Hospital for Children and Adolescents, Research Program for Developmental and Reproductive Biology, University of Helsinki, Biomedicum Helsinki, 00290 Helsinki, Finland
Submitted 2 December 2002 ; accepted in final form 28 February 2003
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ABSTRACT |
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xanthine oxidase; hypoxia; hyperoxia; ischemia reperfusion
Xanthine oxidoreductase (XOR; EC 1.1.3.22 [EC] ) catalyzes the oxidation of hypoxanthine to xanthine and on to uric acid, which are the final reactions of purine catabolism in humans. Under physiological conditions, the enzyme functions as a dehydrogenase (XDH) and uses NAD+ as the electron acceptor, but it can be converted into an oxidase (XO), with molecular oxygen as the electron acceptor, under a variety of conditions such as tissue ischemia (12, 36). Because the substrates of XOR accumulate in hypoxia (34), and because the oxidase form has been proposed as a major source of reactive oxygen metabolites in reperfused/reoxygenated tissues (8, 24), the regulation of XOR by oxygen is of interest.
Hypoxia has been reported to increase XOR activity in cultured rat and bovine endothelial cells (11, 28, 38, 39). The mechanism of the increase, however, is uncertain, because some studies have reported increases in XOR mRNA transcript levels (11, 39), whereas another study failed to show any alterations in XOR gene expression by hypoxia (28). Furthermore, phosphorylation of XOR in hypoxia has been suggested to account for increased XOR activity (21). On the other hand, oxygen metabolites generated in the course of catalysis (37) or added in vitro (2) may inactivate the enzyme. In the interpretation of any studies on the expression and regulation of XOR, differences between species constitute a major problem (27).
XOR is a homodimer with a subunit molecular weight of 150 kDa. Each
subunit has a molybdopterin cofactor, two iron-sulfur clusters of the [2Fe-2S]
ferredoxin type, and a bound flavin adenine dinucleotide (FAD), all of which
are potential targets for inactivation by oxygen or its metabolites. In this
study, we show that XOR activity is increased in human bronchial epithelial
cells cultured in a hypoxic atmosphere, whereas XOR protein, mRNA, and
promoter activity remain unchanged. However, the induction of XOR in hypoxic
cells is only apparent and represents reversal of oxygen-induced inactivation
of the molybdenum center of the XOR enzyme, which occurs in the normal
atmosphere of standard cell culture conditions.
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MATERIALS AND METHODS |
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Experimental design. When the cells grown in normoxia (21% O2) reached confluence, fresh medium was provided, and the cells were exposed to either hypoxia, normoxia, or hyperoxia for specified periods of time (448 h). To achieve hypoxia or hyperoxia, a preanalyzed gas mixture (5% CO2, specified O2%, balance N2; AGA, Finland) was infused into airtight humidified chambers (Billups-Rothenburg, Del Mar, CA), and gas flow was adjusted so that the oxygen concentration remained stable during the incubation. The chambers were maintained in an incubator at 37°C for the duration of the exposure. After exposure, the cells were rinsed twice with phosphate-buffered saline and mechanically harvested with a plastic policeman. The harvested cells were centrifuged, and the cell pellet was resuspended in 500 µl of 50 mM potassium phosphate buffer, pH 7.8, containing 0.5 mM dithiothreitol and 1 mM EDTA and sonicated on ice. For cobalt treatment (224 h), CoCl2 was added to fresh medium and the cells were maintained in normoxia.
Assays of enzyme activities. XOR activity was measured by using [14C]xanthine (0.1 mM, specific activity 58 mCi/mmol) (NEN; Life Science Products, Boston, MA) as substrate and separating the product uric acid by HPLC as described (31). For total XOR activity (XDH + XO), NAD+ (400 µM) was present, whereas for XO assay, NAD+ was omitted. In the case of purified XOR, XO activity was measured spectrophotometrically by detecting absorbance change over 3 min at 295 nm corresponding to uric acid production (40). Total protein was determined with the Bio-Rad DC protein assay (Bio-Rad, Hercules, CA).
The electron transfer activity from xanthine (0.05 mM) to the artificial substrate 2,6-dichlorophenolindophenol (DCPIP) was determined spectrophotometrically by monitoring the absorbance of DCPIP (0.05 mM) at 600 nm. NADH oxidation was measured spectrophotometrically by monitoring the absorbance of NADH (0.1 mM) at 340 nm. All activity measurements were performed in 50 mM potassium phosphate buffer, pH 7.8, containing 0.5 mM dithiothreitol and 1 mM EDTA.
Inactivation of XOR. The effect of oxygen metabolites, generated by the enzyme itself, on XOR activity was studied by using purified bovine XOR (14.4 U/ml, Biozyme Laboratories, South Wales, UK), diluted 1:10,000 with 50 mM potassium phosphate buffer. The enzyme was incubated for 30 min with hydrogen peroxide (30 or 90 mM) (Merck, Darmstadt, Germany). XOR activity was measured over 3 min at the start and at the end of the incubation. Reduction of DCPIP was determined from the same samples.
Quantification of XOR protein. XOR protein concentrations were determined by ELISA as described (33).
Western blot analysis. Cells were harvested and sonicated in 50 mM potassium phosphate buffer, pH 7.8, containing 0.5 mM dithiothreitol and 1 mM EDTA, followed by denaturation by heating in 2-mercaptoethanol-containing loading buffer (5 min at 95°C). The samples (20 µg of protein) were separated by SDS-PAGE for 60 min at 140 V constant voltage and transferred onto Immobilon-P membranes (Millipore, Bedford, MA) by electrophoretic transfer at 30 V constant voltage overnight. The membrane was blocked with 5% bovine skimmed milk for 60 min and probed with polyclonal rabbit anti-hXOR antibody (diluted 1:300) (33) for 2 h, followed by horseradish peroxidase-conjugated secondary antibody (diluted 1:5,000) (Jackson ImmunoResearch, West Grove, PA). The antibodies were visualized using the enhanced chemiluminescence detection kit (Amersham Pharmacia Biotech, Amersham, UK). High-range protein size standards (Bio-Rad) were visualized by staining with 0.25% Coomassie blue.
Ribonuclease protection assay. To prepare total cellular RNA,
cells were rinsed twice with phosphate-buffered saline, detached with a
plastic policeman into 4 M thiocyanate buffer, immediately frozen at
80°C, and then extracted with the acid phenol-chloroform method
(5). Ribonuclease protection
assay (RPA) was carried out according to the manufacturer's protocol (RPA II
kit; Ambion, Austin, TX), using 20 µg of total RNA and hybridizing
overnight at 42°C with a 32P-labeled 384 bp cRNA probe
(corresponding to the nucleotides 405789 of the human XOR cDNA)
(32), specific activity 4
x 108 cpm/µg, 60.000 cpm/sample. To control for the amount
of RNA, a parallel assay was performed using a 171-bp -actin cRNA probe
(pTRI-
-actin; Ambion). After RNase A + T1 digestion, the
protected fragments were separated on 5% polyacrylamide/8 M urea gels and
exposed to Kodak BioMax MR autoradiography film (Eastman Kodak Co, Rochester,
NY). The X-ray films were scanned (Scan Jet 6300C), and the XOR/
-actin
ratios were analyzed with Scion Image beta 4.0.2 analysis software (Scion,
Frederick, MD).
Promoter constructs and reporter gene analysis. Human XOR gene promoter fragments (XOR1, XOR2, XOR4, and XOR5) were isolated as previously described (23). The nucleotide sequence data for the promoter of the human XOR gene have been deposited in the GenBank database under GenBank accession no. AF203979 [GenBank] . A hypoxia-responsive reporter gene construct (HRE-luc) carrying three tandem copies of the erythropoietin hypoxia-responsive element coupled to luciferase was kindly provided by Dr. Pekka Kallio (20).
The XOR promoter constructs and HRE-luc were transiently transfected into
293T cells using the FuGene6 transfection reagent (Roche Molecular
Biochemicals, Indianapolis, IN) according to the manufacturer's instructions.
The cells were seeded onto 12- or 6-well plates (2 x 105 or 5
x 105 cells per well, respectively) and transfected 24 h
later with 0.33 or 1 µg of XOR promoter constructs or HRE-luc,
respectively. In all experiments, 0.17 or 0.5 µg pCMV (Clontech, Palo
Alto, CA), producing
-galactosidase, was cotransfected to monitor for
transfection efficiency and empty pGL3-basic vector was used as a control.
Medium was changed 16 h after transfection, and the cells were further
incubated for 24 h in either 21 or 0.5% oxygen. Luciferase and
-galactosidase activity were determined as described
(23).
Assessment of cell injury. Cell suspension was incubated with 0.4% trypan blue for 5 min at room temperature. The cells were counted using a Bürker cell counting chamber, and trypan blue-negative cells were considered viable.
Statistics. All values are shown as means ± SD. Experiments were obtained in triplicate for each experimental and control condition. The differences between two groups were compared using unpaired Student's t-test. All tests are two-tailed, and statistical significance is assumed at P < 0.05.
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RESULTS |
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Cell viability. After incubation, under any of the oxygen concentrations used, over 95% of the cells excluded trypan blue, indicating that cell viability was not compromised.
Cobalt. Because the hypoxic induction of erythropoietin and other
HIF-1-regulated genes can be mimicked by cobalt
(4,
29), BEAS-2B cells were
incubated with CoCl2 (75 µM) for 24 h. This treatment did not
alter XOR activity, suggesting that a signaling pathway involving the
HIF-1
transcription factor may not be relevant here.
Substrate induction. To rule out enzyme induction by substrate accumulating in hypoxic cells (34), exogenous hypoxanthine (100 µM) was added to the medium of BEAS-2B cells grown in normoxia (21% O2). No increase in XOR activity was found (data not shown).
Effect of hypoxia on XOR promoter-driven transcription. The human
XOR promoter carries putative HIF-1 and activator protein (AP-1) sites
(13,
44), known to mediate effects
of changes in oxygen tension. To study the transcriptional activation of XOR,
we prepared a set of constructs containing variable lengths of the human XOR
promoter coupled to a luciferase reporter gene
(Fig. 2)
(23). The effects of hypoxia
on XOR promoter activity were studied in 293T cells rather than BEAS-2B cells,
because the latter could not be effectively transfected with the constructs.
No XOR activity is measurable in normoxic 293T cells, but the maximally active
XOR5 construct showed a ninefold increase in luciferase production compared
with the promoterless pGL3-basic
(23). However, exposure to
0.5% O2 for 24 h did not change XOR promoter activity in cells
transfected with any of the constructs, compared with cells grown under
normoxic conditions, whereas a control construct carrying three erythropoietin
hypoxia-responsive elements (HRE-luc) showed a 15-fold induction of luciferase
production in hypoxia (Fig.
2).
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Effect of hypoxia on XOR mRNA. Although transcriptional control
could not be shown, hypoxia could affect the stability of XOR mRNA. To
investigate this possibility, XOR mRNA levels were assessed using RPA. No
increase in XOR mRNA was observed in BEAS-2B cells exposed to 3% O2
for 4, 11, and 24 h, compared with cells before hypoxic exposure as determined
by XOR/-actin ratios (Fig.
3A).
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Effect of hypoxia on XOR protein. To further evaluate the basis for the increased XOR activity, we examined the effect of 3% O2 on the amount of XOR protein in BEAS-2B cells. As determined by ELISA, the amount of XOR protein relative to total cellular protein remained constant after exposure to 3% O2 for up to 48 h (Fig. 3B). Western blots showed no additional bands compared with cells grown in 21% O2, and the major 150-kDa XOR band remained unaltered (Fig. 3C).
Effect of reoxygenation on XOR activity. To evaluate whether the elevated XOR activity in hypoxic cells was sustained in normoxia, confluent cultures of BEAS-2B cells were exposed to 3% O2 for 24 h and then continued in 21% O2 for another 24 h. Reoxygenation of previously hypoxic cells resulted in a decline of total XOR activity (XDH + XO) to control (21% O2) levels, whereas continued culture at 3% O2 caused no further elevation (Fig. 4). No alteration in the ratio of XO to total XOR enzyme activity was seen (data not shown).
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Effect of hyperoxia on XOR activity and protein. Because all our data were compatible with posttranslational responses to varying oxygen levels, the effects of true hyperoxia were investigated. Incubation of BEAS-2B cells in 95% O2 abolished XOR activity after 24 h of incubation (Fig. 5A), whereas XOR protein concentrations, as determined by ELISA, remained unchanged compared with cells grown in 21% O2 (Fig. 5B). Also, in a Western blot analysis of cells grown in 95% O2 for 24 h, the intensity of the 150-kDa band was constant and there were no additional bands compared with cells grown in 21% O2 (Fig. 3C), indicating that inactivation of the enzyme was not due to proteolysis.
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Mechanism of inactivation of XOR by oxygen. Reactive oxygen metabolites, generated either externally or by the catalytic reaction itself, have been shown to inactivate XOR (2, 37), and they are probably involved in oxygen-induced enzyme inactivation. To assess the mechanism of this inactivation, the function of the main redox centers of XOR was evaluated. NADH oxidation, which is solely dependent on the FAD center, was 18 nmol · min1 · mg1 protein in sonicates of cells grown at 21% oxygen, and this rate was not significantly altered by a 24-h exposure to 3% (25 nmol · min1 · mg protein1) or to 95% (21 nmol · min1 · mg protein1) oxygen, whereas xanthine to urate activity was totally abolished in 95% oxygen. Thus the FAD center appears uninfluenced by oxygen. To study the effect of oxygen metabolites on the molybdenum center of XOR, purified bovine XOR was incubated with H2O2 for 30 min. Addition of increasing concentrations of H2O2 progressively decreased XO activity as determined by the oxidation of xanthine to urate (Fig. 6). In parallel, XO lost its ability to transfer electrons from xanthine to DCPIP (Fig. 6), suggesting that H2O2 reacted with the molybdenum center, because DCPIP directly accepts an electron from reduced molybdenum (16).
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DISCUSSION |
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It was recently suggested that in rat endothelial cells, protein kinase p38 and casein kinase can phosphorylate XOR and thereby increase its activity after 4 h in hypoxia (21). However, we could show no change in XOR activity in human epithelial cells grown for 4 h in 3% O2 compared with cells grown in 21% O2, whereas by 24 h there was a marked increase. Because this was not accompanied with any change in mRNA or protein, activation of preexisting enzyme appears the most likely explanation. This conclusion is supported by the lack of activation of the XOR promoter reporter gene constructs in hypoxia, as well as by the reversal of the "induction" upon return of the cultures to 21% oxygen and total disappearance of activity at 95% O2, again without any change in XOR protein concentrations.
Cultured human bronchial epithelial cells thus exhibit a reversible inactivation-reactivation cycle of XOR as a function of ambient oxygen concentration. The inactivation of purified bovine XOR during xanthine oxidation and its prevention by oxygen metabolite scavengers suggest that the enzyme protein is susceptible to self-generated oxygen metabolites. Exogenously generated oxygen metabolites such as superoxide anion, hydrogen peroxide, or hydroxyl radical decreased purified XOR activity, and the inactivation was diminished by simultaneous addition of oxygen metabolite scavengers superoxide dismutase, catalase, and dimethylsulfoxide (37).
We have recently shown that increased intracellular iron increases XOR
activity at the transcriptional level
(22) and that hydroxyl radical
scavengers do not change basal or iron-induced XOR activity. If increased
intracellular iron generates hydroxyl radicals, an apparent discrepancy
between iron induction and oxygen inactivation exists. However, the activity
of XOR overexpressed in cultured cells decreased 30% without changes in
XOR protein levels, when the cells were incubated for 24 h with 1 mM ferric
ammonium citrate (unpublished data). Thus it is possible that oxidative stress
caused by exogenously added iron could decrease XOR activity, as we propose
for the oxygen effect in this study, but this inactivation seems to be
overridden by transcriptional induction of endogenous XOR by iron.
To account for the inactivation, any of the three types of redox centers involved in intramolecular electron transfer could potentially be damaged by oxygen metabolites. Because the rate of NADH oxidation was similar in cells grown in hyperoxia compared with those grown in normoxia or hypoxia, the binding site of NAD/NADH, i.e., the FAD cofactor, appears not to be affected. Iron-sulfur centers are basically sensitive to oxygen (1), and their reversible alteration has been implicated in the inactivation of quinolinate synthase and 3-hydroxyanthranilate oxidase (6). Although we cannot definitely rule out this mechanism, a more likely site of oxygen-induced damage is the molybdenum cofactor. Molybdopterin forms part of the binding site of the purine substrate, and the essential sulfur atom attached to the molybdenum atom can be replaced by oxygen as a result of treatment of XOR with cyanide (25) or nitric oxide (15). The resulting desulfo-XOR can be reactivated in vitro (25), and molybdenum cofactor sulfurase, the enzyme responsible for reinsertion of the sulfur atom at the active site in vivo, has recently been cloned and characterized in human tissues (14). Modification of the cyanolyzable sulfur at the molybdenum center has been shown to accompany the inactivation of chicken liver XOR by H2O2 (2). This is in line with our findings that a similar treatment of bovine XOR results in a parallel decrease of xanthine oxidation and DCPIP reduction, pointing to involvement of the molybdenum site (16). To account for the response of cultured cells to variations in the oxygen environment, reversible alterations at the molybdenum center thus appear the most likely mechanism.
In a human patient with molybdenum cofactor sulfurase mutation, the phenotype was that of classic xanthinuria, i.e., XOR deficiency (14). It is thus possible that a reversible inactivation-reactivation cycle functions in normal human tissues, with the sulfurase rescuing the enzyme after autoinactivation during catalysis.
We found that the increase in XOR activity in hypoxia was not associated with XDH-to-XO conversion. This contrasts with the findings in freshly isolated rat Kupffer cells, in which total XOR activity was unchanged but XDH was converted into XO during anoxic incubation (42). However, most of the studies using cultured cells have shown no conversion of XDH to XO in hypoxia, in agreement with our results (11, 28, 39).
For convenience, cells are usually cultured in "normal" (21%
O2) atmosphere containing 5% CO2, which means that in
monolayer culture the cells will equilibrate to the approximate oxygen tension
of 140 mmHg (18.7 kPa). This is far higher than in most tissues in vivo,
because the estimated physiological oxygen tensions are 3035 mmHg for
the liver (43), 2030
mmHg for the renal cortex (7),
20 mmHg for the cerebral cortex
(41), and 1217 mmHg for
epicardium and myocardium
(30). Sensing mechanisms for
detection of and response to reduced oxygen availability
(35) should only be triggered
below these threshold levels of oxygen tension. In HeLa cells, the levels of
HIF-1 protein and HIF-1 DNA-binding activity started to increase
exponentially when the oxygen concentration was decreased below 5%, with
half-maximal responses between 1.5 and 2% and maximal at 0.5% O2
(19). Against this background,
hypoxia in cell culture represents an oxygen atmosphere below 5%, and
conventional culture exposes cells to significant hyperoxia. This results in
increased reactive oxygen metabolite production, which is positively
correlated with mitochondrial oxygen tension without a threshold level
(3). Oxygen metabolites have
been implicated in the inactivation of XOR
(37), which is compatible with
the finding that XOR activities of cultured endothelial cells were negatively
correlated with ambient oxygen tension over a wide range below 21%
(38), with no threshold at
around 5% as shown for HIF activation.
The rate of proliferation and the final density of cultured cells are known to be markedly lower at 20% than at 23% oxygen (26), which may be due to detrimental effects of hyperoxia. Therefore, application of the terms hypoxia, normoxia, and hyperoxia in cell culture is not only a semantic question but requires careful consideration of the underlying physiological conditions. We show here one example of a reduced protein function under typical cell culture conditions, but many more are likely to be discovered.
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ACKNOWLEDGMENTS |
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This study was supported by the Sigrid Jusélius Foundation, Finska Läkaresällskapet (N. Linder), Medicinska understöds-föreningen Liv och Hälsa (N. Linder), Svenska kulturfonden (N. Linder), and the Helsinki Biomedical Graduate School, University of Helsinki (E. Martelin).
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FOOTNOTES |
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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. Section 1734 solely to indicate this fact.
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