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Biology of Ageing Laboratories, University of Geneva, Geneva, Switzerland
ABSTRACT I. INTRODUCTION A. Reactive Oxygen Species B. Physiological Sources of ROS C. ROS-Generating NADPH Oxidase: A Historical Overview II. THE NOX FAMILY OF NADPH OXIDASE: INTRODUCING THE PLAYERS A. The NOX Family Members 1. NOX2: prototype NOX 2. NOX1 3. NOX3 A) P22PHOX. B) ABSENCE OF CYTOSOLIC SUBUNITS. C) NOXO1. D) NOXA1. E) P47PHOX AND P67PHOX. F) RAC. 4. NOX4 A) P22PHOX. B) NOX4 DOES NOT REQUIRE CYTOSOLIC SUBUNITS. C) RAC. 5. NOX5 6. DUOX1 and DUOX2 B. NOX Subunits and Regulatory Proteins 1. p22phox 2. Organizer subunits: NOXO1, p47phox 3. Activator subunits: p67phox and NOXA1 4. p40phox 5. Rac GTPases C. NOX Inhibitors 1. Diphenylene iodonium 2. Apocynin 3. AEBSF 4. Neopterin 5. gp91ds-tat D. Polymorphisms in NOX Enzymes and Subunits 1. NOX1–5 and DUOX 2. Subunits and regulatory proteins III. PHYSIOLOGICAL FUNCTION OF ROS AND NOX FAMILY NADPH OXIDASES A. Host Defense and Inflammation 1. ROS-dependent killing A) SUPEROXIDE. B) HYDROGEN PEROXIDE AND PEROXIDASE. C) RNS. D) OTHER ROS. 2. Inactivation of microbial virulence factors 3. Regulation of pH and ion concentration in the phagosome A) PHAGOSOMAL PH. B) PHAGOSOMAL ION CONCENTRATION. 4. NOX enzymes and proton channels 5. Anti-inflammatory activity B. Cellular Signaling 1. Inhibition of phosphatases 2. Activation of kinases 3. Regulation of ion channels 4. Ca2+ signaling A) PLASMA MEMBRANE CALCIUM CHANNELS. B) CALCIUM RELEASE FROM INTRACELLULAR STORES. C) CALCIUM PUMPS. C. Gene Expression D. Cellular Death and Cellular Senescence E. Regulation of Cell Growth 1. Cellular senescence 2. Cellular growth F. Oxygen Sensing 1. Kidney 2. Carotid body 3. Pulmonary oxygen sensing 4. Others G. Biosynthesis and Protein Cross-Linking H. Regulation of Cellular Redox Potential I. Reduction of Metal Ions J. Regulation of Matrix Metalloproteinases K. Angiogenesis L. Cross-Talk With the Nitric Oxide System IV. NADPH OXIDASES IN SPECIFIC ORGAN SYSTEMS: PHYSIOLOGY AND PATHOPHYSIOLOGY A. Adipose Tissue B. Biology of Reproduction 1. Testis, spermatocytes, and fertilization 2. Prostate 3. Ovary 4. Uterus, placenta, and preeclampsia A) UTERUS. B) PLACENTA. C) PREECLAMPSIA. C. Cardiovascular System 1. Vascular system 2. Heart A) MYOCARDIAL INFARCTION AND MYOCARDIAL REPERFUSION INJURY. B) ISCHEMIC PRECONDITIONING. C) CARDIAC HYPERTROPHY, FIBROSIS, AND HEART FAILURE. D) ATRIAL FIBRILLATION. 3. Shock and related pathologies D. Central Nervous System 1. Microglia 2. Oligodendrocytes 3. Astrocytes 4. Neurons 5. NOX enzymes in pathologies of the CNS A) ISCHEMIC STROKE. B) ALZHEIMER'S DISEASE, PARKINSON'S DISEASE, AND HIV DEMENTIA. E. Endocrinology 1. Thyroid 2. Endocrine pancreas F. Gastrointestinal System and Liver 1. Stomach and Helicobacter 2. Colon 3. Liver A) HEPATOCYTES. B) HEPATIC STELLATE CELLS. C) KUPFFER CELLS. D) LIVER CIRRHOSIS AND ALCOHOLIC LIVER DISEASE. E) HEPATIC CANCER. F) LIVER ISCHEMIA AND REPERFUSION INJURY. G. Kidney and Urinary Tract 1. ROS and NOX in kidney physiology A) RENAL BLOOD FLOW. B) CELL FATE. C) GENE EXPRESSION. 2. ROS and NOX in kidney pathophysiology A) DIABETIC NEPHROPATHY. 3. High salt and hypertension 4. Others H. Lung and Airways 1. Airway epithelium 2. Alveolar cells 3. Lung vasculature and pulmonary hypertension 4. Pulmonary fibroblasts and pulmonary fibrosis 5. Asthma and chronic obstructive pulmonary disease I. Musculoskeletal System 1. Bone A) OSTEOCLASTS. B) OSTEOBLASTS. C) CARTILAGE. 2. Skeletal muscle J. Platelets and Leukocytes 1. Hematopoietic stem cells 2. Granulocytes 3. Macrophages 4. Dendritic cells 5. B lymphocytes 6. T lymphocytes 7. Platelets K. Sensory Organs 1. Eye A) LENS. B) RETINA. 2. Inner ear L. Skin 1. Keratinocytes, fibroblasts, and melanocytes 2. ROS and wound healing V. CONCLUSIONS GRANTS ACKNOWLEDGMENTS REFERENCES
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I. INTRODUCTION |
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Reactive oxygen species (ROS) are oxygen-derived small molecules, including oxygen radicals [superoxide (O2
–), hydroxyl (OH), peroxyl (RO2), and alkoxyl (RO·)] and certain nonradicals that are either oxidizing agents and/or are easily converted into radicals, such as hypochlorous acid (HOCl), ozone (O3), singlet oxygen (1O2), and hydrogen peroxide (H2O2). Nitrogen-containing oxidants, such as nitric oxide, are called reactive nitrogen species (RNS). ROS generation is generally a cascade of reactions that starts with the production of superoxide. Superoxide rapidly dismutates to hydrogen peroxide either spontaneously, particularly at low pH or catalyzed by superoxide dismutase. Other elements in the cascade of ROS generation include the reaction of superoxide with nitric oxide to form peroxynitrite, the peroxidase-catalyzed formation of hypochlorous acid from hydrogen peroxide, and the iron-catalyzed Fenton reaction leading to the generation of hydroxyl radical (468, 874).
ROS avidly interact with a large number of molecules including other small inorganic molecules as well as proteins, lipids, carbohydrates, and nucleic acids. Through such interactions, ROS may irreversibly destroy or alter the function of the target molecule. Consequently, ROS have been increasingly identified as major contributors to damage in biological organisms. In 1956, Harmann made his ground-breaking observations on the role of ROS in the aging process (350), and the concept of ROS as agents of cellular damage became widely accepted in theories of aging (73). Yet, at least one beneficial function of ROS production was also realized quite early, namely, the importance of ROS in host defense. This point became particularly clear when the link was made between deficiency in ROS generation and reduced killing ability in leukocytes. However, over the last decades, a second important concept of ROS has been evolving. In fact, ROS are involved not only in cellular damage and killing of pathogens, but also in a large number of reversible regulatory processes in virtually all cells and tissues. This review discusses both the physiological and pathophysiological role of ROS generated by the NADPH oxidase family of enzymes.
B. Physiological Sources of ROS
The physiological generation of ROS can occur as a byproduct of other biological reactions. ROS generation as a byproduct occurs with mitochondria, peroxisomes, cytochrome P-450, and other cellular elements (50, 307, 314, 356, 588, 636, 715, 791, 874). However, the phagocyte NADPH oxidase was the first identified example of a system that generates ROS not as a byproduct, but rather as the primary function of the enzyme system. The discovery of other members of the NOX family of NADPH oxidases demonstrated that enzymes with the primary function of ROS generation are not limited to phagocytes. In fact, the ROS-generating enzymes described in this review are found in virtually every tissue. This review focuses on novel homologs of the phagocyte NADPH oxidase. A complete coverage of the biochemistry and physiology of the phagocyte NADPH oxidase itself is beyond the scope of this review. However, we give an overview of the phagocyte NADPH oxidase to serve as a framework to understand the specific features of novel NOX isoforms (see sect. IIA1).
C. ROS-Generating NADPH Oxidase: A Historical Overview
Although the NADPH oxidase was not yet identified, a respiratory burst by cells had already been described by the first half of the 20th century. These early observations were done in sea urchin eggs (938), phagocytes in 1933 (51), and spermatocytes in 1943 (565). In 1959, Sbarra and Karnovsky (787) demonstrated that the phagocyte respiratory burst was an energy-requiring process that depended on glucose metabolism. Shortly after, in 1961, Iyer et al. (419) showed that the phagocyte respiratory burst results in the generation of hydrogen peroxide. There was a major controversy over whether the main substrate for the enzyme system was NADPH or NADH. In 1964, Rossi and Zatti (755) correctly proposed that an NADPH oxidase was responsible for the respiratory burst. In 1970, Klebanoff (466) demonstrated a contribution of myeloperoxidase to the respiratory burst-dependent antimicrobial activity of phagocytes. In 1973, Babior et al. (43) reported that the initial product of the respiratory burst oxidase was superoxide and not hydrogen peroxide.
A second important line of study that led to the discovery of the phagocyte NADPH oxidase came from clinical research. In 1957, Berendes et al. (83) recognized a new and relatively rare syndrome in young boys who suffered from recurrent pyogenic infections that was accompanied with granulomatous reaction, lymphadenopathy, and hypergammaglobulinemia. The genetic disorder is now referred to as chronic granulomatous disease (CGD). Quie et al. (721) showed that CGD phagocytes have diminished bactericidal capacity, although many phagocyte functions, such as chemotaxis, phagocytosis, and degranulation, were found to be intact in CGD phagocytes. In 1967, it was recognized that the respiratory burst was absent in the phagocytes of CGD patients (47, 385, 721).
Further characterization of ROS generation by phagocytes revealed that this enzyme system 1) produced superoxide and its downstream metabolite hydrogen peroxide; 2) was insensitive to cyanide, distinguishing it from mitochondria and myeloperoxidase (MPO); 3) was present in phagocytes from MPO-deficient patients, but absent in those of CGD patients; and 4) was selective for NADPH over NADH by a factor of 100 (42).
The identification of proteins responsible for ROS production in phagocytes was the next challenge. A breakthrough occurred in 1978, when Segal, Jones, and colleagues (798, 799) identified cytochrome b558, which was missing in the leukocytes of many CGD patients. In the late 1980s, the gene coding for the catalytic subunit of the phagocyte NADPH oxidase, commonly referred to as gp91phox, was cloned by Royer-Pokora et al. (762) and Teahan et al. (865). In the novel NOX terminology, gp91phox is called NOX2.
However, it was rapidly understood that NOX2 was not the only component of the phagocyte enzyme. In 1987, the transmembrane protein p22phox was discovered as the membrane subunit associated with NOX2 (216, 693, 795). The development of a cell-free system allowed activation of the phagocyte NADPH oxidase using purified cytosol and membrane fractions (110, 373). This system provided the tools to discover the cytosolic subunits p47phox and p67phox (660, 923) and to define the roles of the small GTP-binding proteins Rac1 and Rac2 (4, 470). In 1993, Wientjes et al. (951) described a third cytosolic subunit, p40phox.
In parallel with the progress toward understanding the phagocyte NADPH oxidase, a series of observations suggested that enzyme systems similar to the phagocyte NADPH oxidase exist in many other cell types, including fibroblasts (600), various tumor cells (855), and vascular smooth muscle (321). Fibroblasts from NOX2-deficient patients had normal ROS generation, suggesting that the phagocyte NADPH oxidase was not the source of ROS in fibroblasts (249). However, the molecular identity of the NADPH oxidase-like systems in nonphagocytic cells remained obscure. That was fundamentally changed by the availability of the human genome sequence. Two groups independently identified a first homolog of NOX2, which was initially referred to as mitogenic oxidase 1 (mox-1; Ref. 841) or NADPH oxidase homolog 1 (NOH-1; Ref. 55); this isoform has been named NOX1 in the novel terminology.
The identification of NOX1 was quickly followed by the cloning of NOX3 (143, 454), NOX4 (294, 813), and NOX5 (56, 143). In parallel with the identification of NOX1 to NOX5, two very large members of the NOX family were discovered, namely, DUOX1 and DUOX2, initially also referred to as thyroid oxidases (189, 228).
The identification of the new NOX/DUOX proteins was not, however, always followed by an immediate demonstration of their biochemical function. Indeed, the closest NOX2 homolog, NOX1, is usually inactive when transfected by itself. A search for homologs of the cytosolic subunits of the phagocyte NADPH oxidase (p47phox and p67phox) led to the cloning of a novel set of cytosolic subunits, NOXO1 and NOXA1 (53, 297, 857). Similarly, heterologous expression of DUOX enzymes is only successfully achieved since the identification of the DUOX maturation factors DUOXA1 and DUOXA2 (319).
Is the NOX family complete? Given the essentially complete databases of several mammalian genomes, it appears likely that most specific elements of the NOX system have now been identified: seven NOX isoforms, two organizer subunits (p47phox, NOXO1), two activator subunits (p67phox, NOXA1), and two DUOX-specific maturation factors (DUOXA1 and DUOXA2). There is, however, some space for continuing gene discovery: nonessential, modulatory subunits such as p40phox might have gone unrecognized, and the possible existence of an unidentified p22phox homolog has been suggested (844).
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II. THE NOX FAMILY OF NADPH OXIDASE: INTRODUCING THE PLAYERS |
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The description of NOX family members in this review focuses on mammalian NOX homologs and subunits (Table 1). Some features that are invariable for mammalian NOX enzymes (e.g., electron transfer to oxygen) might be different in nonmammalian organisms (e.g., ferric reductases in yeast).
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NOX2, also known as gp91phox, is the prototype NADPH oxidase. Its biochemical features have been extensively studied and abundantly reviewed in the recent past (44, 175, 652, 750, 917). Thus an extended coverage of NOX2 goes beyond the scope of this review. However, we summarize the most important features of NOX2, with a particular focus on the properties of NOX2 that allow a better understanding of other NOX isoforms.
Much of what is known about the topography and structure of the NOX isoforms is derived from studies on NOX2. Yet, even for NOX2, the suggested topographical features are deduced from indirect data and hence are putative assignments. Definitive assignments will have to await crystallographic studies, which unfortunately have not been achieved yet. Still, it is likely that the basic features, outlined in Figure 1, are correct. Hydropathy plots predict between four and six transmembrane domains for NOX2 (365, 800). Phage display library screening provide experimental data defining the extracellular domains (118, 406, 644). Antibody mapping studies demonstrate a cytoplasmic localization of the COOH terminus (119, 406, 757). Sequencing data and antibody mapping confirm a cytoplasmic NH2 terminus (676, 865), over the alternative suggestion that the NH2 terminus is proteolytically cleaved (143). Taken together, the available data suggest that NOX2 has six transmembrane domains and that its COOH terminus and its NH2 terminus are facing the cytoplasm.
Human NOX2 is a highly glycosylated protein that appears as a broad smear on SDS-PAGE reflecting the heterogeneity of glycosylation. The fully glycosylated form runs with an apparent molecular mass of 
70–90 kDa. Removal of the carbohydrates by endoglycosidase F leaves a protein that runs at 55 kDa, demonstrating the extent of glycosylation (351). The carbohydrate chains are composed of N-acetylglucosamine and galactose and, to a lesser extent, frucose, mannose, and glucose (351). A mutagenesis approach demonstrates that the carbohydrates are bound to asparagine residues (132Asn, 149Asn, and 240Asn) in the second and third predicted extracellular loops (929).
The activation of NOX2 occurs through a complex series of protein/protein interactions (Fig. 2; for more detailed recent reviews, see Refs. 328, 652, 844). NOX2 constitutively associates with p22phox. Indeed, the NOX2 protein is unstable in the absence of p22phox, and phagocytes from p22phox-deficient patients have no detectable NOX2 protein (217, 692, 828). Activation of NOX2 requires translocation of cytosolic factors to the NOX2/p22phox complex (Fig. 3). The present working model is as follows. Phosphorylation of p47phox leads to a conformational change allowing its interaction with p22phox (327, 843). It is thought that p47phox organizes the translocation of other cytosolic factors, hence its designation as "organizer subunit." The localization of p47phox to the membrane brings the "activator subunit" p67phox into contact with NOX2 (342) and also brings the small subunit p40phox to the complex. Finally, the GTPase Rac interacts with NOX2 via a two-step mechanism involving an initial direct interaction with NOX2 (214), followed by a subsequent interaction with p67phox (476, 508). Once assembled, the complex is active and generates superoxide by transferring an electron from NADPH in the cytosol to oxygen on the luminal or extracellular space.
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In the first step, electrons are transferred from NADPH to FAD, a process that is regulated by the activation domain of p67phox (658). NOX2 is selective for NADPH over NADH as a substrate, with Km values of 40–45 µM versus 2.5 mM, respectively (160). In the second step, a single electron is transferred from the reduced flavin FADH2 to the iron center of the inner heme. Since the iron of the heme can only accept one electron, the inner heme must donate its electron to the outer heme before the second electron can be accepted from the now partially reduced flavin, FADH. The force for the transfer of the second electron, while smaller (31 vs. 79 mV), is still energetically favorable. However, the transfer of the electron from the inner heme to the outer heme is actually against the electromotive force between these two groups. To create an energetically favorable state, oxygen must be bound to the outer heme to accept the electron (175, 223, 917).
NOX2 was first described in neutrophils and macrophages and is often referred to as the phagocyte NADPH oxidase. NOX2 is still widely considered to have a very limited, essentially phagocyte-specific tissue expression (e.g., Ref. 844), yet when tissue distribution of total mRNA from various organs is investigated, NOX2 appears to be among the most widely distributed among the NOX isoforms (Table 2). It is described in a large number of tissues, including thymus, small intestine, colon, spleen, pancreas, ovary, placenta, prostate, and testis (143). Mostly this wide tissue distribution is due to the presence of phagocytes and/or blood contamination in the tissues from which total mRNA has been extracted. However, there is now also increasing evidence at both the message and the protein level for expression of NOX2 in nonphagocytic cells, including neurons (806), cardiomyocytes (372), skeletal muscle myocytes (426), hepatocytes (739), endothelial cells (313, 434, 538), and hematopoietic stem cells (704).
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In cells other than phagocytes, the subcellular distribution varies depending on the specific cell type. In smooth muscle cells, NOX2 is found to colocalize with the perinuclear cytoskeleton (538). In hippocampal neurons, NOX2 is suggested to be localized in the membranes of synaptic sites (866), possibly playing a role in superoxide-dependent long-term potentiation and memory function.
The human and mouse NOX2 gene is located on the X chromosome. At the transcriptional level, a promoter region sufficient for NOX2 expression in monocytes lies in the 450-bp region before the transcriptional start site (818). However, additional regulatory regions located up to 60 kb upstream from the start site also influence expression, with sensitive regions at –13, –15, –28, and –29 kb (545). NOX2 gene expression is regulated by repressing and activating factors. Repressing factors include the CCAAT displacement protein (CDP) (563, 819), HOXA10, and Meis1 (76). Activating factors include PU.1, Elf-1, YY1, and interferon regulatory factors IRF1 and IRF2 (243, 422, 850, 924), NF
B (28), HOXA9 (76), and PB1 (76). The PU.1 transcription factor appears to have a particularly important role in the expression of NOX2, as PU.1-deficient mice fail to express NOX2 (850). Mutations in the NOX2 promoter region at positions –53, –52, or –50 interfere with PU.1 binding, and CGD patients with such mutations do not express NOX2 in neutrophils, monocytes, and B lymphocytes. However, the importance of the PU.1 promoter may be somewhat cell specific, as these patients do express NOX2 in eosinophils (850).
NOX2 gene expression is inducible. This has been demonstrated in phagocytes in response to interferon-
(mRNA) (654), in myofibroblasts after carotid artery injury (mRNA) (856), and in cardiomyocytes after acute myocardial infarction (protein) (488). NOX2 expression is also increased in response to angiotensin II in adipose tissue (mRNA) (359), aorta (mRNA) (359), heart (mRNA) (359), resistance artery vascular smooth muscle cells (mRNA and protein) (881), and pancreatic islets (protein) (648). Note however that an increase in NOX2 levels is not invariably due to transciptional activation. In the case of resistance arteries, the angiotensin II-induced NOX2 elevation may be due to increased de novo protein synthesis, regulated at a posttranscriptional level (881).
NOX1 was the first homolog of NOX2 (then called gp91phox) to be described (55, 841). NOX1 and NOX2 genes appear to be the result of a relatively recent gene duplication, as the number and the length of the exons is virtually identical between the two genes (53). Similarly, at the protein level, there is a high degree of sequence identity (60%) between NOX1 and NOX2 (55, 841). The human and mouse NOX1 gene is located on the X chromosome. An alternatively spliced form of NOX1 lacks exon 11 (55, 296, 353). It appears that this splice variant encodes a protein incapable of producing superoxide (296). The existence of a second very short isoform of NOX1 had been suggested; however, this turned out to be an artifact due to the formation of a stable loop in the NOX1 mRNA (see Refs. 55, 296, 353). In mouse, several other splice variants due to the use of alternative promoters have been described (29). Most studies suggest a molecular mass of NOX1 in the range of 55–60 kDa (24, 178, 425). If these values are correct, NOX1 is most likely not N-glycosylated, despite the presence of two NXT/S consensus glycosylation sites in the extracellular domains.
The message for NOX1 is most highly expressed in colon epithelium (53, 841, 854); however, it is also expressed in a variety of other cell types, including vascular smooth muscle cells (510, 841), endothelial cells (14, 473), uterus (55, 841), placenta (178), prostate (55, 841), osteoclasts (516), retinal pericytes (576), as well as in several cell lines, such as the colon tumor cell lines Caco-2 (159, 701), DLD-1 (701), and HT-29 (701) and the pulmonary epithelial cell line A549 (320). Expression of NOX1 in the gastric mucosa is species dependent. It was not found in human stomach (770) but is functionally expressed in guinea pig stomach pit cells and mucosal cells (444, 447, 868).
In addition to its constitutive expression in a variety of tissues, the NOX1 message is induced under many circumstances. In vascular smooth muscle, platelet-derived growth factor (PDGF), prostaglandin F2
, and angiotensin II induce NOX1 expression (441, 510, 841, 955). NOX1 is upregulated in restenosing carotid artery after injury (856) and in prostate in response to castration (859). Other conditions where NOX1 upregulation is observed include interferon- addition to Caco-2 colonocytes (295), Ras expression in rat kidney cells (614), BMP4 stimulation of endothelial cells (823), and Helicobacter pylori lipopolysaccharide stimulation of guinea pig gastric mucosal cells (443). Studies on the inducible expression of NOX1 in the vascular system suggest an involvement of epidermal growth factor (EGF) receptor transactivation and involvement of ATF-1, phosphatidylinositol 3-kinase, and protein kinase C (PKC)-
(252). In renal mesangial cells, nitric oxide downregulates NOX1 (mRNA and protein) (707).
The 5'-region of the human NOX1 gene contains binding elements for signal transducers and activators of transcription (STATs), interferon regulatory factor (IRF), AP-1, NFB, CREB, CBP/p300 elements (498), and GATA factors (107). Constitutive expression of NOX1 in intestinal epithelial cells depends on the GATA-binding sites (107), while interferon--enhanced expression is regulated by the binding of activated STAT1 dimers to the -activated sequence (GAS) element (498). Note that the GATA sites are within the 520-bp region upstream of the transcription initiation site, while the GAS elements is located at –3818 to –3810 bp (498). In the mouse NOX1 gene, additional promoters up to 110 kb upstream from the transcription initiation site give rise to NOX1 splice variants (29). Within the colon, there is a gradient of NOX1 expression with levels being low in the proximal and high in the distal colon (302, 854). However, at this point, it is not clear whether this gradient is constitutive or secondary to bacterial colonization.
Data on the subcellular localization of NOX1 are scarce, mainly because the generation of high quality antibodies against NOX1 (and other NOX isoforms) turned out to be a challenge and some of the antibodies used were not subjected to rigorous validation protocols. With this limitation in mind, there are several studies reporting a subcellular localization of NOX1; in keratinocytes, there was a weak cytoplasmic and a strong nuclear staining (134). One study in vascular smooth muscle suggests an ER pattern (425), while another describes punctate patches along cell surface membranes, possibly corresponding to a caveolar localization (378).
In studies using a cell-free system, NOX1 is selective for NADPH over NADH as a substrate (977).
After the initial discovery of NOX1, it was not immediately obvious whether NOX1 was indeed a superoxide generating enzyme. While one group reported a very low level of superoxide generation in NOX1-transfected cells without the need of a stimulus (841), other groups did not observe such ROS generation by NOX1 alone (53, 295, 297). The discovery of colon homologs of the cytosolic subunits of the phagocyte NADPH oxidase resolved the issue (53, 146, 297, 857): superoxide generation by NOX1 depends on cytosolic subunits. The novel cytosolic subunits were named NOXO1 (NOX organizer 1 = p47phox homolog) and NOXA1 (NOX activator 1 = p67phox homolog). Details concerning these proteins are discussed below. The discovery of the subunit dependence of NOX1 introduced new complexities. First, in transfected cells, NOX1 is also able to use the p47phox and p67phox subunits, suggesting that cytosolic subunits are not specific for a given NOX protein (53). For example, it might be possible that p47phox acts as a subunit of NOX1 in the vascular system (see below). Second, while expression systems using the mouse proteins suggest a constitutive activity of the NOX1/NOXO1/NOXA1 system, studies using human proteins show only a weak constitutive activity, and full activation depends on activation through the PKC activator phorbol 12-myristate 13-acetate (PMA) (297, 857). There are indeed significant differences between the mouse and the human proteins, in particular, in the region of the phox homology domain which is distinct in human NOXO1. Yet, at this point, it is not clear whether the difference in PKC dependence is really due to a difference between the mouse and human NOX1/NOXO1/NOXA1 system or whether this reflects some experimental details, such as cell lines or the use of transient versus stable expression systems.
In addition to its dependence on cytosolic subunits, NOX1 requires the membrane subunit p22phox (24, 446). The p22phox dependence of NOX1 might be less stringent than the one observed for NOX2 and NOX3 (844). There is now also ample evidence for an involvement of the small GTPase Rac in the regulation of NOX1 activity (144, 443, 615, 687, 844, 857, 892). Rac binds to the TPR domain of the activator subunit NOXA1 (144, 857, 892), but in analogy with NOX2, Rac activation of NOX1 might be a two-step process that also includes a direct binding to NOX1.
NOX3 was described in 2000 based on its sequence similarity to other NOX isoforms (454), although the first studies on the function of the protein did not appear until 2004 (54, 677). NOX3 shares 56% amino acid identity with NOX2. The gene for human NOX3 is located on chromosome 6. Sequence alignment and hydropathy plot analysis predict the overall structure of NOX3 to be highly similar to that of NOX1 and NOX2, in terms of transmembrane domains, the length of the extracellular loops, NADPH- and FAD-binding sites, and the localization of the heme-coordinating histidines (143, 454). To date, no splice variants of NOX3 have been reported.
Two different approaches led to the definition of NOX3 as an NADPH oxidase of the inner ear. A characterization of the "head tilt" mutant mouse through reverse genetics revealed underlying mutations in the NOX3 gene (677); as the head tilt mouse has vestibular defects, a functional role of NOX3 in the inner ear was established. Based on an EST clone derived from the inner ear, another study performed detailed analysis of NOX3 distribution by real time PCR and in situ hybridization and found very high NOX3 expression in the inner ear, including the cochlear and vestibular sensory epithelia and the spiral ganglion (54). Low levels of NOX3 can also be detected in other tissues, including fetal spleen (454), fetal kidney (54, 143), skull bone, and brain (54).
Nothing is currently published about the promoter region of NOX3; however, given the highly restricted tissue distribution, it seems likely that the expression is under the control of a distinct set of regulatory factors. Similarly, nothing is known about the subcellular localization of NOX3.
Our present knowledge on subunit dependence of NOX3 is as follows.
A) P22PHOX. NOX3 is a p22phox-dependent enzyme. NOX3 expression stabilizes the p22phox protein (891) and leads to p22phox translocation to the plasma membrane (892). In functional studies, p22phox is required for NOX3 activation (446, 891), and truncated p22phox inhibits ROS generation by NOX3 (446). Yet, there remain some doubts about the in vivo relevance of p22phox for NOX3 function, as no vestibular dysfunction has been reported for p22phox-deficient CGD patients.
B) ABSENCE OF CYTOSOLIC SUBUNITS. In the absence of cytosolic subunits, heterologously expressed NOX3 was found either to be inactive (147), weakly active (54, 892), or substantially active (891).
C) NOXO1. An enhanced activation of NOX3 in the presence of NOXO1 was found in all studies (54, 147, 891, 892). The strongest argument for a crucial role of NOXO1 in NOX3 activation however comes from in vivo studies demonstrating that inactivation of NOXO1 mimicks the phenotype of NOX3-deficient mice (463). Thus there can be little doubt that NOXO1 is in vivo an essential partner of NOX3.
D) NOXA1. The results on the requirement for NOXA1 are contradictory: while some studies found enhancement of NOX3 activity through NOXA1 (54, 892), others did not (147, 891). Thus the results of heterologous expression studies depend on the experimental conditions, and in vivo data with NOXA1-deficient animals will be necessary to clarify the issue.
E) P47PHOX AND P67PHOX. In heterologous expression studies, p47phox and p67phox are capable of activating NOX3 (54, 147, 891, 892); however, the physiological relevance of these finding is, at least with respect to p47phox, questionable as the loss of NOXO1 function in mice suffices to mimick the NOX3 deletion phenotype (463) and no vestibular symptoms have been described in p47phox-deficient mice or patients.
F) RAC. The Rac dependence of NOX3 is also still a matter of debate. Two studies suggest a Rac independence (144, 891), while the results of a third study suggest an effect (892). The differences in the findings may be due to a less strict requirement for Rac in NOX3 activation, or to the presence of endogenous Rac in the cells which found Rac independence.
Is the NOX3 system activation dependent or constitutively active? Present biochemical evidence would rather argue in favor of a constitutive activation: NOXO1, the key subunit for NOX3 activation, constitutively activates NOX3 in reconstituted systems (see references above). However, this is much less clear from a physiological point of view: why should there be a constitutively active ROS-generating system in the inner ear? Thus in vivo studies will be necessary to clarify the point.
NOX4 was originally identified as an NADPH oxidase homolog highly expressed in the kidney (294, 813). While NOX1-NOX3 represent an evolutionarily closely related subgroup of NOX enzymes, NOX4 is more distant, sharing only 39% identity to NOX2. The gene for human NOX4 is located on chromosome 11. The existence of four NOX4 splice variants has been suggested (315). NOX4 antibodies recognize two bands, one of 75–80 kDa and a second of 65 kDa from both endogenous NOX4-expressing cells (smooth muscle and endothelium) and NOX4-transfected Cos7 cells (378, 403, 813). The subcellular distribution of the two bands was distinct (378). The fact that two molecular masses are detected and that NOX4 contains four putative N-glycosylation sites might suggest that NOX4 is glycosylated, although treatment with N-glycosidase F failed to reduce the protein to a single band (813).
In addition to its strong expression in the kidney, NOX4 mRNA is also found in osteoclasts (969, 973), endothelial cells (14, 392, 901), smooth muscle cells (247, 383, 425, 510, 699, 836), hematopoietic stem cells (704), fibroblasts (170, 176, 211), keratinocytes (135), melanoma cells (105), and neurons (900).
Induction of NOX4 mRNA expression is observed under the following conditions: in response to endoplasmic reticulum stress (699), shear stress (402), carotid artery injury (856), hypoxia and ischemia (842, 900), and transforming growth factor (TGF)-
1 and tumor necrosis factor (TNF)- stimulation of smooth muscle (619, 836). Upregulation of NOX4 (mRNA and protein) has been reported in response to angiotensin II (377, 955, 964) (but one study found an angiotensin II-induced downregulation in NOX4 mRNA, Ref. 510). The angiotensin II-induced upregulation of NOX4 mRNA was prevented by pigment epithelium-derived factor (PEDF) (964). Downregulation of NOX4 mRNA and protein is observed in response to PPAR- ligands (403).
In vascular smooth muscle, NOX4 is described in proximity to focal adhesions (378). In transfected cells, NOX4 localization is mostly observed in the endoplasmic reticulum (ER), whether green fluorescent protein (GFP)-tagged NOX4 is used (901) or distribution is assessed by immunofluorescence (584). While a functional role for NOX4 in the ER is entirely possible, such a localization may also represent an accumulation at its site of synthesis. Puzzling observations come from vascular smooth muscle and endothelial cells, where NOX4 expression in the nucleus is suggested by several lines of arguments (immunofluorescence, electron microscopy, nuclear Western blots, and nuclear ROS generation) (378, 494). It is however difficult to understand how a protein that spans the membrane six times can be found in a presumably membrane-free space, such as the interior of the nucleus.
Our present knowledge on subunit dependence of NOX4 is as follows.
A) P22PHOX. NOX4 is a p22phox-dependent enzyme. NOX4 colocalizes and coimmunoprecipitates with p22phox; NOX4 also stabilizes the p22phox protein (24). Importantly, functional studies also demonstrate a p22phox requirement for NOX4-dependent ROS generation (446, 584). p22phox mutants lacking the proline-rich COOH terminus are still fully active in supporting NOX4 activity, while such mutants are not sufficient for NOX1, -2, and -3 activation.
B) NOX4 DOES NOT REQUIRE CYTOSOLIC SUBUNITS. NOX4 does not require cytosolic subunits for its activity, and upon heterologous expression, it is active without the need for cell stimulation (294, 584, 813).
C) RAC. In heterologously NOX4-expressing cells, Rac is not required for activity (584). Yet, at least in some endogenously NOX4-expressing cells, a Rac requirement has been documented (311, 410). Whether such a Rac requirement reflects a direct Rac/NOX4 interaction or is rather indirect remains to be seen.
As discussed above, NOX4 might be a constitutively active enzyme. However, not all available data favor this concept. NOX4 activation is observed under the following experimental conditions: 1) lipopolysaccharide (LPS)-stimulated HEK293 cells (686), 2) insulin-stimulated adipocytes (569), 3) angiotensin II- or high glucose-stimulated mesangial cells (311, 410), and 4) PMA-stimulated vascular endothelial cells (494). Mechanisms of NOX4 activation might include a direct binding of TLR4 to NOX4 (686). The angiotensin II and the high glucose stimulation are attributed to a Rac1-dependent NOX4 activation (311, 410).
A peculiarity of NOX4 is the fact that upon its heterologous expression, hydrogen peroxide, rather than superoxide, is detected (584). This should not, however, be taken as proof of direct hydrogen peroxide generation by the enzyme. Indeed, the most likely explanation is that the localization of the enzyme within intracellular organelles results in the release of superoxide into the lumen of the organelles where it rapidly dismutates into hydrogen peroxide. It is then the nonpolar hydrogen peroxide that is able to diffuse through membranes and reach the extracellular space.
NOX5 was discovered in 2001 by two groups. Cheng et al. (143) described it as a cDNA predicting a protein with 565 amino acids, while Banfi et al. (56) described it as cDNA predicting a protein of over 700 amino acids. The human NOX5 gene is located on chromosome 15. The NOX5 isoforms described by Banfi et al. (NOX5, -, -, and -) distinguish themselves from the NOX1–4 enzymes by the presence of a long intracellular NH2 terminus containing a Ca2+-binding EF hand domain (56, 58). The fifth isoform described by Cheng et al. (NOX5
or NOX5-S) lacks the EF-hand region and therefore has an overall structure more similar to NOX1–4 (143). On immunoblots, NOX5 is described as an 85-kDa protein (103). This would be consistent with its predicted molecular mass and suggests that the protein is not glycosylated. As seen for NOX2, NADH cannot replace NADPH as a cytoplasmic electron donor for NOX5 (58).
NOX5 mRNA expression is described in testis, spleen, lymph nodes, vascular smooth muscle, bone marrow, pancreas, placenta, ovary, uterus, stomach, and in various fetal tissues (56, 143, 770). Within the testis, the NOX5 message is localized to pachytene spermatocytes. Within the spleen, NOX5 shows a distinct localization within the mantle zone, which is rich in mature B cells, and in the periarterial lymphoid sheath area, which is enriched with T lymphocytes (56). Interestingly, NOX5 could not be detected within circulating lymphocytes (56). These data are based on mRNA expression; no data on the tissue distribution or subcellular distribution of the NOX5 protein are published. Presently there is also no information on the NOX5 promoters or on factors controlling gene expression of the EF-hand expressing NOX5 isoforms (-). However, a recent study shed first light on mechanisms regulating expression of the NOX5 isoform: acid induces NOX5 expression in Barrett's esophageal adenocarcinoma cells through mechanisms involving the cAMP response element binding protein CREB (274).
Nothing is known about the activation of the EF hand-deficient NOX5; thus the activation mechanisms summarized below are based on studies using EF hand-containing NOX5 isoforms. NOX5 does not require p22phox for activity, as demonstrated by siRNA suppression of p22phox leading to a decrease in the activity of NOX1 to NOX4, but not of NOX5 (446). NOX5 does not require cytosolic organizer or activator subunits (56) and has been shown to function in a cell-free system without the requirements of any cytosolic proteins (58). As predicted by the presence of EF hands, activation of NOX5 is mediated by an increase in the cytoplasmic Ca2+ concentration (58). The Ca2+-binding domain of NOX5 behaves as an independent folding unit and undergoes conformational changes in response to Ca2+ elevations (58). This is thought to activate the enyzme through an intramolecular protein-protein interaction between the Ca2+-binding region and the catalytic COOH terminus of the enzyme (56, 58).
For several novel NOX isoforms, the identification of the protein preceded the definition of its function. In the case of DUOX1 and DUOX2, the situation was reversed. It had been known for a long time that thyroid epithelial cells produce H2O2 at the apical plasma membrane in a Ca2+- and NADPH-dependent manner (88). Researchers in the thyroid field were actively looking for an NADPH oxidase. It took 15 years from the discovery of this function to the identification of DUOX proteins (originally called thyroid oxidase). They were identified from thyroid gland by two groups using different methods: purification and partial sequencing of the DUOX2 enzyme followed by rapid amplification of cDNA ends polymerase chain reaction (RACE PCR) (228) and low-temperature hybridization of a thyroid cDNA phage library with a NOX2 probe (189, 228). The genes for both human DUOX isoforms are located on chromosome 15. The two DUOX genes are somewhat unusual in their arrangement. They are arranged in a head-to-head configuration, separated by a relatively short (16 kb) region with the direction of transcription away from one another (675).
In addition to a NOX1–4 homology domain and an EF-hand region, DUOX proteins have a seventh transmembrane domain at the NH2 terminus with an ecto-facing peroxidase like domain. Within the NOX backbone, DUOX isoforms share 50% identity with NOX2 (189). An NH2-terminally truncated form of DUOX2 mRNA has been found in rat thyroid cell lines (625).
DUOX enzymes are glycosylated. Both DUOX1 and DUOX2 have two N-glycosylation states: the high mannose glycosylated form found in the ER, which runs by gel electrophoresis at 180 kDa, and a fully glycosylated form found at the plasma membrane that runs at 190 kDa (188, 624). Carbohydrate content analysis of plasma membrane DUOX revealed specific oligosaccharides indicative of Golgi apparatus processing (623). When totally deglycosylated, the molecular mass of both DUOX1 and DUOX2 drops to 160 kDa (188).
It is not clear whether the peroxidase homology domain of DUOX enzymes functions as a peroxidase. One study suggests that DUOX peroxidase homology domains, when expressed as recombinant proteins, have a peroxidase function (239). However, from a structural point of view, this is surprising. Indeed, the DUOX peroxidase homology domains lack many amino acid residues identified as essential for peroxidase function (168, 181, 653). The fact that a peroxidase is usually coexpressed in DUOX expressing systems, e.g., thyroid peroxidase in the thyroid gland and lactoperoxidase in salivary glands, also questions the peroxidase function of DUOX. This is particularly well documented for the thyroid, where thyroid peroxidase deficiency leads to severe hypothyroidism, due to a lack of peroxidase-dependent hormone synthesis (690). Still, the peroxidase homology region of DUOX2 seems to be of functional importance, as hypothyroidism in patients with mutations in the extracellular domain has been reported (918).
Based on its homology with NOX2 and the fact that heme enzymes are monoelectron transporters, DUOX enzymes should generate superoxide. However, a generation of hydrogen peroxide by thyrocytes has been detected in many studies. This led to a heated debate over the question of whether the thyroid oxidase directly generates hydrogen peroxide or whether the hydrogen peroxide generation occurs via a superoxide intermediate (231, 527, 645, 646). In a recent study, the immature, partially glycosylated form of DUOX2 generated superoxide, while the mature form generated hydrogen peroxide (27). The authors speculate that posttranslational modifications favor intramolecular dismutation of superoxide to hydrogen peroxide. Taken together, it is likely that the primary product of DUOX enzymes is superoxide and that a rapid dismutation precludes in many instances the detection of a superoxide intermediate. The substrate selectivity for human DUOX has not been defined; however, the GXGXXPF sequence typical of NADPH over NADH substrate selectivity is conserved, and in the sea urchin DUOX homolog Udx1 has been shown to favor the substrate NADPH to NADH (957).
Both DUOX1 and DUOX2 are highly expressed in the thyroid (189, 228). In addition, DUOX1 has been described in airway epithelia (271, 299, 794) and in the prostate (931).
DUOX2 is found in the ducts of the salivary gland (299); in rectal mucosa (299); all along the gastrointestinal tract including duodenum, colon, and cecum (230, 246); in airway epithelia (271, 794); and in prostate (931).
Induction of DUOX enzymes has been described. DUOX1 is induced in response to interleukin (IL)-4 and IL-13 in respiratory tract epithelium (352). DUOX2 expression was induced in response to interferon- in respiratory tract epithelium (352), in response to insulin in thyroid cell lines (625), and during spontaneous differentiation of postconfluent Caco-2 cells (246). Some studies (624, 625), but not others (189), find effects of forskolin, an adenylate cyclase activator, on DUOX expression. The putative promoters of DUOX1 and DUOX2 do not resemble each other and differ from promoters of other known thyroid-specific genes. The DUOX1, but not the DUOX2, promoter is GC rich and has putative SP-1 binding sites (675).
In thyrocytes, DUOX enzymes localize to the apical membrane (189, 228), although it appears that substantial amounts are found intracellularly, presumably in the ER (188). In airway epithelia, DUOX enzymes also localize to the apical membrane, as revealed by staining with an antibody that recognizes both DUOX1 and DUOX2 (794).
When heterologously expressed, DUOX enzymes tend to be retained in the ER, and superoxide generation can be measured only in broken cell preparations (27). This observation led to the discovery of DUOX maturation factors, which are ER proteins termed DUOXA1 and DUOXA2 (319). DUOX maturation factors seem to be crucial in overcoming ER retention of DUOX enzymes. DUOX enzymes do not require activator or organizer subunits; however, the p22phox requirement is still a matter of debate. DUOX enzymes coimmunoprecipitate with p22phox (931), but there is no evidence for enhanced DUOX function upon coexpression of p22phox (27, 188, 931).
Studies on the activation of heterologously expressed DUOX2 in membrane fractions indicated that the enzyme 1) does not require cytosolic activator or organizer subunits and 2) can be directly activated by Ca2+, suggesting that
