Posttranslational inactivation of human xanthine oxidoreductase by oxygen under standard cell culture conditions

Nina Linder, Eeva Martelin, Risto Lapatto, and Kari O. Raivio

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


    ABSTRACT
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Xanthine oxidoreductase (XOR) catalyzes the final reactions of purine catabolism and may account for cell damage by producing reactive oxygen metabolites in cells reoxygenated after hypoxia. We found a three- to eightfold higher XOR activity in cultured human bronchial epithelial cells exposed to hypoxia (0.5–3% O2) compared with cells grown in normoxia (21% O2) but no difference in XOR protein or mRNA. XOR promoter constructs failed to respond to hypoxia. The cellular XOR activity at 3% O2 returned to basal levels when the cells were returned to 21% O2, and hyperoxia (95% O2) abolished enzyme activity with no change in XOR protein. Our data suggest reversible enzyme inactivation by oxygen or its metabolites. NADH was normally oxidized by the oxygen-inactivated enzyme, which rules out damage to the flavin adenine dinucleotide cofactor. Hydrogen peroxide partially inactivated the molybdenum center of XOR, as shown by a parallel decrease in XOR-catalyzed xanthine oxidation and dichlorophenolindophenol reduction. We conclude that the transcription or translation of XOR is not influenced by hypoxia or hyperoxia. Instead, the molybdenum center of XOR is posttranslationally inactivated by oxygen metabolites in "normal" (21% O2) cell culture atmosphere. This inactivation is reversed in hypoxia and accounts for the apparent induction.

xanthine oxidase; hypoxia; hyperoxia; ischemia reperfusion


IN ADDITION TO ITS PHYSIOLOGICAL EFFECTS at the level of the whole organism, hypoxia has profound cellular effects that are mediated by altered activity and expression of proteins (4). These changes prepare the cell to cope with a shortage of oxygen, and they are mediated by several different mechanisms. Stabilization of the {alpha}-subunit of hypoxia-inducible factor 1 (HIF-1) upon exposure to low oxygen tension results in increased cellular levels of the heterodimeric HIF-1 protein and transcriptional activation of several genes, including erythropoietin, vascular endothelial growth factor (VEGF), glucose transporter-1 (GLUT-1), and several glycolytic enzymes (9, 17, 18, 29). Another mechanism is posttranscriptional through increased stability of mRNA, which has been documented for VEGF, GLUT-1, and tyrosine hydroxylase (4). Stabilization of the iron-regulatory protein 2 in hypoxia increases its RNA binding activity and protects the target mRNA from degradation (10).

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.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Cell cultures. SV40-transformed normal human bronchial epithelial cells (BEAS-2B) were obtained from the American Type Culture Collection (Manassas, VA) and cultured in serum-free, hormone-supplemented bronchial epithelial cell growth medium (Cytotech ApS, Hellebaek, Denmark). Human embryonic kidney cells (293T) (from Dr. Kalle Saksela, University of Tampere, Finland) were grown in Dulbecco's modified Eagle's medium (DMEM) supplemented with penicillin (50 IU/ml), streptomycin (50 µg/ml), and 10% fetal bovine serum. The standard or normoxic atmosphere was 21% oxygen (O2) and 5% CO2 at 37°C. For subculture, the cells were detached with 1% trypsin/EDTA. If not otherwise specified, all reagents were purchased from Sigma (St. Louis, MO).

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 (4–48 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 (2–24 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 405–789 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 {beta}-actin cRNA probe (pTRI-{beta}-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/{beta}-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{beta} (Clontech, Palo Alto, CA), producing {beta}-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 {beta}-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.


    RESULTS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Effect of hypoxia on XOR activity. XOR activity in BEAS-2B cells grown at 21% O2 increased circa twofold after subculture until confluence (data not shown) and then decreased (Fig. 1, time 0 to 48). Therefore, exposures to 3 or 0.5% O2 were started at confluence for 4, 24, and 48 h, after which intracellular XOR activities were measured. No change in XOR activity was detected after 4 h of incubation in 3% O2 (data not shown). However, after 24 h of exposure to 3% O2, XOR activity was threefold compared with normoxic (21% O2) timed controls, and after 48 h the difference was eightfold (Fig. 1). After 24 h of exposure to 0.5% O2, XOR activity was threefold, compared with normoxic (21% O2) timed controls, and after 48 h the difference was sixfold. Compared with preexposure activity, there was apparent slight induction in cells grown in 3% O2, but not in cells grown in 0.5% O2. XO represented ~20% of the total XDH + XO activity in all oxygen concentrations. There was some batch-to-batch variability in the basal XOR activity of BEAS-2B cells, but the pattern of the oxygen responses was always similar.



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Fig. 1. Hypoxia increases xanthine oxidoreductase (XOR) activity. Confluent cultures of human bronchial epithelial cells (BEAS-2B) were exposed to normal (21%) or hypoxic (3% or 0.5%) O2 atmosphere for 24 or 48 h. Total XOR enzyme activity (dehydrogenase and oxidase) was measured. Each bar represents means ± SD of 3 replicate wells in one of at least 3 experiments showing a similar pattern. ***P < 0.001 compared with timed controls in normoxia.

 

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{alpha}-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{alpha} 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{alpha} 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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Fig. 2. Hypoxia does not have an effect on XOR promoter-driven transcription. 293T cells were transfected with XOR promoter luciferase constructs of the indicated lengths (XOR1, XOR2, XOR4, XOR5), vector pGL3-basic alone, or 3 tandem repeats of the erythropoietin hypoxia-responsive element, coupled to a luciferase reporter gene (HRE-luc). After a change of medium 16 h posttransfection, the cells were exposed for 24 h to normoxia or hypoxia, followed by assay of luciferase and {beta}-galactosidase. The bars show fold induction of luciferase activity relative to {beta}-galactosidase activity in cells exposed to 0.5% O2 compared with cells exposed to 21% O2. The data are means ± SD of 3 independent experiments, each performed in triplicate.

 

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/{beta}-actin ratios (Fig. 3A).



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Fig. 3. Effect of hypoxia and hyperoxia on XOR mRNA and protein. A: mRNA levels in BEAS-2B cells before hypoxic exposure (0 h) and after being exposed to 3% O2 for the indicated times and then processed for ribonuclease protection assay (RPA). The intensity of the band corresponding to the protected XOR mRNA fragment compared with the {beta}-actin control was not increased as a function of time of exposure, as indicated by the calculated XOR/{beta}-actin ratio. Representative results from 1 experiment are shown. B: amount of XOR protein in cells grown for 24 or 48 h in 21% O2 or in 3% O2. After exposure, the cells were processed for assay of XOR protein with ELISA using polyclonal anti-XOR antibody. Each bar represents means ± SD of 3 replicate wells in 1 of at least 3 independent experiments showing similar results. C: Western blot analysis of XOR in BEAS-2B cells exposed for 24 h to 21% O2 (lanes 2 and 4), for 24 h to 95% O2 (lane 3), or for 24 h (lane 5) or 48 h (lane 6) to 3% O2. Molecular mass markers are shown in lane 1. The main band of XOR protein at 143 kDa is not altered under any of the conditions. Representative results from 1 experiment are shown.

 

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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Fig. 4. Effect of reoxygenation on XOR enzyme activity. BEAS-2B cells were grown at either 21 or 3% O2 for 24 h, the latter then either continuing at 3 or at 21% O2 for another 24 h. At the end of the experiment, the cells were processed and total XOR enzyme activity (dehydrogenase and oxidase) was measured. Each bar represents means ± SD of 3 replicate wells in 1 of at least 3 experiments showing similar results. ***P < 0.001 compared with controls.

 

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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Fig. 5. XOR enzyme activity and protein in hyperoxia. A: total XOR enzyme activity (dehydrogenase and oxidase) in confluent cultures of BEAS-2B cells exposed to either 95% O2 or to 21% O2 for the indicated times. Each bar represents means ± SD of 3 replicate wells in 1 of at least 3 experiments showing similar results. ***P < 0.001 compared with timed controls. B: amount of XOR protein in cells grown for 24 h in 95% O2 or in 21% O2. After exposure, the cells were processed for assay of XOR protein with ELISA using polyclonal anti-XOR antibody. Each bar represents means ± SD of 3 replicate wells in 1 of at least 3 experiments showing similar results.

 

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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Fig. 6. Effect of hydrogen peroxide on xanthine oxidation and dichlorophenolindophenol (DCPIP) reduction by XOR. A: urate formation from xanthine, and B: DCPIP reduction after exposure of purified XOR to hydrogen peroxide (30 or 90 mM) for the indicated times. Xanthine oxidase (XO) enzyme activity was measured spectrophotometrically at 295 nm and DCPIP reduction at 600 nm. The data are means ± SD of 3 experiments, each performed in duplicate. *P < 0.05; ***P < 0.001 compared with controls.

 


    DISCUSSION
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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Although a consensus exists that XOR activity is elevated in cultured cells exposed to hypoxia, the mechanism of the elevation is controversial. The finding that cycloheximide does not block the hypoxic increase in XOR activity suggests that new protein synthesis is not required (11, 28). XOR mRNA levels have been reported to be either unchanged (28) or elevated (11, 39), but in both of the latter studies only after enzyme activity had already increased. Discrepancies in the published reports may be due to cell type-specific responses, variations in experimental conditions, or species differences.

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 30–35 mmHg for the liver (43), 20–30 mmHg for the renal cortex (7), 20 mmHg for the cerebral cortex (41), and 12–17 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{alpha} 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 2–3% 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.


    ACKNOWLEDGMENTS
 
We thank J. Palvimo and M. Saksela for methodological advice and R. Löfman and S. Lindén for technical assistance.

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).


    FOOTNOTES
 

Address for reprint requests and other correspondence: N. Linder, Research Program for Developmental and Reproductive Biology, Univ. of Helsinki, Biomedicum Helsinki, Rm. B524b, Haartmaninkatu 8, 00290 Helsinki, Finland. (E-mail: nina.linder{at}hus.fi).

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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