ascorbate levels were also elevated after 60 min
DHA treatment (4.60±0.01 mM) and returned to baseline 48 h after
addition of DHA. Subsequently, we examined the effect of DHA on
H2O2-induced apoptosis.
Jurkat cells were preincubated with DHA, washed, and apoptosis was
induced with 100 µM H2O2,
as described before (17)
. Surprisingly, short- (20 min to
2 h) and long-term (24 h to 48 h) DHA pretreatment had
opposing effects on cell survival. Preincubation with DHA for 20 min to
2 h enhanced
H2O2-induced apoptosis
(Figs. 1B, C
). In contrast, pretreatment with DHA for
36 h strongly inhibited cell death (Fig. 1B, C
). When
Jurkat cells were pretreated with DHA for 36 h, washed, and
repeatedly preincubated with DHA for 1 h,
H2O2-induced apoptosis was
reduced (Fig. 1C
). These studies suggest that the effect of
long-term pretreatment was dominant over short-term effects of DHA.
Similar findings were obtained in H9 human T cells (data not shown).
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The mechanism of action of DHA on
H2O2-induced apoptosis was
likely to involve GSH metabolism (2)
. Short-term (less
than 8 h) incubation with DHA had no effect on GSH levels, whereas
pretreatment with DHA for 24 h to 48 h increased
intracellular GSH levels up to threefold (Fig. 2A
). Maximum GSH elevation, from a baseline of 7.65 ±
1.2 ng/ml to 25.36 ± 3.05 ng/ml (P<0.001), was
observed after 48 h stimulation with 800 µM DHA (Fig. 2A
). Five-carbon products of DHA catabolism can enter the
PPP (23)
. Previous studies revealed that activity of PPP
enzymes TAL (17)
and G6PD influence GSH levels and
sensitivity to apoptosis dependent on formation of ROIs
(18)
. Therefore, the effect of DHA on these enzyme
activities was also evaluated. G6PD, 6PGD, and TAL activities were
increased after incubation with DHA in a time- (Fig. 2A
) and
concentration-dependent manner (Fig. 2B
). G6PD activity
started to rise 24 h after addition of DHA. Incubation with 800
µM DHA doubled G6PD activity after 48 h (Fig. 2A
).
This was accompanied by a 5.9-fold increase in G6PD protein levels with
respect to actin levels based on densitometry of Western blots (Fig. 2C
). 6PGD and TAL activities were stimulated to a lesser
extent (Fig. 2A, B
). Since DHA can be reduced back to
ascorbate at the expense of GSH, we studied the time course of
DHA-induced changes in GSH and GSSG levels. Preincubation with DHA for
less than 24 h did not influence intracellular GSH or GSSG levels
or PPP enzyme activities (Fig. 2A, D
). HPLC measurement of
GSH and GSSG levels after 48 h of preincubation with DHA showed a
selective increase of GSH levels (Fig. 2D
). This indicated
that augmented PPP activities after long-term DHA pretreatment were not
triggered by an initial GSH oxidation, i.e., depletion of GSH or
increased GSSG content.
|
Disruption of the mitochondrial membrane potential
(m) has been proposed as the point of no
return in apoptotic signaling (28
, 38
, 39)
. Mitochondrial
membrane permeability is subject to regulation by an oxidation
reduction equilibrium of ROIs, pyridine nucleotides (NADH/NAD +
NADPH/NADP), and GSH levels (40)
. Since DHA stimulated PPP
enzyme activities and augmented GSH levels, its effect on
m was also investigated.
m and mitochondrial ROI production were
monitored with respect to externalization of PS, an early event in
PCD (41
, 42)
. As expected, incubation with
H2O2 resulted in PS
externalization (Fig. 3A, B)
. m was diminished in annexin
V-positive cells (Fig. 3A
). In correlation with previous
results, DiOC6 fluorescence was increased in
annexin V-negative cells (Fig. 3A
), suggesting that an
elevation of m preceded PS externalization
in Jurkat cells (19
, 20)
. Control cells stained with JC-1
showed green fluorescence whereas
H2O2-treated cells gained
red JC-1 fluorescence (Fig. 3C
), consistent with a higher
m (20
, 29
, 30)
.
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DHA pretreatment for 36 h inhibited key checkpoints of
H2O2-induced apoptosis,
attenuated the augmentation of m in annexin
V-negative cells (Fig. 3A
), and reduced PS externalization
(Figs. 3a
and 3b)
. By contrast, 1 h DHA pretreatment stimulated
H2O2-induced elevation of
m in annexin V-negative cells and
accelerated PS externalization (Figs. 3A, C
). One hour of
DHA pretreatment increased m even in the
absence of H2O2 (Fig. 4A, B
). This effect of DHA on m
was not associated with increased production of ROIs (Fig. 4C, D
). The effect of DHA on ROI levels was assessed by flow cytometry
using oxidation-sensitive fluorescent probes DHR and HE, as described
earlier (17
, 20
, 43
, 44)
. DHR is nonfluorescent,
uncharged, and readily taken up by cells whereas R123, the product of
DHR oxidation, is fluorescent, positively charged, and binds
selectively to the inner mitochondrial membrane of living cells
(44)
. Fluorescence of this dye is an indicator of
mitochondrial ROI production and maintenance of the membrane integrity
and transmembrane potential. HE is oxidized into ethidium by ROIs and
remains in the cytosol (45)
. Thus, R123 fluorescence
correlates with mitochondrial ROI levels whereas ethidium fluorescence
reflects cytosolic ROI levels. Short-term (1 h) or long-term (36 h)
treatment with DHA alone has not increased R123 (Fig. 3D
)
and ethidium fluorescence (Fig. 4D
), thus excluding the
possibility of an DHA-induced oxidative stress.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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. Vitamin C
functions as an antioxidant by reacting directly with ROIs or
regenerating vitamin E from -tocopheroxyl radical (1
, 46)
. Thus, it protects cell membranes from external oxidants and
has a GSH-sparing effect. However, vitamin C cannot be efficiently
transported into blood cells. In fact, ascorbate concentrations above
the normal serum level of 50 µM induce oxidative stress and apoptosis
of lymphoid and myeloid cells (3
, 4
, 7)
. The existence of
DHA has long been considered evidence of ascorbate oxidation (24
, 25)
. Blood cells and related tumor cell lines preferentially
import DHA over ascorbate (12)
. The present study provides
evidence that DHA can increase GSH levels through stimulation of the
PPP. A maximum of 3.3-fold GSH elevation was observed after 48 h
stimulation with 800 µM DHA. DHA may be a more potent stimulator of
GSH levels than N-acetyl cysteine. GSH levels could only be increased
up to twofold by treatment with N-acetyl cysteine, at an optimal
concentration of 3 mM, in Jurkat cells (17)
. Thus,
intracellular presence of DHA may not only represent an accumulation of
the oxidized form of vitamin C, but may have a role in influencing GSH
levels through the PPP. Products of DHA metabolism,
2,3-diketo-L-gulonate and its decarboxylation products, L-xylonate and
L-lyxonate, can enter the nonoxidative branch of PPP
(21
22
23)
. Activity of the PPP is vitally important by
providing NADPH for biosynthetic reactions and maintenance GSH in a
reduced state and, thus, regulating the redox homeostasis of the cell.
Accordingly, treatment of Jurkat cells with DHA increased G6PD, 6PGD,
and TAL activities. These changes were accompanied by a severalfold
increase in GSH level and protection from
H2O2-induced cell death.
DHA-induced elevations of enzyme activities were accompanied by a
2.6 ± 0.2-fold increase in G6PD protein levels with respect to
actin levels, suggesting that DHA may stimulate gene expression. Carbon
flux via the PPP has been implicated in regulating expression of other
glucose-metabolizing enzymes, pyruvate kinase (47)
,
glucose 6-phosphatase, and phosphoenolpyruvate carboxykinase
(48)
. While xylulose 5-phosphate (X5P) has been proposed
as a key intermediate, its effect on the PPP and potential involvement
of X5P metabolites remain to be determined (47
, 48)
.
GSH and GSSG levels were not affected by pretreatment of Jurkat cells
with DHA for less than 24 h. Along the same line, DHA did not
influence intracellular ROI levels as estimated by DHR and HE
fluorescence. DHA treatment, up to 1 mM, did not affect cell viability.
Our data correlate with observations by others showing that DHA has no
pro-oxidant effects and it may directly neutralize ROIs by undergoing a
peroxidative decarboxylation reaction (49)
. Thus, the
possibility that increased PPP enzyme activities were triggered by a
DHA-induced oxidative stress could clearly be excluded.
Stimulation by DHA of activities of the NADPH-generating enzymes G6PD
and 6PGD may be responsible for elevation of GSH levels. Genetically
enforced augmentation of G6PD activities can increase GSH levels and
resistance to apoptosis signals (17
18
19)
. In turn,
increased GSH can directly stimulate expression of 
-glutamylcysteine
synthetase, the rate-limiting enzyme of de novo GSSG
synthesis (50)
, and further support GSH production.
Pretreatment with DHA from 20 min to 2 h increased
m and enhanced toxicity of
H2O2. Short-term
(< 2 h) DHA pretreatment increased m,
even in the absence of
H2O2. This effect of DHA on
m was not associated with increased
production of ROIs. Elevation of m, which
precedes PS externalization (20)
, was clearly enhanced by
short-term DHA treatment. This transient effect of DHA may be related
to its rapid metabolism (25)
. Once transported into the
cell, DHA is reduced back to ascorbate or rapidly hydrolyzed
(25)
. Ascorbate passively diffuses through the inner
mitochondrial membrane (51)
. The resultant increase in
ascorbate or short-chain sugar content (52
, 53)
may be
responsible for the transient elevation of
m and susceptibility to
H2O2-induced cell death.
Short-chain sugars, glyceraldehyde, dihydroxyacetone, erythrose
(53)
, and ribose 5-phosphate (52)
can enhance
oxidative killing (53)
and cause apoptosis of lymphocytes
(52)
. Thus, DHA metabolism in the nonoxidative branch of
the PPP may increase the concentration of short-chain sugars and
temporarily enhance toxicity of
H2O2. A combination of
long- and short-term treatment regimens revealed an overall protective
effect of DHA against cell death.
A direct redox relationship between ascorbate and glutathione has long
been established (2)
. This latter relationship is based on
the glutathione-sparing effect of ascorbate in the event of oxidative
stress (2)
. The present work is reveals an indirect
relationship between DHA and glutathione via the PPP. DHA can elevate
GSH levels through stimulation of the PPP pathway. This antioxidant
effect of DHA is clearly distinct from the previously recognized
glutathione-sparing effect of ascorbate. The effect of DHA on the PPP
and GSH synthesis is also significant with respect to the
pro-oxidant/proapoptotic effects of ascorbate on myeloid and lymphoid
cells (3
, 4)
. By stimulation of GSH levels, DHA may have a
important role in antioxidant defenses of cells preferentially
importing DHA over ascorbate. The data identify DHA and its sugar
metabolites as potential targets for development of novel therapeutics
aimed at enhancing antioxidant defenses.
|
ACKNOWLEDGMENTS |
|---|
Received for publication August 20, 1999.
Revision received November 23, 1999.
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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F. Petrat, S. Paluch, E. Dogruoz, P. Dorfler, M. Kirsch, H.-G. Korth, R. Sustmann, and H. de Groot Reduction of Fe(III) Ions Complexed to Physiological Ligands by Lipoyl Dehydrogenase and Other Flavoenzymes in Vitro: IMPLICATIONS FOR AN ENZYMATIC REDUCTION OF Fe(III) IONS OF THE LABILE IRON POOL J. Biol. Chem., November 21, 2003; 278(47): 46403 - 46413. [Abstract] [Full Text] [PDF]
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