INVITED REVIEW
Reactive species mechanisms of cellular hypoxia-reoxygenation injury

Chuanyu Li and Robert M. Jackson

Department of Veterans Affairs Medical Center, Birmingham 35233; and University of Alabama at Birmingham, Birmingham, Alabama 35294


    ABSTRACT
TOP
ABSTRACT
BACKGROUND
ROS MEDIATE REOXYGENATION...
INTRACELLULAR SOURCES OF ROS...
ANTIOXIDANT DEFENSE MECHANISMS
RNS MEDIATE HYPOXIA-...
ROLE OF MITOCHONDRIA IN...
REACTIVE SPECIES IN...
REACTIVE SPECIES MECHANISMS OF...
SUMMARY AND CONCLUSIONS
REFERENCES

Exacerbation of hypoxic injury after restoration of oxygenation (reoxygenation) is an important mechanism of cellular injury in transplantation and in myocardial, hepatic, intestinal, cerebral, renal, and other ischemic syndromes. Cellular hypoxia and reoxygenation are two essential elements of ischemia-reperfusion injury. Activated neutrophils contribute to vascular reperfusion injury, yet posthypoxic cellular injury occurs in the absence of inflammatory cells through mechanisms involving reactive oxygen (ROS) or nitrogen species (RNS). Xanthine oxidase (XO) produces ROS in some reoxygenated cells, but other intracellular sources of ROS are abundant, and XO is not required for reoxygenation injury. Hypoxic or reoxygenated mitochondria may produce excess superoxide (O<UP><SUB>2</SUB><SUP>−</SUP></UP>) and release H2O2, a diffusible long-lived oxidant that can activate signaling pathways or react vicinally with proteins and lipid membranes. This review focuses on the specific roles of ROS and RNS in the cellular response to hypoxia and subsequent cytolytic injury during reoxygenation.

anoxia; ischemia; reperfusion


    BACKGROUND
TOP
ABSTRACT
BACKGROUND
ROS MEDIATE REOXYGENATION...
INTRACELLULAR SOURCES OF ROS...
ANTIOXIDANT DEFENSE MECHANISMS
RNS MEDIATE HYPOXIA-...
ROLE OF MITOCHONDRIA IN...
REACTIVE SPECIES IN...
REACTIVE SPECIES MECHANISMS OF...
SUMMARY AND CONCLUSIONS
REFERENCES

Scope and Goals of the Review

This review focuses on reactive oxygen (ROS) and nitrogen species (RNS)-mediated mechanisms that lead to cellular injury during hypoxia and reoxygenation. Pathophysiology due to hypoxia per se is addressed when relevant to reoxygenation in the cellular context. Emphasis is on intracellular mechanisms that generate ROS and RNS and their roles in cellular signaling or as mediators of cell death.

Clinical Importance of Ischemia-Reperfusion Injury

Both hypoxia (lack of oxygen relative to metabolic needs) and reoxygenation (reintroduction of oxygen to hypoxic tissue) are important in human pathophysiology because they occur in a wide variety of important clinical conditions. Prominent examples of tissue hypoxia that predispose to injury during reoxygenation include circulatory shock, myocardial ischemia, stroke, and transplantation of organs (67, 76, 102). Because diseases due to ischemia (e.g., myocardial infarction and stroke) are exceedingly common causes of morbidity and mortality and because organ transplantation is increasingly frequent, understanding the role of ROS and RNS in reoxygenation injury has the potential to lead to therapies that could improve public health. Cellular models of hypoxia-reoxygenation have provided useful tools for the study of reactive species-mediated mechanisms of cellular dysfunction in ischemia-reperfusion injury (127).

Cellular Hypoxia and Reoxygenation Injury

Cellular necrosis inevitably follows extended periods of anoxia (i.e., oxygen absent) or severe hypoxia (i.e., oxygen supply decreased relative to metabolic demand). Hypoxic tolerance of various cell types differs, depending on the metabolic rate and intrinsic adaptive mechanisms of the tissue. Sublethal hypoxia, which may be transient and have no apparent consequences, can be followed by enhanced resistance to reoxygenation injury (conditioning), recovery, or exacerbated cellular injury (reoxygenation injury). Posthypoxic injury is due to a combination of changes in cellular energy charge, oxidant generating systems, and antioxidant defenses (76).

Cellular hypoxia appears to be a key signal that activates transcriptional regulators, including hypoxia-inducible factor-1 (HIF-1) (109), nuclear factor-kappa B (NF-kappa B), activator protein 1 (AP-1), and some mitogen-activated protein kinase (MAPK) pathways (46). A redox-sensitive human antioxidant response element induces gene expression in response to low oxygen concentrations in some malignant cells (124). Two overlapping manifestations of cell death, necrosis and apoptosis, can be initiated by cellular hypoxia-reoxygenation (105).

Cells undergo specific changes in enzyme activities, mitochondrial function, cytoskeletal structure, membrane transport, and antioxidant defenses in response to hypoxia, which then collectively predispose to reoxygenation injury (1). In contrast to the adaptive effects of sublethal hyperoxia on ion transport (74), hypoxia downregulates proteins that maintain alveolar ion transport, including the Na+-K+-ATPase (sodium pump) and the epithelial Na+ channel (ENaC) (23). Hypoxia causes time- and concentration-dependent decreases in alpha -, beta -, and gamma -subunits of ENaC mRNA and decreases both beta 1- and alpha 1-subunits of the Na+-K+-ATPase. Hypoxia itself also impairs cation transport in both A549 lung epithelial cells and rat alveolar cells (73). Oxidative inhibition of membrane Na+-K+- ATPase activity by H2O2 produced as a result of hypoxia-reoxygenation may be an important mechanism leading to swelling and cytolysis (52). In lung hypoxia-reoxygenation injury (e.g., after lung transplantation), reduced transepithelial Na+ transport and fluid resorption by the alveolar epithelium would increase pulmonary edema formation and impair its clearance.

A number of mitochondrial enzymes, including cytochrome oxidase and manganese superoxide dismutase (Mn SOD), decrease in activity in hypoxia (81, 101, 113), with predicted effects on oxygen metabolism. Cellular hypoxia inhibits expression of the multisubunit cytochrome oxidase (complex IV), the final intramitochondrial site of oxidative phosphorylation. Cytochrome oxidase activity of aerobic mouse lung macrophages decreases ~40% when incubated anaerobically for 96 h (113). Loss of cytochrome oxidase activity leads to cellular injury during reoxygenation, because absence of the final electron acceptor increases ROS production by more proximal complexes (33).

Cytoskeletal changes occurring in hypoxia would likely alter endothelial and epithelial permeability. ATP-depleted endothelial cells display shortening and disassembly of F-actin filaments (41), which lead to increased endothelial permeability (71). Hypoxia-reoxygenation has specific effects on the cytoskeleton of renal epithelial cells (88), which influence translational motion of membrane lipids. Cellular hypoxia may cause membrane protein aggregation, alterations in protein polarization (altered apical-basolateral orientation), protein degradation, and changes in molecular chaperones or growth factors. Renal epithelial cells develop increased membrane permeability, because of changes in transmembrane adhesion molecules, in a coordinated response characterizing the ischemic phenotype (12).

McCord (76) proposed the seminal model to explain how reoxygenation worsens ischemic (i.e., hypoxic) injury through increased ROS production. Superoxide radical (O<UP><SUB>2</SUB><SUP>−</SUP></UP>), generated by xanthine oxidase (XO), was postulated initially to be the ROS responsible, but now the entire spectrum of ROS and RNS [including peroxynitrite (ONOO-) (7)] has been implicated in posthypoxic cellular injury. ROS generated by hypoxia or reoxygenation are now recognized as interacting with physiological signal transducers (19, 46, 104, 110) rather than behaving as simple reactants that peroxidize membrane lipids, oxidize DNA, or denature enzyme proteins.

Free radicals (defined chemically as molecules containing an odd number of electrons) (33), such as O<UP><SUB>2</SUB><SUP>−</SUP></UP>, the hydroxyl radical (·OH), and other ROS and RNS species (especially ONOO-) are detected during, and likely account for, some manifestations of postischemic injury (Table 1). Perhaps of more physiological importance, both ROS and RNS can affect signal transduction in posthypoxic cells, and ROS are able to initiate cell death programs in the form of apoptosis or necrosis. Diverse sources of ROS (e.g., enzymes, mitochondria) exist normally within cells, some of which produce excess reactive species during hypoxia-reoxygenation. Excess ROS from endogenous sources can account for autocrine and paracrine cellular injury during reoxygenation.

                              
View this table:
[in this window]
[in a new window]
 
Table 1.   Important mechanisms and sequelae of cellular reoxygenation injury


    ROS MEDIATE REOXYGENATION INJURY
TOP
ABSTRACT
BACKGROUND
ROS MEDIATE REOXYGENATION...
INTRACELLULAR SOURCES OF ROS...
ANTIOXIDANT DEFENSE MECHANISMS
RNS MEDIATE HYPOXIA-...
ROLE OF MITOCHONDRIA IN...
REACTIVE SPECIES IN...
REACTIVE SPECIES MECHANISMS OF...
SUMMARY AND CONCLUSIONS
REFERENCES

General evidence for involvement of ROS in hypoxia-reoxygenation injury includes detection of lipid peroxidation and protein nitration products in reperfused brains (117), protection of various reperfused organs by antioxidant enzymes including SOD (76), and inhibition of postischemic injury by allopurinol, an XO inhibitor structurally related to purines (91). ROS (including ·OH radical) have been confirmed by electron paramagnetic resonance and spin trapping to occur in reoxygenated endothelial cells (135).

Neutrophils are important sources of ROS, but activated neutrophils are not required for reoxygenation injury. Injury to cultured endothelial cells, cardiac myocytes, hepatocytes, and other cell types occurs after anoxia-reoxygenation in vitro, even in the absence of neutrophils. Endothelial cells themselves subjected to anoxia-reoxygenation release superoxide anions (O<UP><SUB>2</SUB><SUP>−</SUP></UP>) into the extracellular medium, as demonstrated by SOD-inhibitable, extracellular cytochrome c reduction and other assays (108). The quantity of O<UP><SUB>2</SUB><SUP>−</SUP></UP> produced by reoxygenated cells depends on the duration of both anoxia and reoxygenation (99).

In vitro reoxygenation decreased the viability of rat hepatocytes as a function of time in hypoxia (26). Extracellular SOD and catalase completely prevented reoxygenation injury, confirming involvement of at least extracellular ROS. Electron transport chain inhibitors cyanide and antimycin A increased the severity of cellular injury in this model, suggesting that mitochondria may be a source of ROS (27). Reoxygenated hepatocytes released increased quantities of O<UP><SUB>2</SUB><SUP>−</SUP></UP> detected by oxidation of a nonspecific probe, lucigenin. Antimycin A, an inhibitor that increases mitochondrial O<UP><SUB>2</SUB><SUP>−</SUP></UP> production from complex III, increased both lucigenin chemiluminescence and lactate dehydrogenase (LDH) release (15). Inhibition of lucigenin oxidation by diphenyliodonium implicates a membrane NAD(P)H oxidase as the source of O<UP><SUB>2</SUB><SUP>−</SUP></UP>. Neurons obtained from spontaneously hypertensive rats, which are stroke prone, also generate increased ROS after reoxygenation. Neuronal injury is inhibited by mitochondrial accumulation of vitamin E, a lipid-soluble antioxidant that inhibits membrane lipid peroxidation (116).

Renal proximal tubular cells produce increased ROS [7-fold increase by 2',7'-dichlorofluorescein (DCF) fluorescence] after hypoxia-reoxygenation. Excess ROS appear capable of injuring the cells (89), because increased lipid peroxidation after reoxygenation is prevented by addition of SOD, catalase, dimethylthiourea (a radical scavenger), or deferoxamine (an iron chelator).

Hypoxia-reoxygenation also stimulates vascular endothelial cells to release extracellular O<UP><SUB>2</SUB><SUP>−</SUP></UP> (118). The anion channel blocker DIDS, which prevents extracellular release of O<UP><SUB>2</SUB><SUP>−</SUP></UP> through anion channels, protected from reoxygenation injury as well as extracellular SOD did. Injury to reoxygenated endothelial cells appears to involve generation of ·OH by surface-associated iron that reacts with H2O2, because iron chelation decreased cellular injury.

Hypoxia Apparently Increases ROS Production

Oxidants are produced in excess during reoxygenation, but ROS production also may increase in the reduced state that characterizes cellular hypoxia. Pulmonary artery smooth muscle cells, cardiomyocytes, and several other cell types produce increased ROS in hypoxia that are usually detected as oxidation of the fluorescent probe DCF (54, 133). Hypoxic pulmonary artery smooth muscle cells significantly increased DCF fluorescence fivefold above that of normoxic cells (55), and simulated ischemia (hypoxia and low glucose) of embryonic ventricular myocytes increased DCF fluorescence threefold over that of normoxic cells (133). Simulated in vitro ischemia of cardiomyocytes also increased dihydroethidine (DHE) oxidation due to ROS, so these observations are not simply an artifact related to the use of oxidizable DCF. Both myxothiazole and rotenone inhibited the DHE oxidation, suggesting that mitochondrial complexes III and I produced increased ROS. SOD inhibited the DHE oxidation, providing indirect evidence that the mitochondrial electron transport chain generated O<UP><SUB>2</SUB><SUP>−</SUP></UP> during in vitro ischemia, probably from the ubisemiquinone site (6). Hypoxia-induced tyrosine hydroxylase gene expression also is paralleled by increased ROS production in PC-12 cells (42), as shown by increased oxidation of dihydrorhodamine.

Human lung epithelial cells likewise have been observed to increase ROS production significantly after 24-h incubation in hypoxia (<1% O2) (69). Examples of typical data from such experiments are shown in Fig. 1, which demonstrates increased DCF fluorescence in hypoxia-preexposed lung epithelial cells by both fluorescent microscopy and flow cytometry. The sulfhydryl compound N-acetylcysteine and the antioxidants ebselen (a peroxide scavenger) and Tempol (a superoxide scavenger) inhibited DCF fluorescence. Ebselen was the most effective at inhibiting the DCF signal from reoxygenated cells, suggesting that peroxides (probably H2O2) accounted for the most of the increased DCF fluorescence.


View larger version (40K):
[in this window]
[in a new window]
 
Fig. 1.   To test whether hypoxia preexposure increased cellular reactive oxygen species (ROS) production, we exposed cells to hypoxia (<1% O2) for 24 h. 2',7'-Dichlorofluorescein (DCF) fluorescence was then measured by flow cytometry. Fluorescent microscopic images of previously hypoxia H441 cells stained with DCF are shown below corresponding flow cytometry data. Hypoxia preexposure per se increased epithelial DCF fluorescence. As reported in the text, N-acetylcysteine, ebselen, and Tempol all inhibited the DCF fluorescence increase due to hypoxia, indicating a role for oxidants in production of the fluorescent signal.

Hypoxia preexposure appears to increase cellular ROS production, probably from mitochondrial electron transport complexes. Excess ROS and RNS production has been well documented during reoxygenation. In vitro experiments have demonstrated that hypoxia-reoxygenation is sufficient to cause injury to various cell types, even in the absence of activated neutrophils. Both inhibitor (antioxidant) studies and detection of lipid and protein oxidation have confirmed a role for endogenously generated reactive species in cellular reoxygenation injury.


    INTRACELLULAR SOURCES OF ROS DURING HYPOXIA-REOXYGENATION
TOP
ABSTRACT
BACKGROUND
ROS MEDIATE REOXYGENATION...
INTRACELLULAR SOURCES OF ROS...
ANTIOXIDANT DEFENSE MECHANISMS
RNS MEDIATE HYPOXIA-...
ROLE OF MITOCHONDRIA IN...
REACTIVE SPECIES IN...
REACTIVE SPECIES MECHANISMS OF...
SUMMARY AND CONCLUSIONS
REFERENCES

Xanthine Dehydrogenase-Oxidase

Two closely related enzymes potentially capable of ROS generation, aldehyde oxidase and XO, exist in most animals. Activities are concentrated mainly in the liver and intestine (63). XO is the more important source of both O<UP><SUB>2</SUB><SUP>−</SUP></UP> and H2O2, but XO varies widely in abundance among cell types, organs, and species. XO normally occurs in vivo as an NAD+-dependent dehydrogenase [xanthine dehydrogenase (XD)] incapable of ROS production. XD activity converts by sulfhydryl oxidation or limited proteolysis (conditions that exist during reoxygenation) to an oxidase that produces both O<UP><SUB>2</SUB><SUP>−</SUP></UP> and H2O2 (91). XO is phosphorylated in hypoxic microvascular endothelial cells through a mechanism involving p38 MAPK and casein kinase II, and phosphorylation appears to be necessary for hypoxia-induced enzyme activation (54). However, reoxygenation injury also unquestionably occurs in cells in which XO activity is exceedingly low or absent. For example, human and rabbit hearts produce only minute quantities of urate because of their extremely low XO activities, yet both species develop myocardial infarctions and ischemia-reperfusion injury (82).

Rat tissues contain relatively high levels of XD-XO activity. Rat pulmonary endothelial cells are protected equally from reperfusion injury by antioxidant enzymes or allopurinol, confirming that XO is an important source of ROS during reoxygenation (1, 99). XO also appears to be responsible for reoxygenation-induced O<UP><SUB>2</SUB><SUP>−</SUP></UP> formation and injury of rat kidney cells. Rat proximal tubule epithelial cells subjected to 60-min hypoxia and 30-min reoxygenation released increased quantities of LDH, whereas XO inhibitors allopurinol and dietary tungsten, which depletes XO activity, attenuated both injury and O<UP><SUB>2</SUB><SUP>−</SUP></UP> production (37).

Ferrylhemoglobin

Hemoglobin and myoglobin, which are released into plasma after trauma, can mediate endothelial cell oxidative stress, and a ferrylhemoglobin intermediate has been detected in cellular hypoxia-reoxygenation models. Incubation of hemoglobin (Hb) or myoglobin during hypoxia-reoxygenation with endothelial cells causes transient oxidation of Hb to the reactive ferryl species (Fe4+). Lipid peroxidation, which results from exposure of endothelial cells to ferrylhemoglobin, increased after reoxygenation. Incubation of endothelial cells with Hb also caused a dose-dependent decrease in intracellular glutathione (GSH), confirming an oxidative stress (78). Nitric oxide (·NO) acts as an antioxidant to inhibit lipid peroxidation and injury due to ferrylhemoglobin, because 400 µM arginine inhibited both lipid peroxidation and formation of ferryl intermediate (3). Hemoproteins oxidized to their ferryl forms could account for heme-mediated injury of endothelial cells that have been stressed by hypoxia, reoxygenation, and glutathione depletion.

NADPH Oxidase

Endothelial cells express a membrane NADPH oxidase, similar to the membrane enzyme complex that produces the respiratory burst in granulocytes. The membrane oxidase complex is an important source of ROS in ischemic mouse lungs (4) and probably other reoxygenated tissues. ROS produced by lungs exposed to normoxic ischemia or high-K+ buffer (24 mM) emanate from the membrane-associated NADPH oxidase complex in the vascular endothelial cells. p47phox, a subunit of the NAPDH oxidase, is detected by Western blotting of endothelial proteins. Endothelial cells adapted to flow in vitro for 2-7 days showed a nearly twofold increase in ROS production during simulated no-flow ischemia, compared with continuously perfused cells (129). Absence of flow causes flow-adapted cells to activate both NF-kappa B and AP-1. Transcription factor activation appears to be dependent on transduction of mechanical signals rather than altered oxygenation, showing that ischemia rather than hypoxia is the more important determinant in flow-adapted cells. In contrast, in isolated perfused lungs, anoxia-reoxygenation rather than ischemia-reperfusion causes XO to produce ROS (4).

The GTPase binding protein Rac1 regulates the membrane NADPH oxidase that produces ROS during mouse liver ischemia-reperfusion. A dominant negative Rac1 construct completely inhibited ischemia-reperfusion induced ROS production, NF-kappa B activation, and liver necrosis (86). The dominant negative gene product (N17rac1) inhibits the intracellular ROS burst after reoxygenation. N17rac1 expression protects smooth muscle cells, fibroblasts, endothelial cells, and ventricular myocytes from hypoxia-reoxygenation-induced death (58), indicating an important role for Rac1. A constitutively active Rac1 mutant does not increase intracellular ROS. Rac1 GTPase is, therefore, a necessary but not sufficient component of the pathway leading to ROS production during reoxygenation (87).

Multiple sources of ROS exist with cells and may account for increased reactive species production during reoxygenation. Although XD was identified early as a source of ROS during reoxygenation, other intracellular sources including redox cycling of iron and NAD(P)H oxidases have been demonstrated to produce ROS and contribute to oxidant stress during reoxygenation.


    ANTIOXIDANT DEFENSE MECHANISMS
TOP
ABSTRACT
BACKGROUND
ROS MEDIATE REOXYGENATION...
INTRACELLULAR SOURCES OF ROS...
ANTIOXIDANT DEFENSE MECHANISMS
RNS MEDIATE HYPOXIA-...
ROLE OF MITOCHONDRIA IN...
REACTIVE SPECIES IN...
REACTIVE SPECIES MECHANISMS OF...
SUMMARY AND CONCLUSIONS
REFERENCES

Antioxidants Inhibit Cellular Reoxygenation Injury

Endogenous antioxidant systems are critically important in limiting reoxygenation-induced cellular damage. The preponderance of data shows that exogenous antioxidant enzymes and low-molecular-weight ROS scavengers inhibit reoxygenation injury in cellular models. However, studies employing scavengers do not by themselves identify specific ROS or RNS responsible for reoxygenation injury, and they do not exclude other mechanisms. The level and duration of hypoxia are themselves important in regulating levels of both the antioxidant enzymes and low-molecular-weight ROS scavengers. including glutathione, which ultimately might determine the extent of reoxygenation injury (60, 81, 112).

Reoxygenation (after 12-min hypoxia) of hippocampal slices in vitro increased LDH release and lipid peroxidation significantly. Exogenous SOD and catalase inhibited injury, demonstrating involvement of ROS (43), which appeared to arise from prostaglandin synthesis, XO, and mitochondria, on the basis of the inhibitor profile. O<UP><SUB>2</SUB><SUP>−</SUP></UP> is generated at complex III during reoxygenation (13), and antimycin A (a mitochondrial complex III inhibitor) typically increases its production. Antimycin A increased reoxygenation injury, as would be predicted if increased mitochondrial O<UP><SUB>2</SUB><SUP>−</SUP></UP> production were responsible. Antimycin A is a quinone analog that binds to cytochrome b562 and blocks transfer of electrons to the ubiquinone. It thus increases the complex III semiquinone concentration, which donates single electrons to oxygen.

Further indirect evidence from inhibitor studies for the involvement of ROS is that sodium salicylate, a nonspecific radical scavenger, prevents reoxygenation injury of rat livers perfused with hypoxic buffer (24). Livers released increased LDH and developed increased protein carbonyl and malondialdehyde content during reoxygenation. Salicylate (2 mM) inhibited development of markers of oxidant stress in reperfused livers, including LDH release, carbonyl formation, and malondialdehyde production.

Endogenous low-molecular-weight antioxidants are important in protection from reoxygenation injury. Thioredoxin (TRX; human T cell leukemia-derived factor) proteins act as disulfide oxidoreductases and electron donors for thioredoxin peroxidases. Thioredoxins function as disulfide bond reductants, similar in mechanism to N-acetylcysteine. Oxidized thioredoxin is reduced by thioredoxin reductases requiring NADPH.

Thioredoxin protects lung cells from reoxygenation injury. Viability of murine endothelial cells cultured in thiol-free medium decreased significantly after hypoxia-reoxygenation, but injury was diminished by exogenous TRX (100 µM). Because TRX does not decrease cellular ROS production or inhibit loss of GSH, it rather appears to act directly as a scavenger in reoxygenation models (47).

Effects of Hypoxia on Antioxidant Enzymes

Alveolar (AM) or peritoneal macrophages (PM) exist in high (PO2 > 100 mmHg) or low (PO2 < 20 mmHg) oxygen environments, respectively. Activities of the enzymes of oxidative phosphorylation and glycolysis differ significantly, with oxidative phosphorylation being higher in aerobic AM and glycolysis higher in hypoxic PM. Hypoxia decreases apparent expression of cytochrome oxidase in AM, a change in the electron transport system that would predispose to O<UP><SUB>2</SUB><SUP>−</SUP></UP> formation (101). The antioxidant enzymes SOD, glutathione peroxidase (GP), and catalase are all two- to threefold higher in aerobic AM than in hypoxic PM (113), apparently reflecting the different levels of oxygenation. Similar changes in antioxidant enzyme phenotype occur when normally aerobic cells are cultured in hypoxia (48, 59, 101).

Mitochondrial DNA content, reflecting the abundance of mitochondria, does not change significantly in rat skeletal muscle cells cultured in hypoxia (81). However, a number of mitochondrial enzymes in these cells and in lung macrophages decrease in a coordinated fashion in hypoxia. Citrate synthase, NAD-isocitrate dehydrogenase, malate dehydrogenase, and cytochrome oxidase all decreased significantly after incubation of cells in hypoxia. (Mn SOD activity was not measured.) These data suggest a loss of mitochondrial enzyme activities due to hypoxia without a change in the number of mitochondrial. Brain capillary endothelial cells similarly respond to hypoxia preexposure by decreasing activities of GP, glutathione reductase (GR), catalase, and SOD, as well as cellular total GSH content (94). Such changes, which represent possibly adaptive downregulation of antioxidant defenses by unspecified mechanisms in hypoxia, would predispose to increased ROS production by reoxygenated mitochondria.

Global ischemia of isolated rabbit hearts decreases the rate of oxidative phosphorylation but does not eliminate H2O2 production. Hypoxia-reoxygenation significantly decreased activities of SOD and GP by ~40% in mitochondria isolated from rabbit hearts (112). Glucose-free hypoxia, used to model ischemia in vitro, also resulted in significantly depressed SOD and GP activities in perfused rat hearts. Rat hearts hypertrophied due to chronic pressure overload had increased SOD and GP activities (on a per mg protein basis), which appeared biologically important during recovery. Yet, during hypoxia, SOD activity decreased significantly in both control and hypertrophied hearts (60). Similarly, hypoxic exposure of cardiac myocytes (30 min) in vitro also caused a decrease in Mn SOD and GP activities but caused no change in catalase activity (59).

The response of alveolar type II (ATII) cells in primary culture to hypoxia exemplifies the effects of hypoxia on antioxidant defenses. Exposure of isolated rabbit ATII cells to hypoxia in vitro (<1% O2 for 24 h) caused a significant decrease in Mn SOD activity and protein content (103). Mn SOD and Cu,Zn SOD mRNA expression also decreased significantly in ATII cells cultured in hypoxia for 24 h (48). The decrease in ATII cell Mn SOD mRNA expression (-69% compared with air controls by semiquantitative PCR) was greater than the decrease in Cu,Zn SOD mRNA expression. The decrease in Mn SOD mRNA content was due, in part, to decreased Mn SOD mRNA stability in the hypoxic ATII cells. Mn SOD enzyme specific activity did not change.

Hypoxia-induced decreases in cellular antioxidant enzymes, especially Mn SOD, have potentially important biological consequences because they could lead indirectly to cellular reoxygenation injury by exacerbating oxidant stress (69). LDH release from lung epithelial cells maintained in hypoxia (<1% O2) for 24 h before treatment with antimycin A plus carbonyl cyanide p-trifluoromethoxyphenylhydrazone (FCCP), an in vitro oxidant stress that increases mitochondrial O<UP><SUB>2</SUB><SUP>−</SUP></UP> production, was significantly greater than LDH release from vehicle-treated cells. The sulfhydryl antioxidant N-acetylcysteine (1 mM) significantly inhibited the increase in LDH release due to antimycin A plus FCCP.

The general response to hypoxia, therefore, appears to be decreased activity or expression of antioxidant defenses. Hypoxia-induced downregulation of antioxidant enzymes leads to reoxygenation injury, but the mechanism by which antioxidant enzyme expression is regulated by hypoxia is not known with certainty.


    RNS MEDIATE HYPOXIA-REOXYGENATION INJURY
TOP
ABSTRACT
BACKGROUND
ROS MEDIATE REOXYGENATION...
INTRACELLULAR SOURCES OF ROS...
ANTIOXIDANT DEFENSE MECHANISMS
RNS MEDIATE HYPOXIA-...
ROLE OF MITOCHONDRIA IN...
REACTIVE SPECIES IN...
REACTIVE SPECIES MECHANISMS OF...
SUMMARY AND CONCLUSIONS
REFERENCES

·NO Itself Can Cause Cellular Injury

Both ROS and RNS mediate some aspects of reoxygenation injury, but ·NO can produce either damaging or protective effects (49). In many cell types, inducible nitric oxide synthase (iNOS) is induced by hypoxia. iNOS induction during hypoxia is controlled by HIF-1 (51), a basic helix-loop-helix-PAS heterodimer regulated by oxygen tension (125). ·NO reacts rapidly with O<UP><SUB>2</SUB><SUP>−</SUP></UP> when both are produced simultaneously. The reaction product, peroxynitrite anion (ONOO-), is a potent oxidant that, when protonated (pKa 7.49; ONOOH), has ·OH radical-like reactivity (7). ONOO- nitrates protein tyrosine residues to form 3-nitrotyrosine (3-NT) in reoxygenated tissues, but alternate pathways of tyrosine nitration utilizing myeloperoxidase and H2O2 exist and probably predominate (122). Thus finding 3-NT in reoxygenated cells does not unequivocally identify ONOO- as the nitrating species, because nitrite (NO<UP><SUB>2</SUB><SUP>−</SUP></UP>) can react with peroxidase and H2O2 to form nitrogen dioxide (NO2·), which also contributes strongly to tyrosine nitration. Diverse cellular proteins may be nitrated during reoxygenation, but the specific functional consequences of most nitration reactions are unknown.

Human umbilical vein endothelial cells exposed to anoxia-reoxygenation release decreased quantities of prostacyclin (PGI2) in response to thrombin, calcium ionophore A-23187, or arachidonic acid (40). Endothelial cells decrease prostacyclin production during short (15 min) hypoxia exposures (126). Decreased cyclooxygenase activity is due, in part, to ROS, because SOD and catalase inhibit the decrease in PGI2 production. In addition, nitration of PGI2 synthase correlates with decreased PGI2 formation after reoxygenation. ONOO- apparently nitrates and inactivates PGI2 synthase, leaving unmetabolized prostaglandin H2 (PGH2), which can cause vasospasm, platelet aggregation, and thrombus formation by stimulating the thromboxane A2 (TXA2)/PGH2 receptor (134).

· NO itself may be cytotoxic, especially when present at supraphysiological concentrations (57). ·NO can react reversibly with enzyme 4Fe-4S (iron-sulfur) centers, yielding inactive 4Fe-4S-·NO derivatives. ·NO thus inactivates mitochondrial aconitase, NAD-ubiquinone oxidoreductase (complex I), and succinate-ubiquinone reductase (complex II), resulting in inhibited mitochondrial respiration (32). ·NO inhibits respiration directly by binding cytochrome oxidase. ·NO-derived reactants, especially ONOO-, also potentiate inflammation by inactivating Mn SOD protein through nitration of its tyrosine residues (72), as well as by depleting low-molecular-weight antioxidants like GSH and ascorbate.

·NO Protects Cells From Hypoxia-Reoxygenation Injury

In contrast to its cytotoxic effects, ·NO more often acts as an antioxidant and attenuates reoxygenation injury. ·NO is capable of inhibiting lipid peroxidation chain reactions, and ·NO has clear protective effects in many cellular hypoxia-reoxygenation models. Hypoxia preexposure increases iNOS mRNA in myocardial cells and protects against subsequent damage from prolonged hypoxia (98). ·NO from hippocampal constitutive NOS (cNOS) may be involved in neuroprotection afforded by hypoxic preconditioning, because NOS inhibition blocks the development of preconditioning after anoxia exposures (17).

Activation of cGMP-dependent pathways by ·NO appears to be protective. For example, L-arginine, an NOS substrate, limits myocardial cell death due to hypoxia-reoxygenation through a cGMP-dependent mechanism (2). L-Arginine increases cGMP in cardiac myocytes and inhibits LDH release due to reoxygenation (2). Cardiomyocytes preconditioned with 90 min of simulated ischemia (i.e., near anoxia and glucose-free medium) followed by 30-min reoxygenation were protected from subsequent ischemia (98). The NOS inhibitor NG-nitro-L-arginine methyl ester (L-NAME) blocked preconditioning, and the ·NO donor S-nitroso-N-acetyl-L-penicillamine (SNAP) protected cells from reoxygenation. Preconditioning required synthesis of ·NO and was cGMP-dependent, but protection due to ·NO appeared independent of protein kinase C (PKC) activation or ATP-sensitive K+ (KATP) channels. Similarly, decreased cNOS expression due to reoxygenation appears to be related to development of apoptosis of coronary artery endothelial cells. Anoxia-reoxygenation of the cells significantly decreased cNOS and Bcl-2, whereas it increased Fas expression (70).

The protective effects of ·NO depend critically on the PO2 and balance of ·NO and O<UP><SUB>2</SUB><SUP>−</SUP></UP> production (45). When perfused rabbit lungs were ventilated with N2 and subjected to ischemia-reperfusion, NOS inhibition decreased, whereas an NOS substrate increased, the degree of lung injury. In contrast, the ·NOS inhibitor L-NAME significantly worsened injury of lungs ventilated with air. Protective effects of N2 ventilation may be due to a lower rate of ·NO production or altered peroxynitrite production.

Consistent with apparent antioxidant effects of ·NO noted above, we observed that inhibition of endogenous NOS activity markedly worsens lung epithelial cell reoxygenation injury in vitro. Cultured, human lung epithelial cells were incubated with aminoguanidine (50 µM), an inhibitor of iNOS, during reoxygenation. iNOS inhibition markedly increased LDH release, suggesting that ·NO inhibits reoxygenation injury [similar results have been found with NG-monomethyl-L-arginine (L-NMMA)]. An example of such data is shown in Fig. 2. Similarly, L-NMMA significantly increased cytolysis of Mn SOD-deficient mouse lung fibroblasts (-/-) during reoxygenation (Jackson and Li, unpublished data), again suggesting that ·NO, probably derived from iNOS, modulates oxidative stress in reoxygenated lung cells.


View larger version (17K):
[in this window]
[in a new window]
 
Fig. 2.   Nitric oxide synthase (NOS) inhibitors increase lung epithelial cytolysis. To test the hypothesis that nitric oxide (·NO) might attenuate lung epithelial cell reoxygenation injury, H441 lung epithelial cells were incubated with the inducible NOS (iNOS) inhibitor aminoguanidine (50 µM) (4 bars at right) during reoxygenation. In the absence of aminoguanidine (4 bars at left), antimycin A and FCCP (Anti A + FCCP) caused a moderate increase in lactose dehydrogenase (LDH) release from hypoxia-preexposed cells (solid bars) compared with vehicle (veh)-treated, hypoxia-preexposed controls. Inhibition of iNOS significantly increased LDH release from both air controls (shaded bars) and hypoxia-preexposed cells (solid bars) exposed to the oxidant stress, suggesting that ·NO inhibits reoxygenation injury (similar results obtained with NG-monomethyl-L-arginine). These data are consistent with others in the literature showing a protective and apparently antioxidant effect of ·NO in cellular hypoxia-reoxygenation models. *P < 0.05 compared with the respective vehicle by ANOVA.

Effects of ·NO on Mitochondrial Function During Hypoxia-Reoxygenation

·NO has important effects on cellular respiration and mitochondrial ROS production, which could either contribute to or protect from hypoxia-reoxygenation injury. Cellular respiration is regulated by ADP availability to F2-ATPase, O2 availability to cytochrome oxidase, and the concentration of ·NO (10). ·NO participates in redox reactions in the mitochondrial matrix, which regulate both intramitochondrial ·NO concentration itself and production of other reactive species (O<UP><SUB>2</SUB><SUP>−</SUP></UP> and ONOO-). Exogenously produced ·NO markedly decreased O2 uptake of isolated rat heart mitochondria (50% inhibition at 4 µM ·NO) (96). ·NO inhibits cytochrome oxidase between cytochromes b and c. Binding at this site accounts for its inhibitory effects on O2 uptake. ·NO also regulates O<UP><SUB>2</SUB><SUP>−</SUP></UP> production, detected as H2O2. Mitochondrial ·NO is metabolized by pathways leading to ONOO- formation and ubiquinol oxidation and only secondarily by reversible binding to cytochrome oxidase (95). Whereas low concentrations of ·NO inhibit cytochrome oxidase, higher concentrations of ·NO inhibit other respiratory chain complexes, possibly by nitrosylation, oxidation of protein thiols, or disruption of Fe-S centers (11).

In contrast, ONOO- irreversibly inhibits mitochondrial complexes I, II, IV, and V, as well as aconitase. Decreased maximal uptake rate and increased affinity constant for O2 of the enzyme characterize ONOO- inhibition of cytochrome oxidase (111). Intramitochondrial ONOO- reacts with NADH, changing the mitochondrial redox state. ONOO- treatment of submitochondrial particles leads to ubisemiquinone autooxidation increasing O<UP><SUB>2</SUB><SUP>−</SUP></UP> production.

Mitochondrial proteins such as Mn SOD may be nitrated chemically by high concentrations of ONOO- (200 µM in vitro). However, the intramitochondrial steady-state concentration of ONOO- remains around 2 nM (121), indicating that this mechanism may be less relevant in vivo. Mitochondrial creatine kinase (CK) exists predominantly as an octamer, and ischemia decreases the ratio of octamers to dimeric forms and so inhibits CK activity. An identical decrease in CK activity and change in structure occurs after ONOO- exposure that inhibits transport of high-energy phosphate out of mitochondria, resulting in impaired cardiac performance (114).

Although ·NO can injure cells directly or after reaction with O<UP><SUB>2</SUB><SUP>−</SUP></UP> to from the oxidant ONOO-, much evidence indicates that ·NO is protective against cellular hypoxia-reoxygenation injury. Probable protective mechanisms include innate antioxidant properties of ·NO: increasing cellular cGMP, and partial inhibition of cellular respiration.


    ROLE OF MITOCHONDRIA IN HYPOXIA-REOXYGENATION INJURY
TOP
ABSTRACT
BACKGROUND
ROS MEDIATE REOXYGENATION...
INTRACELLULAR SOURCES OF ROS...
ANTIOXIDANT DEFENSE MECHANISMS
RNS MEDIATE HYPOXIA-...
ROLE OF MITOCHONDRIA IN...
REACTIVE SPECIES IN...
REACTIVE SPECIES MECHANISMS OF...
SUMMARY AND CONCLUSIONS
REFERENCES

Mitochondrial ROS Production

ROS and possibly RNS generated by mitochondria may damage the organelles themselves and other cellular constituents during reoxygenation. Protein carbonyls, a marker of oxidant stress, are detected in mitochondrial proteins after 10-min hypoxia and 5-min reoxygenation (100). The mitochondrial electron transport chain generates superoxide anions radicals (O<UP><SUB>2</SUB><SUP>−</SUP></UP>) during reoxygenation that dismutate rapidly either enzymatically or spontaneously to form H2O2, which reacts with iron to form hydroxyl radicals (·OH) (30). Mitochondrial oxidative phosphorylation, uncoupled respiration, and ADP-to-oxygen ratios are all decreased by excess O<UP><SUB>2</SUB><SUP>−</SUP></UP> during reoxygenation. The mitochondrial outer membrane also contains monoamine oxidase, which catalyzes oxidative deamination of biogenic amines and is itself a source of H2O2 (13).

Effects of Hypoxia on Mitochondrial ROS Production

The respiratory electron transport chain becomes reduced (i.e., the complexes harbor electrons) during anoxia, and the reduced state potentiates O<UP><SUB>2</SUB><SUP>−</SUP></UP> production (20). Mitochondria generate excess partially reduced oxygen species (primarily O<UP><SUB>2</SUB><SUP>−</SUP></UP> and its dismutation product, H2O2) during reoxygenation, which react with respiratory chain proteins (especially those containing Fe-S centers) (120) or diffuse as H2O2 out of mitochondria. H2O2 produced by mitochondria also might function as a signal transducer, allowing mitochondria to act as oxygen sensors (84) that regulate extramitochondrial signaling pathways.

Effects of Reoxygenation on Mitochondrial Function

Decreased Mn SOD activity, protein, or gene expression caused by hypoxia could influence the steady-state concentration of mitochondrial O<UP><SUB>2</SUB><SUP>−</SUP></UP> production. Antimycin A-sensitive generation of O<UP><SUB>2</SUB><SUP>−</SUP></UP> originates from the ubiquinone-cytochrome c reductase (complex III), specifically from the Qo semiubiquinone. When Mn SOD-depleted submitochondrial particles are inhibited with myxothiazole or stigmatellin, O<UP><SUB>2</SUB><SUP>−</SUP></UP> is detected by coelenterazine oxidation. Mn SOD eliminates ROS generated at the Qi site, but in the absence of Mn SOD, O<UP><SUB>2</SUB><SUP>−</SUP></UP> from that site is free to react (97). This leads to increased O<UP><SUB>2</SUB><SUP>−</SUP></UP> production, a situation that might occur if Mn SOD synthesis were inhibited (e.g., by hypoxia) or if Mn SOD were inactivated (e.g., by ONOO-).

Mitochondrial complex I dysfunction occurs after reoxygenation of hypoxic mitochondria (62). Complex I defects increase cellular production of ROS, which may influence Mn SOD expression (93). Reoxygenated kidney tubule cells develop energy deficits due to complex I dysfunction that occur before onset of the mitochondrial permeability transition (MPT) or loss of cytochrome c. Supplementation with citric acid cycle metabolites that anaerobically generate ATP may prevent energy deficits (125). Anaerobic metabolism of alpha -ketoglutarate and aspartate generate sufficient ATP to maintain mitochondrial membrane potential. Proximal tubules are protected from hypoxia-reoxygenation-induced mitochondrial injury by anaerobic metabolism of citric acid cycle intermediates and of succinate (130). Whereas complex I dysfunction occurs as a result of ROS produced during reoxygenation, substrates that bypass complex I provide sufficient energy to maintain viability of reoxygenated cells (131).

Transient, but large, cellular and mitochondrial Ca2+ fluxes occur during hypoxia-reoxygenation. Elevated intracellular Ca2+ concentration ([Ca2+]i) causes mitochondrial depolarization. Anoxia depolarizes guinea pig ventricular myocyte mitochondria, and recovery of normal mitochondrial membrane potential is essential to avoid hypercontracture. Reoxygenation caused significant elevation in intramitochondrial [Ca2+], the amplitude of which correlated with the extent of hypercontracture (28). Intramitochondrial [Ca2+] also increases markedly ([Ca2+]i > 350 nM) during rat cardiomyocyte hypoxia-reoxygenation, whereas cytosolic [Ca2+] falls. Myocyte mitochondrial Ca2+ uptake during hypoxia occurs largely through the Na+/Ca2+ exchanger, rather than the Ca2+ uniporter (38). Extracellular catalase inhibited the increase in myocyte [Ca2+] and [Na+] content and LDH release due to hypoxia-reoxygenation, indicating that extracellular H2O2 contributed to the ion fluxes.

Release of Ca2+ from storage sites stimulates Ca2+-dependent proteases, nucleases, and phospholipases that trigger apoptosis. Ca2+ leaves mitochondria during reoxygenation through pore formation, reversal of uniport influx carrier, the Ca2+/H+ antiport system, or channel-mediated Ca2+ pathways. Increased intracellular Ca2+ also appears to be related to oxidation of adenine nucleotides or by thiol oxidation, because overexpression of the antiapoptotic protein Blc-2, which has antioxidant properties, in mitochondrial membranes inhibits Ca2+ efflux due to oxidants (31).

The long-lived oxidant H2O2 stimulates human aortic endothelial cell [Ca2+] oscillations. Reoxygenation similarly initiates [Ca2+] oscillations, which depend on Ca2+ release from the intracellular pool and require extracellular Ca2+. The Ca2+ oscillations caused by NADPH oxidase-derived H2O2 appear to play an important role in signal transduction (44). Removal of Ca2+ from the medium or Ca2+ chelation prevented killing of rat hepatocytes by rotenone and anoxia, but not by cyanide. Ca2+ depletion prevented the MPT in anoxic or rotenone-treated cells, and Ca2+ influx is required to kill cells after complex I inhibition (92). Although anoxia is associated with mitochondrial Ca2+ influx, hypocalcemic medium does not always prevent reoxygenation injury. Low extracellular Ca2+ at the time of reoxygenation significantly worsened cellular injury of rat hepatocytes (61). When the Na+/H+ exchanger and Na+-HCO<UP><SUB>3</SUB><SUP>−</SUP></UP> symporter were both inhibited, intracellular pH recovery did not occur normally, although Ca2+ oscillations and hypercontracture were attenuated (107).

Preconditioning involves both the KATP channel and ROS production. KATP channel antagonists and antioxidants blocked both preconditioning and ROS production. Transfer and expression of genes encoding ATPase channel subunits render COS-7 monkey kidney cells resistant to hypoxia-reoxygenation injury (50).

Mitochondrial complex III is an important site of O<UP><SUB>2</SUB><SUP>−</SUP></UP> production during hypoxia-reoxygenation. Reoxygenation impairs mitochondrial complex I function and inhibits energy production. Reoxygenation causes increased ROS production and a significant increase in mitochondrial calcium concentration. Mitochondrial Ca2+ fluxes may be associated with the activation of intracellular proteases that trigger apoptosis.


    REACTIVE SPECIES IN INTRACELLULAR SIGNALING
TOP
ABSTRACT
BACKGROUND
ROS MEDIATE REOXYGENATION...
INTRACELLULAR SOURCES OF ROS...
ANTIOXIDANT DEFENSE MECHANISMS
RNS MEDIATE HYPOXIA-...
ROLE OF MITOCHONDRIA IN...
REACTIVE SPECIES IN...
REACTIVE SPECIES MECHANISMS OF...
SUMMARY AND CONCLUSIONS
REFERENCES

ROS act as signaling molecules in various cell types (46, 119), participating in or modifying physiological events related to receptor-ligand binding and transcriptional activation (35, 39, 90). Intracellular signaling pathways are implicated in cell death following reoxygenation, although the specific pathway leading to apoptosis or necrosis might vary among cell types. Several protein kinase signaling pathways involved in cellular reoxygenation injury are shown in Fig. 3. ROS interact with a number of specific molecular targets in reoxygenated cells, including extracellular signal-regulated kinases (ERK) and MAPK that mediate proliferation, stress-activated protein kinases (SAPK) implicated in apoptosis, NF-kappa B, and several caspases (132). The GTPase binding protein Rac1 regulates some kinases [e.g., SAPK, p38 MAPK, and c-Jun NH2-terminal kinases (JNK)]. These kinases may be regulated further by ROS from NAD(P)H oxidase, which requires Rac1 for activity (46). Inhibition of Rac1 lessens both ROS production and reoxygenation injury (58).


View larger version (24K):
[in this window]
[in a new window]
 
Fig. 3.   Protein kinase signaling pathways possibly involved in reoxygenation injury. The mitogen-activated protein kinases (MAPK), which phosphorylate target molecules at threonine-X-tyrosine domains, have been implicated in cellular hypoxia-reoxygenation models. GTPase binding proteins and kinases experimentally implicated in cellular reoxygenation injury models are shown in bold italics. Hypoxia or reoxygenation appears to activate the c-Jun NH2-terminal kinase (JNK) and p38 kinase pathways. During reoxygenation, protein kinase C-zeta is activated by phosphoinositide 3-kinase, which induces MAPK activation through MEK1/2. As noted in the text, permeability changes in a cell culture model due to hypoxia and low glucose (simulated ischemia) could be blocked by PKC and PKG inhibition or by inhibition of p38 MAPK-1 (89). RNS, reactive nitrogen species.

ROS also regulate or participate in growth, apoptosis, and the adaptive response to injury or stress (35). Platelets exposed to anoxia-reoxygenation generate O<UP><SUB>2</SUB><SUP>−</SUP></UP> and ·OH, which activate arachidonic acid metabolism via phospholipases A2 and C (66). Platelet activation is associated with inositol 1,3,4-trisphosphate and TXA2 production, and platelet activation is inhibited by antioxidants.

Stress Kinases, JNK, and AP-1

ROS produced during reoxygenation appear to stimulate SAPK, including JNK and p38 MAPK (65). Anoxia followed by hyperoxia (reoxygenation) causes apoptosis of mouse embryo fibroblast (NIH/3T3) cells. Fibroblast arrest in G1 during anoxia was reversed on reoxygenation. c-jun and c-fos expression increased during anoxia. AP-1 binding increased markedly during reoxygenation, as did poly(ADP-ribose) polymerase (PARP) cleavage and caspase-3 activation. Induction of c-jun/c-fos (AP-1) expression during hypoxia was followed by PARP activation and histone H1 ADP-ribosylation. Activation of the AP-1 pathway was essential to initiation of programmed cell death (21).

Hypoxia-reoxygenation of cardiac myocytes caused a 10-fold increase in phosphorylation of the c-jun transcription factor. c-jun activation correlated with a decrease in GSH content, indicating that AP-1 activation occurred in association with oxidant stress. JNK activity was inhibited by a free radical spin trap (alpha -phenyl-N-tert-butylnitrone) and N-acetylcysteine, a sulfhydryl compound. Phosphorylation and activation of c-Jun protein are linked directly to intracellular redox status (65). SAPK activation also leads to H2O2-induced apoptosis and upregulation of intercellular adhesion molecule (ICAM)-1 (46).

Supporting evidence for activation of kinase pathways by ROS during reoxygenation includes inhibition of hypoxia-reoxygenation-induced apoptosis by an antisense oligonucleotide targeted to JNK-1. Human kidney cells in vitro undergo apoptosis during reoxygenation after 48-h hypoxia. Reoxygenation causes a time-dependent increase in activation of JNK-1, while p38 MAPK is activated by hypoxia. Activation of the JNK signaling cascade leads to apoptosis during hypoxia-reoxygenation (34).

Transient hypoxia causes apoptosis of developing rat brain neurons due to activation of AP-1-related transcription factors and JNK kinases. AP-1 expression increases in neurons during hypoxia. JNK-1 and JNK-3 are induced after 48-h reoxygenation. Hypoxia exposure increased c-Jun, Jun B, Jun D, c-Fos, and Fos-related protein expression. JNK-1 and JNK-3 both increased transiently 48 h after reoxygenation, when apoptosis occurred (22).

JNK-1 activity translocated to the nucleus increases during reperfusion, at which time JNK-1 is serine-phosphorylated. An upstream kinase, SAPK/ERK kinase 1 (SEK1), activates JNK-1 in the nucleus. Another upstream kinase for the MAPK, MAPK/ERK kinase 1 (MEK1), in contrast, remains localized to the cytosol. Increased JNK-1 activity after ischemia depends on its translocation to the nucleus and activation of upstream protein kinases (probably SEK1) during reoxygenation (80).

Protein Kinase C

PKC-zeta acts upstream of the MAPK. PKC-zeta induces MAPK activity through MEK activation during reoxygenation. PKC-zeta is itself activated by phosphoinositide 3-kinase, which induces MAPK as a result of MEK1/2 activation in the nucleus of reoxygenated cardiomyocytes (79). Immunocytochemical staining of PKC-zeta in nuclei of cardiomyocytes increases when MAPK is phosphorylated. Phosphorylation and activation of MAPK during reoxygenation can be blocked both by PKC inhibition and phosphoinositide 3-kinase inhibitors. Activation of PKC also appears to be involved in the development of preconditioning or tolerance. Simulated ischemic preconditioning of endothelial cells prevented subsequent reoxygenation-induced expression of ICAM-1 associated with activation of PKC and production of NOS and ROS (5).

Chemical hypoxia (produced by the reducing agent thioglycolic acid) and lack of glucose increased permeability of cultured human dermal endothelial cells. Intracellular [Ca2+] increased as well under those conditions. Permeability changes due to hypoxia and low glucose could be blocked by chelating Ca2+, PKC, and protein kinase G (PKG) inhibition or by inhibition of p38 MAPK-1. Ca2+-induced dissociation of cadherin-actin and occludin-actin junctions might mediate increases in endothelial permeability (88), through pathways that involve PKC, PKG, and MAPK.

Ischemic preconditioning of hepatocytes decreases Na+ accumulation during hypoxia and decreases cellular death, as does PKC activation (16). PKC appears to activate H+ extrusion and maintain intracellular pH.

Nuclear Factor-kappa B

NF-kappa B activation is involved in hypoxia-reoxygenation injury, especially in the vascular endothelium, where NF-kappa B activation leads to neutrophil adhesion in vivo (56). NF-kappa B nuclear binding increased in microvascular endothelial cells during reoxygenation but not during hypoxia. NF-kappa B binding and ICAM-1 surface expression were induced, concurrent with ROS production. An ·NO donor molecule, SNAP, blocked NF-kappa B activation and inhibited ICAM-1. Blockade of NOS by L-nitroarginine increased ICAM-1 expression after reperfusion. ·NO thus appears to inhibit ICAM-1 activation by decreasing the level of oxidant stress responsible for NF-kappa B activation (64).

NF-kappa B activation and nuclear translocation is required for development of tolerance to anoxia-reoxygenation (18). Inhibition of ·NO synthase during a second anoxia-reoxygenation challenge prevents both inhibition of polymorphonuclear neutrophil adherence and NF-kappa B activation. Thus, while NF-kappa B activation is associated with the vascular inflammatory response in vivo, NF-kappa B activation is required for the development of tolerance to hypoxia-reoxygenation in vitro.

Reactive species may act as intracellular signals, or they may interact with specific molecules of intracellular signaling cascades. Stress-activated protein kinases, particularly JNK and p38 MAPK, have been most strongly associated with hypoxia-reoxygenation injury. PKC-zeta appears to be responsible for induction of MAPK activity during reoxygenation. NF-kappa B activation by hypoxia-reoxygenation can be blocked by ·NO, and NF-kappa B appears to be required for development of tolerance to hypoxia-reoxygenation.


    REACTIVE SPECIES MECHANISMS OF CELL DEATH DURING REOXYGENATION
TOP
ABSTRACT
BACKGROUND
ROS MEDIATE REOXYGENATION...
INTRACELLULAR SOURCES OF ROS...
ANTIOXIDANT DEFENSE MECHANISMS
RNS MEDIATE HYPOXIA-...
ROLE OF MITOCHONDRIA IN...
REACTIVE SPECIES IN...
REACTIVE SPECIES MECHANISMS OF...
SUMMARY AND CONCLUSIONS
REFERENCES

Apoptosis

Both apoptosis and necrosis (and likely intermediate forms of cell death, as well) are linked to excess intracellular ROS production (70). ROS, for example, mediate PARP cleavage and lead to apoptosis (77). Apoptosis initiated by oxidant stress involves increased mitochondrial permeability and release of cytochrome c (14). Downstream signaling and enzymatic mechanisms, including activation of caspases, are all regulated by oxidant mechanisms (29). Apoptosis depends on the availability of cellular ATP, so reoxygenation could potentiate apoptosis by restoring cellular energy (77). Antioxidants inhibit DNA fragmentation and PARP cleavage, suggesting that ROS are responsible for these manifestations (79). Hypoxia-reoxygenation itself enhances expression of CD95L (APO-1/Fas), a transmembrane protein that induces apoptosis by ligand binding and subsequent activation of caspases (123).

Bcl-2 may exert some of its antiapoptotic effects through apparent antioxidant actions, because Bcl-2 inhibited H2O2 production due to ceramide or tumor necrosis factor-alpha . Bcl-2 overexpression allows cells to adapt to ROS overproduction (31). Apoptosis in an endothelial reperfusion model also required activation of NF-kappa B, because NF-kappa B decoy oligodeoxynucleotides decreased the number of TUNEL (TdT-mediated dUTP nick end labeling)-positive cells. NF-kappa B augments apoptosis by suppressing the antiapoptotic Bcl-2 expression in hypoxia (75).

Bax and p53 are induced in some cell types that undergo apoptosis after hypoxia. Proapoptotic Bax and the interleukin-converting enzyme subfamily of caspases increased after exposure of neurons to hypoxia followed by reoxygenation (8). Induction of Bax and p53 expression preceded neuronal apoptotic death.

Apoptosis observed after in vitro reoxygenation of cardiomyocytes is associated with activation of caspases-3 and -9 and appearance of cytochrome c in the cytosol. Initiation of apoptosis depends on mitochondrial release of a caspase-activating factor (probably cytochrome c); Bcl-2 overexpression blocks both cytochrome c release and activation of caspases-3 and -9 from the reoxygenated cells (53). Hypoxia-reoxygenation decreases Bcl-2 expression and increases Fas protein, changes associated with cardiomyocyte apoptosis and necrosis. Caspase inhibition and Bcl-2 overexpression inhibit apoptosis during reoxygenation, but they do not inhibit death due to hypoxia.

Hypoxia causes accumulation of p53, as well as hypophosphorylated retinoblastoma protein (pRB), in S-phase melanoma cells. p53 protein was higher in S phase than in G1 or G2, while pRB was not cell-cycle dependent. However, hypoxic induction of p53 did not arrest cell growth (25). Hypoxia also induces p53 and enhances apoptosis of cultured trophoblasts. Apoptosis in hypoxic trophoblasts involves alterations in p53 and Bax expression, because both are increased in hypoxia, whereas Bcl-2 is decreased (68). In contrast to the proapoptotic effects of p53, epithelial growth factor and enhanced differentiation protect cells from hypoxia-induced apoptosis (68).

Hypoxia per se is insufficient to initiate apoptosis in some cells. Apoptosis of rapidly contracting hypoxic myocytes occurred only when extracellular pH decreased. Reoxygenation of the hypoxic acidotic myocytes induced apoptosis in 23-30% of cells, independently of changes in p53 expression. Therefore, exogenous lactic acid during reoxygenation can induce programmed cell death without an increase in cellular p53 (128).

Bax protein translocates to mitochondria during reoxygenation and triggers cell death. Maintenance of glucose during hypoxia prevented Bax translocation to mitochondria and reoxygenation injury, in addition to maintaining intracellular ATP levels. Likewise, Bcl-2 overexpression in kidney tubule cells minimized apoptosis. Bcl-2 overexpression preserved mitochondrial integrity and prevented both apoptosis (glucose present) and necrosis (glucose absent), suggesting a common pathway involving mitochondrial dysfunction in cell death (104).

Hypoxic preconditioning activates transcription of Bcl-2 and heat shock protein 70 (HSP70), which are apparently protective, in neurons. Preconditioning occurs, in part, by expression of antiapoptotic gene products including Bcl-2, HSP-70, and regulatory components of the cell cycle (9). Likewise, overexpression of HSP-70 attenuates hypoxic injury of coronary endothelial cells (115). HSP-72 appears to be an important part of the normal response to stress, protecting from hypoxia-reoxygenation. HSP-72 translocates to the cytoskeleton after of hypoxia-reoxygenation (106). Inhibition of HSP-72 using antisense oligonucleotides increases reoxygenation injury of cardiomyocytes (83).

Mitochondria are critical to initiation of apoptosis in reoxygenated cells. Increased ROS production, cytochrome c release from mitochondria, and cell death during reoxygenation are linked. Mitochondria from apoptotic cells produce increased quantities of O<UP><SUB>2</SUB><SUP>−</SUP></UP> because of a switch from normal four-electron reduction of O2 to one-electron reduction after cytochrome c release (14). Overexpression of the antiapoptotic protein Bcl-2 prevents increased O<UP><SUB>2</SUB><SUP>−</SUP></UP> production associated with apoptosis. Loss of cytochrome c from the mitochondrial matrix leads to activation of caspases, which participate in the apoptotic program. Caspases-3, -8, and -9 are activated by hypoxia, possibly through Ca2+-induced activation of other proteases. Serine protease inhibitors suppress cytochrome c-mediated caspase-9 activation and apoptosis during hypoxia-reoxygenation (29).

Necrosis

Necrosis is associated with tissue inflammation, but cytolysis occurs in cell culture reoxygenation models as well. Inhibition of apoptosis, because of lack of energy due to hypoxia-induced ATP depletion, may predispose to necrosis, and mechanisms of cell death typically overlap. DNA "laddering" into 180-bp fragments may occur in necrosis, although nucleosomal laddering is more characteristic of apoptosis. Multiple assays, including morphological examination, are required to define the actual characteristics (apoptotic, necrotic, or overlap) of cell death due to reoxygenation.

Human lung epithelial cells clearly undergo necrosis rather than apoptosis because of excess superoxide production in mitochondria after preexposure to hypoxia (1% O2 for 24 h). The Mn SOD activity of the cell decreases by ~50% after 24 h in hypoxia, as observed previously in ATII cells (103). The mechanism of decreased Mn SOD expression involves decreased stability of Mn SOD mRNA rather than direct inhibition of the Mn SOD promoter (85). ATP content did not change significantly, so cellular injury or lack of apoptosis could not be attributed to lack of energy per se. Antimycin A and FCCP generate excess mitochondrial O<UP><SUB>2</SUB><SUP>−</SUP></UP> and cause necrosis in the hypoxia-preexposed cells. Neither hypoxia (24 h) nor reoxygenation (24-h air following hypoxia) by itself increased cellular LDH release significantly. Cell death rarely was associated with annexin V staining (1-3%), even at early time points during the oxidant stress. The epithelial cells did not display increased Hoechst nuclear staining (a morphological marker of apoptosis), PARP fragmentation, caspase-3 activation, or change in p53 protein levels on Western blots (69).

Cellular injury after reoxygenation was associated with increased ROS production. N-acetylcysteine (1 mM), a sulfhydryl compound, partially inhibited increased DCF fluorescence after hypoxia and protected the lung epithelial cells from cytolysis during incubation with antimycin A and FCCP. Inhibition of both ROS production and cytolytic damage by N-acetylcysteine supports the notion that oxidants produced by the epithelial cells are involved in reoxygenation injury. Thus lung epithelial cells may undergo necrosis directly without preliminary apoptosis, as a result of increased susceptibility to oxidant stress after hypoxia preexposure.

Thus both apoptosis and necrosis occur after hypoxia-reoxygenation injury, and both appear dependent on increased reactive species production. Apoptosis after reoxygenation is associated with activation of caspases, and both caspase inhibition and Blc-2 overexpression inhibit apoptosis. Mitochondria are involved in posthypoxic apoptosis through increased ROS production and cytochrome c release. Cells are capable of undergoing necrosis directly after reoxygenation, and necrosis is also related to increased ROS production.


    SUMMARY AND CONCLUSIONS
TOP
ABSTRACT
BACKGROUND
ROS MEDIATE REOXYGENATION...
INTRACELLULAR SOURCES OF ROS...
ANTIOXIDANT DEFENSE MECHANISMS
RNS MEDIATE HYPOXIA-...
ROLE OF MITOCHONDRIA IN...
REACTIVE SPECIES IN...
REACTIVE SPECIES MECHANISMS OF...
SUMMARY AND CONCLUSIONS
REFERENCES

Adequate oxygenation is critical to cellular viability. The "paradox" of reoxygenation injury can be understood in terms of counteradaptive changes occurring in hypoxia that predispose to cellular dysfunction, apoptosis, and necrosis during reoxygenation. These changes include enhanced reduction of the mitochondrial respiratory chain, conversion of XD to XO, downregulation of antioxidant systems, increased reactive species formation, and activation of signaling pathways. Specific molecular mechanisms and pathways involving ROS and RNS that lead to necrosis or apoptosis during reoxygenation have been identified. Knowledge of these pathways has led to clinical attempts at minimizing reoxygenation injury in transplantation and myocardial reperfusion.


    ACKNOWLEDGEMENTS

We thank M. Kennedy for transcribing the manuscript.


    FOOTNOTES

This research was supported by The Veterans Affairs Research Service, National Institutes of Health Grants HL-57801 and DK-97010, Office of Naval Research Grant N00014-97-1-03-09, and an American Heart Association-Alabama Affiliate Grant-in-Aid.

Address for reprint requests and other correspondence: R. Jackson, Division of Pulmonary, Allergy and Critical Care Medicine, Univ. of Alabama at Birmingham, Rm. 215 Tinsley Harrison Tower, 1900 Univ. Boulevard, Birmingham, AL 35294-0006 (E-mail: rjackson{at}uab.edu).

10.1152/ajpcell.00112.2001


    REFERENCES
TOP
ABSTRACT
BACKGROUND
ROS MEDIATE REOXYGENATION...
INTRACELLULAR SOURCES OF ROS...
ANTIOXIDANT DEFENSE MECHANISMS
RNS MEDIATE HYPOXIA-...
ROLE OF MITOCHONDRIA IN...
REACTIVE SPECIES IN...
REACTIVE SPECIES MECHANISMS OF...
SUMMARY AND CONCLUSIONS
REFERENCES

1.   Acosta, D, and Li CP. Injury to primary cultures of rat heart endothelial cells by hypoxia and glucose deprivation. In Vitro 15: 929-934, 1979[ISI][Medline].

2.   Agullo, L, Garcia-Dorado D, Inserte J, Paniagua A, Pyrhonen P, Llevadot J, and Soler-Soler J. L-Arginine limits myocardial cell death secondary to hypoxia-reoxygenation by a cGMP-dependent mechanism. Am J Physiol Heart Circ Physiol 276: H1574-H1580, 1999[Abstract/Free Full Text].

3.   Agnillo, FD, Wood F, Porras C, Macdonald VW, and Alayash AI. Effects of hypoxia and glutathione depletion on hemoglobin- and myoglobin-mediated oxidative stress toward endothelium. Biomed Biochim Acta 1495: 150-159, 2000.

4.   Al-Mehdi, AB, Zhao G, Dodia C, Tozawa K, Costa K, Muzykantov V, Ross C, Blecha F, Dinauer M, and Fisher AB. Endothelial NADPH oxidase as the source of oxidants in lungs exposed to ischemia or high K+. Circ Res 83: 730-737, 1998[Abstract/Free Full Text].

5.   Beauchamp, P, Richard C, Tamion F, Lallemand F, Lebreton JP, Vaudry H, Daveau M, and Thuillez C. Protective effects of preconditioning in cultured rat endothelial cells: effects on neutrophil adhesion and expression of ICAM-1 after anoxia and reoxygenation. Circulation 100: 541-546, 1999[Abstract/Free Full Text].

6.   Becker, LB, Vanden Hoek TL, Shao ZH, Ch. Li Q, and Schumacker PT. Generation of superoxide in cardiomyocytes during ischemia before reperfusion. Am J Physiol Heart Circ Physiol 277: H2240-H2246, 1999[Abstract/Free Full Text].

7.   Beckman, JS, Beckman TW, Chen J, Marshall PA, and Freeman BA. Apparent hydroxyl radical production by peroxynitrite: implications for endothelial injury from nitric oxide and superoxide. Proc Natl Acad Sci USA 87: 1620-1624, 1990[Abstract].

8.   Bossenmeyer, C, Chihab R, Muller S, Schroeder H, and Daval JL. Hypoxia/reoxygenation induces apoptosis through biphasic induction of protein synthesis in cultured rat brain neurons. Brain Res 787: 107-116, 1998[ISI][Medline].

9.   Bossenmeyer-Pourie, C, Koziel V, and Daval JL. Effects of hypothermia on hypoxia-induced apoptosis in cultured neurons from developing rat forebrain: comparison with preconditioning. Pediatr Res 47: 385-391, 2000[Abstract/Free Full Text].

10.   Boveris, A, Costa LE, Poderoso JJ, Carreras MC, and Cadenas E. Regulation of mitochondrial respiration by oxygen and nitric oxide. Ann NY Acad Sci 899: 120-135, 2000.

11.   Brown, GC. Nitric oxide and mitochondrial respiration. Biochim Biophys Acta 1411: 351-369, 1999[ISI][Medline].

12.   Bush, T, Keller SH, and Nigam SK. Genesis and reversal of the ischemic phenotype in epithelial cells. J Clin Invest 106: 621-626, 2000[Free Full Text].

13.   Cadenas, E, and Davies KJA Mitochondrial free radical generation, oxidative stress and aging. Free Radic Biol Med 29: 222-230, 2000[ISI][Medline].

14.   Cai, J, and Jones DP. Superoxide in apoptosis. J Biol Chem 273: 11401-11404, 1998[Abstract/Free Full Text].

15.   Carceni, P, Ryu HS, Van Theil DH, and Borle AB. Source of oxygen free radicals produced by rat hepatocytes during postanoxic reoxygenation. Biomed Biochim Acta 1268: 249-254, 1995.

16.   Carini, R, De Cesaris MG, Spendore R, Bagnati M, and Albano E. Ischemic preconditioning reduces Na+ accumulation and cell killing in isolated rat hepatocytes exposed to hypoxia. Hepatology 31: 166-172, 2000[ISI][Medline].

17.   Centeno, JM, Orti M, Salom JB, Sick TJ, and Perez-Pinzon MA. Nitric oxide is involved in anoxic preconditioning neuroprotection in rat hippocampal slices. Brain Res 836: 62-69, 1999[ISI][Medline].

18.   Cepinskas, G, Lush CW, and Kvietys PR. Anoxia/reoxygenation-induced tolerance with respect to polymorphonuclear leukocyte adhesion to cultured endothelial cells: a nuclear factor-kappa B-mediated phenomenon. Circ Res 84: 103-112, 1999[Abstract/Free Full Text].

19.   Chakraborti Das, S, Mondal M, Roychoudhury S, and Chakraborti S. Oxidants mitochondria and calcium: an overview. Cell Signal 11: 77-85, 1999[ISI][Medline].

20.   Chance, B, and Williams GR. The respiratory chain and oxidative phosphorylation. Adv Enzymol Relat Subj 17: 65-134, 1956[ISI].

21.   Chen, YC, Tsai S-H, Lin-Shiau S-Y, and Lin J-K. Elevation of apoptotic potential by anoxia-hyperoxia shift in NIH3T3 cells. Mol Cell Biochem 197: 147-159, 1999[ISI][Medline].

22.   Chihab, R, Ferry C, Koziel V, Monin P, and Daval JL. Sequential activation of activator protein-1 related transcription factors and JNK protein kinases may contribute to apoptotic death induced by transient hypoxia in developing brain neurons. Mol Brain Res 63: 105-120, 1998[ISI][Medline].

23.   Clerici, C, and Matthay MA. Hypoxia regulates gene expression of alveolar epithelial transport proteins. J Appl Physiol 88: 1890-1896, 2000[Abstract/Free Full Text].

24.   Colantoni, A, de Maria N, Caraceni P, Bernardi M, Floyd RA, and Van Thiel DH. Prevention of reoxygenation injury by sodium salicylate in isolated-perfused rat liver. Free Radic Biol Med 25: 87-94, 1998[ISI][Medline].

25.   Danielsen, T, Hvidsten M, Stokke T, Solberg K, and Rofstad EK. Hypoxia induces p53 accumulation in the S-phase and accumulation of hypophosphorylated retinoblastoma protein in all cell cycle phases of human melanoma cells. Br J Cancer 78: 1547-1558, 1998[ISI][Medline].

26.   De Groot, H, Anundi I, and Littauer A. Hypoxia-reoxygenation injury and the generation of reactive oxygen in isolated hepatocytes. Biomed Biochim Acta 48: S11-S15, 1989[ISI][Medline].

27.   De Groot, H, and Brecht M. Reoxygenation injury in rat hepatocytes: Mediation by O2/H2O2 liberated by sources other than xanthine oxidase. Biol Chem 372: 35-41, 1991.

28.   Delcamp, TJ, Dales C, Ralenkotter L, Cole PS, and Hadley RW. Intramitochondrial [Ca2+] and membrane potential in ventricular myocytes exposed to anoxia-reoxygenation. Am J Physiol Heart Circ Physiol 275: H484-H494, 1998[Abstract/Free Full Text].

29.   Dong Saikumar, ZP, Patel Y, Weinberg JM, and Venkatachalam MA. Serine protease inhibitors suppress cytochrome c-mediated caspase-9 activation and apoptosis during hypoxia-reoxygenation. Biochem J 347: 669-677, 2000[ISI][Medline].

30.   Du, G, Mouithys-Mickalad A, and Sluse FE. Generation of superoxide anion by mitochondria and impairment of their functions during anoxia and reoxygenation and reoxygenation in vitro. Free Radic Biol Med 25: 1066-1074, 1998[ISI][Medline].

31.   Esposti, MD, Hatzinisiriou I, McLennan H, and Ralph S. Bcl-2 and mitochondrial oxygen radicals. J Biol Chem 27: 29831-29837, 1999.

32.   Forfia, PR, Hintze TH, Wolin MS, and Kaley G. Role of nitric oxide in the control of mitochondrial function. Adv Exp Med Biol 471: 381-388, 1999[ISI][Medline].

33.   Fridovich, I. The biology of oxygen radicals: the superoxide radical is an agent of oxygen toxicity. Superoxide dismutases provide an important defense. Science 201: 875-880, 1978[ISI][Medline].

34.   Garay, M, Gaarde W, Monia BP, Nero P, and Cioffi CL. Inhibition of hypoxia/reoxygenation-induced apoptosis by an antisense oligonucleotide targeted to JNK1 in human kidney cells. Biochem Pharmacol 59: 1033-1043, 2000[ISI][Medline].

35.   Gebhardt, BR, Ries J, Caspary WF, Boehles H, and Stein J. Superoxide: a major factor for stress protein induction in reoxygenation injury in the intestinal cell line Caco-2. Digestion 60: 238-245, 1999[ISI][Medline].

37.   Greene, EL, and Paller MS. Xanthine oxidase produces O<UP><SUB>2</SUB><SUP>−</SUP></UP> in post hypoxic injury of renal epithelial cells. Am J Physiol Renal Fluid Electrolyte Physiol 263: F251-F255, 1992[Abstract/Free Full Text].

38.   Griffiths, EJ, Ocampo CJ, Savage JS, Rutter GA, Hansford RG, Stern MD, and Silverman HS. Mitochondrial calcium transporting pathways during hypoxia and reoxygenation in single rat cardiomyocytes. Cardiovasc Res 39: 423-433, 1998[ISI][Medline].

39.   Hanson, ES, and Leibold EA. Regulation of iron regulatory protein 1 during hypoxia and hypoxia/reoxygenation. J Biol Chem 273: 7588-7593, 1998[Abstract/Free Full Text].

40.   Hempel, SL, Haycraft DL, Hoak JC, and Spector AA. Reduced prostacyclin formation after reoxygenation of anoxic endothelium. Am J Physiol Cell Physiol 259: C738-C745, 1990[Abstract/Free Full Text].

41.   Hinshaw, DB, Armstrong BC, Beals TF, and Hyslop PA. A cellular model of endothelial cell ischemia. J Surg Res 44: 527-537, 1988[ISI][Medline].

42.   Hohler, B, Lange B, Holzapfel B, Goldenberg A, Hanze J, Sell A, Testan H, Moller W, and Kummer W. Hypoxic upregulation of tyrosine hydroxylase gene expression is paralleled, but not induced, by increased generation of reactive oxygen species in PC12 cells. FEBS Lett 457: 53-56, 1999[ISI][Medline].

43.   Horakova, L, Stolc S, Chromikova Z, Pekarova A, and Derkova L. Mechanisms of hippocampal reoxygenation injury. Treatment with antioxidants. Neuropharmacology 36: 177-184, 1997[ISI][Medline].

44.   Hu, Q, and Ziegelstein RC. Hypoxia/reoxygenation stimulates intracellular calcium oscillations in human aortic endothelial cells. Circulation 102: 2541-2547, 2000[Abstract/Free Full Text].

45.   Huang, YCT, Fisher PW, Nozik-Grayck E, and Piantadosi CA. Hypoxia compared with normoxia alters the effects of nitric oxide in ischemia-reperfusion lung injury. Am J Physiol Lung Cell Mol Physiol 273: L504-L512, 1997[Abstract/Free Full Text].

46.   Irani, K. Oxidant signaling in vascular cell growth, death, and survival: a review of the roles of oxygen species in smooth muscle and endothelial cell mitogenic and apoptotic signaling. Circ Res 87: 179-183, 2000[Abstract/Free Full Text].

47.   Isowa, N, Yoshimura T, Kosaka S, Liu M, Hitomi S, Yodoi J, and Wada H. Human thioredoxin attenuates hypoxia-reoxygenation injury of murine endothelial cells in a thiol-free condition. J Cell Physiol 183: 33-40, 2000.

48.   Jackson, R, Parish G, and Ho YS. Effects of hypoxia on expression of superoxide dismutases in cultured ATII cells and lung fibroblasts. Am J Physiol Lung Cell Mol Physiol 271: L955-L962, 1996[Abstract/Free Full Text].

49.   Jessup, JM, Battle P, Waller H, Edmiston KH, Stolz DB, Watkins SC, Locker J, and Skena K. Reactive nitrogen and oxygen radicals formed during hepatic ischemia-reperfusion kill weakly metastatic colorectal cancer cells. Cancer Res 59: 1825-1829, 1999[Abstract/Free Full Text].

50.   Jovanovic, A, Jovanovic S, Carrasco AJ, and Terzic A. Acquired resistance of a mammalian cell line to hypoxia-reoxygenation through cotransfection of Kir6.2 and SUR1 clones. Lab Invest 78: 1101-1107, 1998[ISI][Medline].

51.   Jung, F, Palmer LA, Zhou N, and Johns RA. Hypoxic regulation of inducible nitric oxide synthase via hypoxia inducible factor-1 in cardiac myocytes. Circ Res 86: 319-325, 2000[Abstract/Free Full Text].

52.   Kako, K, Kato M, Matsuoka T, and Mustapha A. Depression of membrane-bound Na+-K+-ATPase activity induced by free radicals and by ischemia of kidney. Am J Physiol Cell Physiol 254: C330-C337, 1988[Abstract/Free Full Text].

53.   Kang, PM, Haunstetter A, Aoki H, Usheva A, and Izumo S. Morphological and molecular characterization of adult cardiomyocyte apoptosis during hypoxia and reoxygenation. Circ Res 87: 118-125, 2000[Abstract/Free Full Text].

54.   Kayyali, U, Donaldson C, Huang H, Abdelnour R, and Hassoun P. Phosphorylation of xanthine dehydrogenase/oxidase in hypoxia. J Biol Chem 276: 14359-14365, 2001[Abstract/Free Full Text].

55.   Killilea, DW, Hester R, Balczon R, Babal P, and Gillespie MN. Free radical production in hypoxic pulmonary artery smooth muscle cells. Am J Physiol Lung Cell Mol Physiol 279: L408-L412, 2000[Abstract/Free Full Text].

56.   Kim, CD, Kim YK, Lee SH, and Hong KW. Rebamipide inhibits neutrophil adhesion to hypoxia/reoxygenation-stimulated endothelial cells via nuclear factor-kappa B-dependent pathway. J Pharmacol Exp Ther 294: 864-869, 2000[Abstract/Free Full Text].

57.   Kim, H, and Kim KH. Role of nitric oxide in oxidative damage in isolated rabbit gastric cells exposed to hypoxia-reoxygenation. Dig Dis Sci 43: 1042-1049, 1998[ISI][Medline].

58.   Kim, KS, Takeda K, Sethi R, Pracyk JB, Tanaka K, Shou YF, Yu ZX, Ferrans VJ, Bruder JT, Kovesdi I, Irani K, Goldschmidt-Clermont P, and Finkel T. Protection from reoxygenation injury by inhibition of rac1. J Clin Invest 101: 1821-1826, 1998[Abstract/Free Full Text].

59.   Kirshenbaum, LA, and Singal PK. Changes in antioxidant enzymes in isolated cardiac myocytes subjected to hypoxia-reoxygenation. Lab Invest 67: 796-803, 1992[ISI][Medline].

60.   Kirshenbaum, LA, and Singal PK. Antioxidant changes in heart hypertrophy: significance during hypoxia-reoxygenation injury. Can J Physiol Pharmacol 70: 1330-1335, 1992[ISI][Medline].

61.   Kobayashi, S, Miescher E, and Clemens MG. A synergistic effect of extracellular hypocalcemic condition for hyperoxic reoxygenation injury in rat hepatocytes. Transplantation 67: 451-457, 1999[ISI][Medline].

62.   Kowaltowski, AJ, and Vercesi AE. Mitochondrial damage induced by conditions of oxidative stress. Free Radic Biol Med 26: 463-471, 1999[ISI][Medline].

63.   Krenitsky, TA, Tuttle JV, Cattau ED, Jr, and Wang P. A comparison of the distribution and electron acceptor specificities of xanthine oxidase and aldehyde oxidase. Comp Biochem Physiol B Biochem Mol Biol 49: 687-703, 1974.

64.   Kupatt, C, Weber C, Wolf DA, Becker BF, Smith TW, and Kelly RA. Nitric oxide attenuates reoxygenation-induced ICAM-1 expression in coronary microvascular endothelium: Role of NF-kappa B. J Mol Cell Cardiol 29: 2599-2609, 1997[ISI][Medline].

65.   Laderoute, KR, and Webster KA. Hypoxia/reoxygenation stimulated Jun kinase activity through redox signaling in cardiac myocytes. Circ Res 80: 336-344, 1997[Abstract/Free Full Text].

66.   Leo, R, Pratico D, Iuliano L, Pulcinelli FM, Ghiselli A, Pignatelli P, Colavita AR, FitzGerald GA, and Violi F. Platelet activation by superoxide anion and hydroxyl radicals intrinsically generated by platelets that had undergone anoxia and then reoxygenation. Circulation 95: 885-891, 1997[Abstract/Free Full Text].

67.   Levinson, RM, Shure D, and Moser KM. Reperfusion pulmonary edema after pulmonary artery thromboendarterectory. Am Rev Respir Dis 134: 1241-1245, 1986[ISI][Medline].

68.   Levy, R, Smith SD, Chandler K, Sadovsky Y, and Nelson DM. Apoptosis in human cultured trophoblasts is enhanced by hypoxia and diminished by epidermal growth factor. Am J Physiol Cell Physiol 278: C982-C988, 2000[Abstract/Free Full Text].

69.  Li C, Wright M, and Jackson R. Reactive species-mediated lung epithelial cell death after hypoxia and reoxygenation. Exp Lung Res. In press.

70.   Li, D, Tomson K, Yang B, Mehta P, Croker BP, and Mehta JL. Modulation of constitutive nitric oxide synthase, bcl-2 and Fas expression in cultured human coronary endothelial cells exposed to anoxia-reoxygenation and angiotensin II: role of AT1 receptor activation. Cardiovasc Res 41: 109-115, 1999[ISI][Medline].

71.   Lum, H, Barr DA, Shaffer JR, Gordon RJ, Ezrin AM, and Malik AB. Reoxygenation of endothelial cells increases permeability by oxidant-dependent mechanisms. Circ Res 70: 991-998, 1992[Abstract].

72.   Macmillan-Crow, LA, Crow JP, Kerby JD, Beckman JS, and Thompson JA. Nitration and inactivation of manganese superoxide dismutase in chronic rejection of human renal allografts. Proc Natl Acad Sci USA 93: 11853-11858, 1996[Abstract/Free Full Text].

73.   Mairbaurl, H, Wodopia R, Eckes S, Schulz S, and Bartsch P. Impairment of cation transport in A549 cells and rat alveolar epithelial cells by hypoxia. Am J Physiol Lung Cell Mol Physiol 273: L797-L806, 1997[ISI][Medline].

74.   Matalon, S, Benos D, and Jackson RM. Biophysical and molecular properties of amiloride-inhibitable Na+ channels in alveolar epithelial cells. Am J Physiol Lung Cell Mol Physiol 271: L1-L22, 1996[Abstract/Free Full Text].

75.   Matsushita, H, Morishita R, Nata T, Aoki M, Nakagami H, Taniyama Y, Yamamoto K, Higaki J, Yasufumi K, and Ogihara T. Hypoxia-induced endothelial apoptosis through nuclear factor-kappa B (NF-kappa B)-mediated bcl-2 suppression: in vivo evidence of the importance of NF-kappa B in endothelial cell regulation. Circ Res 86: 974-981, 2000[Abstract/Free Full Text].

76.   McCord, JM. Oxygen-derived free radicals in postischemic tissue injury. N Engl J Med 312: 159-163, 1985[Abstract].

77.   McGowan, AJ, Ruiz-Ruiz MC, Gorman AM, Lopez-Rivas A, and Cotter TG. Reactive oxygen intermediate(s) (ROI): common mediator(s) of poly(ADP-ribose) polymerase (PARP) cleavage and apoptosis. FEBS Lett 392: 299-303, 1996[ISI][Medline].

78.   McLeod, L, and Alayash AI. Detection of a ferrylhemoglobin intermediate in an endothelial cell model after hypoxia-reoxygenation. Am J Physiol Heart Circ Physiol 277: H02-H99, 1999.

79.   Mizukami, Y, Kobayashi S, Uberall F, Hellbert K, Kobayashi N, and Yoshida KI. Nuclear mitogen-activated protein kinase activation by protein kinase Czeta during reoxygenation after ischemic hypoxia. J Biol Chem 275: 19921-19927, 2000[Abstract/Free Full Text].

80.   Mizukami, Y, Yoshioka K, Morimoto S, and Yoshida KI. A novel mechanism of JNK1 activation. J Biol Chem 272: 16657-16662, 1997[Abstract/Free Full Text].

81.   Murphy, BJ, Robin ED, Tapper DP, Wong RJ, and Clayton DA. Hypoxic coordinate regulation of mitochondrial enzymes in mammalian cells. Science 223: 707-709, 1984[ISI][Medline].

82.   Muxfelt, M, and Schaper W. The activity of xanthine oxidase in heart of pigs, guinea pigs, rabbits, rats, and humans. Basic Res Cardiol 82: 486-492, 1987[ISI][Medline].

83.   Nakano, M, Mann DL, and Knowlton AA. Blocking the endogenous increase in HSP 72 increases susceptibility to hypoxia and reoxygenation in isolated adult feline cardiocytes. Circulation 95: 1523-1531, 1997[Abstract/Free Full Text].

84.   Nemoto, S, Takeda K, Yu ZX, Ferrans VJ, and Finkel T. Role of mitochondrial oxidants as regulators of cellular metabolism. Mol Cell Biol 20: 7311-7314, 2000[Abstract/Free Full Text].

85.   Ohman, T, Parish G, and Jackson RM. Hypoxic modulation of manganese superoxide dismutase promoter activity and gene expression in lung epithelial cells. Am J Respir Cell Mol Biol 21: 119-127, 1999[Abstract/Free Full Text].

86.   Ozaki, M, Deshpande SS, Angkeow P, Bellan J, Lowenstein CJ, Dinauer MC, Goldschmidt-Clermont PJ, and Irani K. Inhibition of the Rac1 GTPase protects against nonlethal ischemia/reperfusion-induced necrosis and apoptosis in vivo. FASEB J 14: 418-429, 2000[Abstract/Free Full Text].

87.   Ozaki, M, Deshpande SS, Angkeow P, Suzuki S, and Irani K. Rac1 regulates stress-induced, redox-dependent heat shock factor activation. J Biol Chem 275: 35377-35383, 2000[Abstract/Free Full Text].

88.   Paller, MS. Lateral diffusion of lipids in renal cells: effects of hypoxia and reoxygenation and role of cytoskeleton. Am J Physiol Cell Physiol 264: C201-C208, 1993[Abstract/Free Full Text].

89.   Paller, MS, and Neumann TV. Reactive oxygen species and rat renal epithelial cells during hypoxia and reoxygenation. Kidney Int 40: 1041-1049, 1991[ISI][Medline].

90.   Park, JH, Okayama N, Gute D, Krsmanovic A, Battarbee H, and Alexander JS. Hypoxia/aglycemia increases endothelial permeability: role of second messengers and cytoskeleton. Am J Physiol Cell Physiol 277: C1066-C1074, 1999[Abstract/Free Full Text].

91.   Parks, DA, and Granger DN. Xanthine oxidase: biochemistry, distribution and physiology. Acta Physiol Scand 548, Suppl: 87-99, 1986.

92.   Pastorino, JG, Snyder JW, Hoek JB, and Farber JL. Ca2+ depletion prevents anoxic death of hepatocytes by inhibiting mitochondrial permeability transition. Am J Physiol Cell Physiol 268: C676-C685, 1995[Abstract/Free Full Text].

93.   Pitkanen, S, and Robinson BH. Mitochondrial complex I deficiency leads to increased production of superoxide radicals and induction of superoxide dismutase. J Clin Invest 98: 345-351, 1996[Abstract/Free Full Text].

94.   Plateel, M, Dehouck MP, Torpier G, Cecchelli R, and Teissier E. Hypoxia increases the susceptibility to oxidant stress and the permeability of the blood-brain barrier endothelial cell monolayer. J Neurochem 65: 2138-2145, 1995[ISI][Medline].

95.   Poderoso, JJ, Lisedero C, Schopfer F, Riobo N, Carreras MC, Cadenas E, and Boveris A. The regulation of mitochondrial oxygen uptake by redox reactions involving nitric oxide and ubiquinol. J Biol Chem 274: 37709-33716, 1999[Abstract/Free Full Text].

96.   Poderoso, JJ, Peralta JG, Lisdero CL, Carreras MC, Radisic M, Schopfer F, Cadenas E, and Boveris A. Nitric oxide regulates uptake and hydrogen peroxide release by the isolated beating rat heart. Am J Physiol Cell Physiol 274: C112-C119, 1998[Abstract/Free Full Text].

97.   Raha, S, McEachern GE, Myint AT, and Robinson BH. Superoxides from mitochondrial complex III: the role of manganese superoxide dismutase. Free Radic Biol Med 29: 170-180, 2000[ISI][Medline].

98.   Rakhit, RD, Edwards RJ, Mockridge JW, Baydoun AR, Wyatt AW, Mann GE, and Marber MS. Nitric oxide-induced cardioprotection in cultured rat ventricular myocytes. Am J Physiol Heart Circ Physiol 278: H1211-H1217, 2000[Abstract/Free Full Text].

99.   Ratych, RE, Chuknyiska RS, and Bulkley GB. The primary localization of free radical generation after anoxia/reoxygenation in isolated endothelial cells. Surgery 102: 122-131, 1987[ISI][Medline].

100.   Reinheckel, T, Korn S, Mohring S, Augustin W, Halangk W, and Schild L. Adaptation of protein carbonyl detection to the requirements of proteome analysis demonstrated for hypoxia/reoxygenation in isolated rat liver mitochondria. Arch Biochem Biophys 376: 59-65, 2000[ISI][Medline].

101.   Robin, ED, Murphy BJ, and Theodore J. Coordinate regulation of glycolysis by hypoxia in mammalian cells. J Cell Physiol 118: 287-290, 1984[ISI][Medline].

102.   Robin, ED, and Theodore J. Are there ischemic lung diseases? Arch Intern Med 142: 1791-1792, 1982[ISI][Medline].

103.   Russell, W, Matalon S, and Jackson R. MnSOD expression in alveolar epithelial cells from hypoxic and hypoperfused lungs. Am J Respir Cell Mol Biol 11: 366-371, 1994[Abstract].

104.   Saikumar, P, Dong Z, Patel Y, Hall K, Hopfer U, Weinberg JM, and Venkatachalam MA. Role of hypoxia-induced bax translocation and cytochrome c release in reoxygenation. Oncogene 17: 3401-3415, 1998[ISI][Medline].

105.   Saikumar, P, Dong Z, Weinberg JM, and Venkatachalam MA. Mechanisms of cell death in hypoxia/reoxygenation injury. Oncogene 17: 3341-3349, 1998[ISI][Medline].

106.   Sakamoto, K, Urushidani T, and Nagao T. Translocation of HSP27 to cytoskeleton by repetitive hypoxia-reoxygenation in the rat myoblast cell line, H9c2. Biochem Biophys Res Commun 251: 576-579, 1998[ISI][Medline].

107.   Schafer, C, Ladilov YV, Siegmund B, and Piper HM. Importance of bicarbonate transport for protection of cardiomyocytes against reoxygenation injury. Am J Physiol Heart Circ Physiol 278: H1457-H1463, 2000[Abstract/Free Full Text].

108.   Schinetti, ML, Sbarbati R, and Scarlattini M. Superoxide production by human umbilical vein endothelial cells in an anoxia-reoxygenation model. Cardiovasc Res 23: 76-80, 1989[ISI][Medline].

109.   Semenza, GL. HIF-1: mediator of physiological and pathophysiological responses to hypoxia. J Appl Physiol 88: 1474-1480, 2000[Abstract/Free Full Text].

110.   Semenza, GL. Cellular and molecular dissection of reperfusion injury ROS within and without. Circ Res 86: 117-118, 2000[Free Full Text].

111.   Sharpe, MA, and Cooper CE. Interaction of peroxynitrite with mitochondrial cytochrome oxidase. J Biol Chem 273: 30961-30972, 1998[Abstract/Free Full Text].

112.   Shlafer, M, Myers CL, and Adkins S. Mitochondrial hydrogen peroxide generation and activities of glutathione peroxidase and superoxide dismutase following global ischemia. J Mol Cell Cardiol 19: 1195-1206, 1987[ISI][Medline].

113.   Simon, LM, Robin ED, Phillips JR, Acevedo J, Axline SG, and Theodore J. Enzymatic basis for bioenergetic differences of alveolar versus peritoneal macrophages and enzyme regulation by molecular O2. J Clin Invest 59: 443-448, 1977[ISI][Medline].

114.   Soboll, S, Brdiczka D, Jahnke D, Schmidt A, Schlattner U, Wendt S, Wyss M, and Wallimann T. Octomer-dimer transitions of mitochondrial creatine kinase in heart disease. J Mol Cell Cardiol 31: 857-866, 1999[ISI][Medline].

115.   Suzuki, K, Sawa Y, Kaneda Y, Ichikawa H, Shirakura R, and Matsuda H. Over expressed heat shock protein 70 attenuates hypoxic injury in coronary endothelial cells. J Mol Cell Cardiol 30: 1129-1136, 1998[ISI][Medline].

116.   Tagami, M, Yamagata K, Ikeda K, Nara Y, Fujino H, Kubota A, Numano F, and Yamori Y. Vitamin E prevents apoptosis in cortical neurons during hypoxia and oxygen reperfusion. Lab Invest 78: 1415-1429, 1998[ISI][Medline].

117.   Tan, S, Zhou F, Nielsen VG, Wang Z, Gladson CL, and Parks DA. Increased injury following intermittent fetal hypoxia-reoxygenation is associated with increased free radical production in fetal rabbit brain. J Neuropathol Exp Neurol 58: 972-981, 1999[ISI][Medline].

118.   Terada, LS. Hypoxia-reoxygenation increases O2 efflux, which injures endothelial cells by an extracellular mechanism. Am J Physiol Heart Circ Physiol 270: H945-H950, 1996[Abstract/Free Full Text].

119.   Thannickal, VJ, and Fanburg BL. Reactive oxygen species in cell signaling. Am J Physiol Lung Cell Mol Physiol 279: L1005-L1028, 2000[Abstract/Free Full Text].

120.   Tong, WH, and Rouault T. Distinct iron-sulfur cluster assembly complexes exist in the cytosol and mitochondria of human cells. EMBO J 19: 5692-5700, 2000[Abstract/Free Full Text].

121.   Valdez, LB, Alvarez S, Arnaz SL, Schopfer F, Carreras MC, Poderoso JJ, and Boveris A. Reactions of peroxynitrite in the mitochondrial matrix. Free Radic Biol Med 29: 349-356, 2000[ISI][Medline].

122.   Van der Vliet, A, Eiserich JP, Halliwell B, and Cross CE. Formation of reactive nitrogen species during peroxidase-catalyzed oxidation of nitrite. J Biol Chem 272: 7617-7625, 1997[Abstract/Free Full Text].

123.   Vogt, M, Bauer MKA, Ferrari D, and Schulze-Osthoff K. Oxidative stress and hypoxia/reoxygenation trigger CD95 (APO-1/Fas) ligand expression in microglial cells. FEBS Lett 429: 67-72, 1998[ISI][Medline].

124.   Waleh, NS, Calaoagan J, Murphy BJ, Knapp AM, Sutherland RM, and Laderoute KR. The redox-sensitive human antioxidant responsive element induces gene expression under low oxygen conditions. Carcinogenesis 19: 1333-1337, 1998[Abstract].

125.   Wang, GL, Jiang BH, Rue EA, and Semenza GL. Hypoxia inducible factor 1 is a basic-helix-loop-helix-PAS heterodimer regulated by cellular O2 tension. Proc Natl Acad Sci USA 92: 5510-5514, 1995[Abstract].

126.   Watkins, MT, Al-Badawi H, Cardenas R, Dubois E, and Larson DM. Endogenous reactive oxygen metabolites mediate sublethal endothelial cell dysfunction during reoxygenation. J Vasc Surg 23: 95-103, 1996[ISI][Medline].

127.   Watkins, MT, Haudenschild CC, Al-Badawi H, Velazquez FR, and Larson DM. Immediate responses of endothelial cells to hypoxia and reoxygenation: An in vitro model of cellular dysfunction. Am J Physiol Heart Circ Physiol 268: H749-H758, 1995[Abstract/Free Full Text].

128.   Webster, KA, Disher DJ, Kaiser S, Hernandez O, Sato B, and Bishopric NH. Hypoxia-activated apoptosis of cardiac myocytes requires reoxygenation or a pH shift and is independent of p53. J Clin Invest 104: 239-252, 1999[Abstract/Free Full Text].

129.   Wei, Z, Costa K, Al-Mehdi AB, Dodia C, Muzyantov V, and Fisher AB. Simulated ischemia in flow-adapted endothelial cells leads to generation of reactive oxygen species in cell signaling. Circ Res 85: 682-689, 1999[Abstract/Free Full Text].

130.   Weinberg, JM, Venkatachalam MA, Roeser NF, and Nissim I. Mitochondrial dysfunction during hypoxia/reoxygenation and its correction by anaerobic metabolism of citric acid cycle intermediates. Proc Natl Acad Sci USA 97: 2826-2831, 2000[Abstract/Free Full Text].

131.   Weinberg, JM, Venkatachalam MA, Roeser NF, Saikumar P, Dong Z, Senter RA, and Nissim I. Anaerobic and aerobic pathways for salvage of proximal tubules from hypoxia-induced mitochondrial injury. Am J Physiol Renal Physiol 279: F927-F943, 2000[Abstract/Free Full Text].

132.   Whitmarsh, AJ, and Davis RJ. Transcription factor AP-1 regulation by mitogen-activated protein kinase signal transduction pathways. J Mol Med 74: 589-607, 1996[ISI][Medline].

133.   Yao, Z, Tong J, Tan X, Li CQ, Shao A, Kin WC., Vanden Hoek TL, Becker LB, Head CA, and Schumacker PT. Role of reactive oxygen species in acetylcholine-induced preconditioning in cardiomyocytes. Am J Physiol Heart Circ Physiol 277: H2504-H2509, 1999[Abstract/Free Full Text].

134.   Zou, MH, and Bachschmid M. Hypoxia-reoxygenation triggers coronary vasospasm in isolated bovine coronary arteries via tyrosine nitration of prostacyclin synthase. J Exp Med 190: 135-139, 1999[Abstract/Free Full Text].

135.   Zweier, JL, Kuppusamy P, Thompson-Gorman S, Klunk D, and Lutty GA. Measurement and characterization of free radical generation in reoxygenated human endothelial cells. Am J Physiol Cell Physiol 266: C700-C708, 1994[Abstract/Free Full Text].


Am J Physiol Cell Physiol 282(2):C227-C241