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Mobley HLT, Mendz GL, Hazell SL, editors. Helicobacter pylori: Physiology and Genetics. Washington (DC): ASM Press; 2001.

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Helicobacter pylori: Physiology and Genetics.

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Chapter 15Evasion of the Toxic Effects of Oxygen

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School of Science, Food and Horticulture, College of Science, Technology and Environment, University of Western Sydney, Sydney, Australia

Oxygen as a Toxic Species

Oxygen is an efficient terminal electron acceptor in respiratory pathways. During aerobic respiration the electron transport chain generates free radical oxygen species as a result of electron leakage; this generation of toxic species is proportional to the oxygen tension (51). In addition, toxic oxygen species (TOS) may be formed exogenously, for example, by chemical processes or through radiation. TOS also result from the oxidative burst of polymorphonuclear leukocytes (PMN). Infection with Helicobacter pylori induces an inflammatory response (gastritis), which leads to an increase in the level of TOS in the gastric mucosa and the gastric juice (4, 2426, 59). This increase in the level of toxic metabolites is probably the result of the generation of the superoxide anion (O2·− ), a reactive TOS, formed as part of the oxidative burst of PMN and enzymic activities of gastric epithelial cells. There is evidence that H. pylori infection leads to increased production of O2·− via NADPH oxidase in gastric cells, stimulated by lipopolysaccharide as well as xanthine oxidase, another mechanism for the generation of oxygen-derived free radicals (8, 80). In response to increased superoxide anion production in gastric tissue, changes have been detected in the level of expression of human superoxide dismutase (SOD) (12). Human gastric SOD exists as a cytoplasmic copper-zinc-superoxide dismutase (Cu, Zn-SOD) found in gland cells of the gastric body and antral mucosa, and as a manganese-superoxide dismutase (Mn-SOD) within mitochondria (63). An increase in the amount and activity of Mn-SOD has been observed in response to H. pylori infection and gastritis, whereas the amount and activity of the Cu, Zn-SOD remained constant or decreased slightly (39). It has been suggested that the induction of Mn-SOD is in response to increased cytokine production within the inflamed gastric mucosa (39). This situation is reversed following successful treatment of the infection (38). The data suggest that within the gastric environment H. pylori may be exposed to increased levels of TOS. In such an environment it is important for bacterial survival that the impact of such TOS be neutralized.

Reducing the Impact of TOS

How do microorganisms manage their exposure to TOS? Several strategies may be adopted, governed in part by determinants such as whether the toxic species are generated endogenously or exogenously. Microorganisms may neutralize TOS by mechanisms that include the enzymes SOD, catalase, peroxidases, and a variety of reductases. Also, they may modulate intracellular oxygen concentration or redox potential, thus minimizing their exposure to oxidative damage, or minimize such damage through the evolution of cellular structures resistant to oxidative damage. Finally, bacterial cells may overcome the effects of oxidative damage through efficient DNA repair mechanisms. There are many studies on such mechanisms in other organisms, and indeed, a great part of our understanding of these mechanisms in H. pylori is based predominantly on comparisons with the systems present in other organisms.

Neutralization of TOS

There are two prominent enzymes that facilitate resistance to oxidative damage in H. pylori, catalase (KatA) and SOD (43, 54, 65, 67, 77). In addition, there is genetic and biochemical evidence for the presence of at least two other enzyme systems involved in resistance to oxidative damage, alkylhydroperoxide reductase (Ahp) and thioredoxin-linked thiol peroxidase (scavengease).

Catalase

Catalase has been studied quite comprehensively in many eukaryotic and prokaryotic systems, beginning with studies in the 1800s (53). It has been used and still is used today as a diagnostic tool in bacterial identification in medical microbiology. The major function of catalase is to protect cells from the damaging effects of hydrogen peroxide (H2O2), catalyzing the dismutation of H2O2 into water and oxygen (equation 1). Consequently, catalase is an extremely important enzyme in an organism's response to oxidative stress.

Image ch15e1.jpg

Hydrogen peroxide is generated as a by-product of aerobic respiration, which uses oxygen as a terminal electron acceptor and can give rise to reactive oxygen species such as O2·− and H2O2. SOD is capable of removing the superoxide anion, but this results in the generation of more H2O2 (equation 2).

Image ch15e2.jpg

Nonetheless, the amounts of hydrogen peroxide and superoxide radicals produced during aerobic respiration are quite small in comparison with the quantity released during the respiratory or oxidative burst produced by PMN.

Exposure to H2O2 can be catastrophic for many organisms, yet the reactions between H2O2 and organic molecules, such as proteins and DNA, remain unclear. This is largely due to the rapid formation of other reactive oxygen species (ROS), which appear to be more reactive than H2O2 (33). The formation of other reactive oxygen radicals is due in part to interactions between H2O2 and reduced metallic ions found in all biological systems. The greatest risk that is posed to any cell, in terms of ROS, occurs when H2O2 reacts with reduced iron or copper ions (83) to form hydroxyl radicals (OH·) in a Fenton reaction (16). The hydroxyl radicals will react with most biological and organic molecules in oxidation reactions. Exposure to hydrogen peroxide, either through direct or indirect action, can result in DNA damage (therefore being mutagenic), lipid damage, and inhibition of the activities of enzymes and other proteins through oxidation. This is a probable explanation for the decreased growth rates observed in bacterial cell cultures in the presence of H2O2 (78).

"Typical" catalases characteristic of eukaryotes are homotetrameric with subunit mass between 55 and 65 kDa, and no heme prosthetic group per subunit (53), as indicated by the strong Soret band at 402 to 406 nm, with minor peaks at 500 to 505, 535 to 540, and 620 to 635 nm (45). Typical catalases differ from catalase-peroxidases found in a number of bacterial species in that they do not display peroxidase activity (32). H. pylori catalase is homotetrameric; each subunit has a mass of 58.7 kDa (as determined by the inferred amino acid sequence) and one heme prosthetic group (43, 54, 65). The enzyme is a mono-functional catalase, i.e., it lacks peroxidase activity (43). The activity of the H. pylori catalase is pH independent, with no difference between pH 5.25 and 8.95 (43). The enzyme has good thermostability, retaining catalytic activity after incubation at 56°C for 1 h (43). These properties are consistent with those of typical eukaryote catalases (43, 58).

Catalase is expressed in the cytoplasm and probably in the periplasm of H. pylori. There is some limited evidence supporting the presence of catalase on the cell surface, a unique occurrence in H. pylori, and possibly owing to autolytic events (43, 68, 71). However, Mori et al. (57) were unable to detect catalase activity in the supernatant of culture media after 24 h of growth, concluding that it was unlikely that the enzyme is secreted into the surrounding environment. The sequence does not show a cleavable N-terminal signal peptide, as is the case with many other periplasmic proteins (41), thus the putative translocation of the enzyme to the periplasm would be Sec-independent. H. pylori catalase is expressed during exponential growth and is not induced when the cells enter stationary phase as is the case with some bacterial catalases/catalase-peroxidases, for example, in Escherichia coli (43, 55).

A unique property of H. pylori catalase is an isoelectric point (pI) in the range of 9.0 to 9.3 (43). To date it is the only basic catalase that has been characterized. Catalases produced by other organisms usually have a pI in the range of 4.5 to 5.0 (58). The basic pI of H. pylori catalase is largely due to the high lysine and arginine content in the enzyme. The release of the genome made it clear that many proteins of H. pylori have a basic pI (2, 82). The biological relevance of the pI of catalase and other proteins of H. pylori has yet to be determined.

Catalase in H. pylori is distinctive because in situ there is a very rapid breakdown of hydrogen peroxide. The formation of oxygen occurs extremely fast, giving an "explosive" appearance of oxygen bubbles that is characteristic of catalase tests performed on whole H. pylori cells. The kinetic properties of the enzyme do not necessarily shed light on the activity observed when whole cells are exposed to H2O2. Helicobacter pylori catalase has a Km of 43 ± 3 mM and a Vmax of 60 ± 3 mmol/min/mg of protein (43). The Km is 3 to 10 times higher than those of other bacterial catalases, suggesting that the enzyme is relatively inefficient. Although the affinity of the protein for the substrate is not as high as those of some other catalases, it is likely that the high activity characteristic of H. pylori cells is due to the amount of enzyme present, typically accounting for >1% of the cells' total protein content (42, 43, 54), and/or the rapid turnover of substrate.

Catalases, like other proteins, are susceptible to damage by hydrogen peroxide, but the catalase of H. pylori appears to be quite stable at very high concentrations of hydrogen peroxide. This property appears to be shared with only a few other catalases, for example, of some Mycobacterium spp. (36, 43, 58). It may be hypothesized that the stability of the catalase of these bacteria in the presence of high concentrations of hydrogen peroxide is an adaptation by these organisms to environments comparatively rich in reactive oxygen species.

The H. pylori catalase gene katA from four different strains of the bacterium has been sequenced (2, 54, 65, 82). Not surprisingly, all sequences are almost identical with very few nucleotide variations. The inferred amino acid sequence of catalase revealed the presence of an NADPH-like binding motif similar to that of bovine liver catalase and other typical catalases. The residues involved in NADPH binding in bovine liver catalase are R-202, D-212, K-236 (all binding to the O2′ phosphate of NADP+), and H-304 (binding to the pyrophosphate group) (34). This sequence appears to be semiconserved in the H. pylori catalase; R-184, D-194, H-218 (conserved replacement), and L-286 (nonconservative change). Whether this sequence allows for NADPH binding remains to be determined. However, other data suggest that H. pylori catalase may bind NADH rather than NADPH (54). The inferred amino acid sequence of the protein reveals an adenylate-binding motif (GXGXXG) consistent with NADH binding, different from the NADPH adenylate-binding motif (GXGXXA) (72).

In typical catalases the presence of NADPH is important to maintain an active enzyme. The dismutation of H2O2 occurs by way of an intermediate form of catalase termed "compound I." The formation and decomposition of this intermediate occurs too rapidly for it to be detected by spectroscopy (47). Compound I (a nominal Fe5+ state) is formed by a two-electron oxidation involving H2O2 (equation 3), which then reacts with a second molecule of H2O2, returning the enzyme to its original state (Fe3+) (equation 4) (23). In the presence of excess H2O2 (or with other hydrogen donors), a second intermediate, termed "compound II," is formed (equation 5). Compound II is the result of the one electron oxidation of catalase (thus forming an Fe4+ intermediate). This enzyme intermediate does not react with H2O2 and thus the accumulation of compound II leads to the deactivation of catalase (11).

Image ch15e3.jpg
Image ch15e4.jpg
Image ch15e5.jpg

Formation of compound II can be reversed or inhibited by NADPH bound to catalase (11, 44). Four molecules of NADPH bind to the tetrameric structure of bovine liver catalase (49). This reduced dinucleotide is not essential for the catalytic action of the enzyme, but it is believed that NADPH reduces compound II via a one electron transfer reaction to yield NADP+ and the active native catalase (11).

All strains sequenced have the same genes flanking katA; upstream is frpB coding for an iron-binding protein, and downstream is an open reading frame (ORF) of unknown function. On the basis of sequence homology, a putative Fur-Box (ferric uptake regulator) has been identified upstream of katA (54, 65). Usually, the Fur protein mediates iron repression in gram-negative bacteria, and it would appear that the expression of katA might be regulated by this putative Fur-Box (Fig. 1). Although limited studies have been performed on the regulation of katA (54, 65), the level of catalase activity drops when H. pylori is grown in blood-based media, as opposed to serum-based media, suggesting that iron availability may have a role in the expression of katA (43). Studies by Bereswill et al. indicate that the H. pylori Fur homolog is functional as an iron-dependent transcriptional repressor (9). In Campylobacter jejuni, a bacterium of the same family as H. pylori, expression of catalase (katA) is repressed by iron, and regulation of catalase appears to be mediated by both Fur and the peroxide stress regulator PerR (84, 85). These findings would support the hypothesis that the putative Fur-Box of H. pylori is functional.

Figure 1

Figure 1

. Diagrammatic representation of katA and surrounding genes. Adapted from Manos et al. (54) with permission.

Catalase is not essential for growth and survival of H. pylori in vitro (54, 65, 89). Vaccine studies indicate that the enzyme is a highly effective antigen, suggesting that it may be essential in vivo (71). However, proof that catalase is essential in vivo remains to be established, as no catalase-negative mutants have been employed in animal model studies.

SOD

SOD catalyzes the dismutation of superoxide ions to hydrogen peroxide, which may be deactivated by catalase or peroxidase. The SOD of H. pylori is a typical prokaryotic iron-containing enzyme (Fe-SOD), consisting of two identical subunits each with an apparent molecular mass of 24 kDa (77). Three electromorphs or isoforms of Fe-SOD have been identified in different strains of H. pylori. These isoforms are the products of mutations leading to an altered pI (10). Unlike other bacteria that may express either an Mn-SOD or Cu, Zn-SOD, these forms of the enzyme were not detected in H. pylori by Spiegelhalder et al. (77), nor are they found in the genome (2, 82).

The different types of superoxide dismutase, Cu, Zn-SOD, Fe-SOD, and Mn-SOD, appear to support various functions in resistance to oxidative stress by cells. The dimeric prokaryotic Cu, Zn-SOD, which differs from the corresponding eukaryotic SOD, is usually expressed in the periplasm of gram-negative bacteria (27, 35). The Cu, Zn-SOD of E. coli is more resistant to inactivation by H2O2 than the eukaryotic enzyme and appears to be an important virulence determinant conferring resistance to oxidative damage induced by the respiratory burst of phagocytic cells (6). Indeed, the Cu, Zn-SOD of Salmonella appears essential to serious systemic disease (33).

In contrast, Mn-SOD is found in the cytosol and does not appear to be a critical virulence determinant. Instead, it appears to fulfill a "housekeeping" role, protecting against superoxide generated endogenously in bacteria such as Bordetella pertussis (40). Similarly, the Fe-SOD are cytosolic enzymes important in the management of endogenously generated superoxide (37). In H. pylori the absence of a leader sequence suggests that its Fe-SOD is also cytosolic (77). Interestingly, Fe-SOD show strong structural conservation between the prokaryotic and eukaryotic enzymes, as is the case for the H. pylori enzyme (37, 77).

In members of the Enterobacteriaceae periplasmic Cu, Zn-SOD appears to be much more important than the cytosolic SOD as a virulence determinant (37). However, there is evidence that Fe-SOD may enhance intracellular survival of C. jejuni (67) and may also be important to the virulence of Trichomonas vaginalis (87). The significance of such observations in relation to the Fe-SOD of H. pylori has yet to be ascertained.

Alkylhydroperoxide reductase

Alkylhydroperoxide reductase (2, 67, 82) catalyzes the reduction of alkylhydroperoxide to the corresponding alcohol. In most bacteria alkylhydroperoxide reductase is a two-component system consisting of the proteins AhpF and AhpO; the latter is responsible for the peroxide reductase activity, while the accessory flavoenzyme, AhpF, possesses NADH or NADPH oxidase activities. The H. pylori gene tsaA is orthologous to E. coli ahpC (69, 70). Although a homolog of ahpF has not been identified in the genome of H. pylori, there is ample experimental evidence for the presence of NADH oxidase activity in the bacterium (74). Niimura et al. demonstrated that in Salmonella enterica serovar Typhimurium, in the absence of AhpF, NADH oxidase or NADH oxidase-like activities coupled to AhpC are sufficient to generate alkylhydroperoxide reductase activity (61, 62).

Little is known about the alkylhydroperoxide reductase of H. pylori. Yet this enzyme may be common within this family of bacteria. Baillon et al. (5) identified a homolog of ahpC in the microaerophile C. jejuni. Like H. pylori, C. jejuni appears to lack ahpF, encoding the large accessory flavoenzyme of alkylhydroperoxide reductase. Importantly however, insertional mutagenesis of ahpC in C. jejuni resulted in an increased sensitivity to oxidative stresses induced by cumene hydroperoxide and atmospheric air (5). These data suggest that it is likely that alkylhydroperoxide reductase is functional in H. pylori.

Thioredoxin-linked thiol peroxidase

Thiol peroxidase (scavengease) belongs to a recently identified family of bacterial antioxidant enzymes possessing thioredoxin-linked activity (92). Direct biochemical evidence for the existence of thiol peroxidase in H. pylori has been provided by an assay for antioxidant activity (88). These findings are supported by data from the genome indicating the presence of the gene HP390 (jhp991) encoding a putative thiol peroxidase (2, 82).

Thiol peroxidase is usually a small protein (~20 to 30 kDa) found in both prokaryotic and eukaryotic organisms including Haemophilus influenzae, Vibrio cholerae, E. coli, streptococci, and Entamoeba histolytica (15, 21, 22). Thiol peroxidase protects from inactivation enzymes sensitive to oxidative stress such as glutamine synthetase, by removing H2O2 in a metal-catalyzed oxidation system (equation 6).

Image ch15e6.jpg

The thiol specificity of the enzyme is determined by the observation that the oxidized form of thiol peroxidase is reactivated (converted back to its sulfhydryl form) by treatment with thiols (15, 60). This observation relates to the finding that one cysteine residue, Cys-94 in the E. coli enzyme, appears to be central to peroxidase activity (21).

In E. coli oxidative stress induces higher levels of expression of the enzyme (48), which is located in the periplasm (21). It has been suggested that thiol peroxidase complements the cytosolic enzymes in protecting bacteria from oxidative damage (21). However, in the amoeba E. histolytica the enzyme is located in the cytosol, not on the surface or extracellularly (15), thus its role may include protection from both endogenously and exogenously generated reactive oxygen metabolites.

Management of Redox Potential

The oxidation-reduction (redox) status of H. pylori is important, as changing the environmental oxygen concentration and hence the redox status of the cell can greatly affect metabolic processes and clinical outcomes. The redox state of a cell may be defined as the sum of the oxidized and reduced molecular species present, but it is usually expressed in relation to the ratio of the oxidized and reduced thiols. Oxidation of thiols leads to an increase in the disulfide forms of both proteins and smaller compounds such as glutathione (γ-glutamylcysteinylglycine), the major free thiol in most cells. Reduced glutathione (GSH) plays an important role in the maintenance of the redox balance of cells, as it can scavenge free radicals and be converted to oxidized glutathione (GSSG). The cycling of glutathione is critical for detoxification of free radicals in many organisms, with GSSG normally converted back to GSH by the enzyme glutathione reductase.

However, there is little evidence that GSH is important to the maintenance of the redox balance in H. pylori. On the basis of genome analyses, the bacterium does not appear to have a homolog of the gene encoding for typical glutathione reductases (2, 82). There is evidence that the major free thiol compound within H. pylori is cysteine (Jorgensen et al., unpublished data). This observation is consistent with data from a number of microaerobic protozoan species that lack detectable levels of glutathione and use cysteine as their major free thiol compound (13, 30, 31, 76). Cysteine appears not to be an appropriate free thiol compound for aerobic organisms, because in the presence of a metal catalyst it is oxidized much faster than glutathione. Indeed, cystine (oxidized cysteine) markedly enhances the cytotoxic response of E. coli to H2O2 and may impair the cell defense machinery through thiol-disulfide exchange reactions at the cell membrane (18). This does not appear to be as critical in microaerophiles. If cysteine is the primary free thiol compound in H. pylori, cycling of oxidized cysteine, that is, the maintenance of a reduced state, may depend on a thioredoxin-like reductase as has been proposed for Giardia duodenalis (14).

H. pylori contains two ORFs encoding putative thioredoxin reductases, designated HP0825 (jhp764) and HP1164 (JHP1091), and two ORFs encoding putative thioredoxins, designated HP0824 (jhp763) and HP1458 (JHP1351) (2, 82). Thioredoxin and thioredoxin reductase form an NADPH-linked thiol-dependent redox system able to reduce proteins selectively. The proteins encoded by HP0825 (jhp764) and HP0824 (jhp763) appear to be typical thioredoxin reductase and thioredoxin components of the thioredoxin system involved in stress response (90). The "alternative" thioredoxin reductase and thioredoxin encoded by HP1164 (JHP1091) and HP1458/JHP1351, respectively, may fulfill the role of the thioredoxin-like reductase of G. duodenalis necessary for the maintenance of free cysteine (14), and hence the redox state of the cell.

Managing the concentration of dissolved intracellular oxygen is another way to regulate the redox potential of the cell. NADH oxidases are used to regulate the oxygen concentration in different microaerobic organisms. This family of enzymes directly reduces molecular oxygen to hydrogen peroxide or water. In the genome of H. pylori no ORF homologous to typical NADH oxidases is apparent, but cytosolic NAD(P)H oxidase activities have been measured in the bacterium (75). However, it is possible that such NAD(P)H oxidase activities are the product of electron leakage from the reduced flavin cofactor of flavoprotein enzymes such as alkylhydroperoxide reductase, thioredoxin reductase, glutathione reductase, mercuric reductase, and dihydrolipoamide dehydrogenase (3, 17, 19, 20, 52, 64, 91).

In addition to the enzyme activities outlined above, the pentose phosphate pathway also plays a role in resistance to oxidative stress; among its several roles, it generates reducing power in the form of NADPH. In yeasts, mutations of enzymes of the pentose phosphate pathway lead to increased sensitivity to oxidative stress, and the pathway is required for the maintenance of the cellular redox state (46, 73). Indeed, in mammalian systems, glucose 6-phosphate dehydrogenase, which catalyzes the first step in the pentose phosphate pathway and which provides reductive potential in the form of NADPH, has been found to be essential in protecting cells against oxidative stress, yet it is not essential for pentose synthesis (66). The pentose phosphate pathway was one of the first complete biochemical pathways identified in H. pylori (56), but its role in the maintenance of the redox status has not been investigated.

Gene Regulation and Repair Mechanisms

A surprising finding in the genome of H. pylori was the absence of homologs of genes encoding the transcription regulatory sigma factors σ32 (heat shock) and σS (stress/stationary-phase) (2, 82). Notwithstanding the absence of genes coding for σ32, homologs of genes encoding GroEL, GroES, DnaK, DanJ, and GrpE were identified in the genome regulated by housekeeping σ70-like sigma factors (1, 2, 7, 79, 82) (reviewed further in chapter 29).

The induction of an inflammatory response by H. pylori infection leads to increased potential for oxidative damage of the bacterium. While H. pylori has the enzymatic capacity to deal with such oxidative stress, no homologs of the oxidative stress regulators OxyR, SoxR, SoxS, or SOS present in other bacteria (28, 29, 86), have been found in H. pylori DNA (2, 82). Together with the absence of sigma factor σS, these data suggest that either H. pylori has adapted to an environment of constant oxidative stress or the bacterium contains novel systems of protection yet to be discovered.

H. pylori appears able to perform mismatch repair, as suggested by the coding capacity for methyl transferases, DNA glycosylases, and MutS and UvrD proteins, involved in error-free and error-prone repairs (2, 82). The RecBCD pathway is the major pathway for recombination in wild-type E. coli cells (50), but this system appears to be absent in H. pylori. Homologous recombination may be performed by H. pylori through the RecF pathway. In E. coli, this pathway generally depends on the RecA, RecJ, RecN, RecR, RecG, and RuvABC proteins, whose genes are present in the H. pylori chromosome (reviewed further in chapter 24); and Thompson and Blaser demonstrated that recA H. pylori mutants were highly sensitive to UV light, methyl methanesulfonate, and exposure to mutagenic antibiotics such as metronidazole (81).

C. jejuni does not encode OxyR and, as discussed above, the regulation of catalase expression in this bacterium appears to be mediated by both Fur and PerR (84, 85). It has been suggested that PerR functions as a nonhomologous substitute for OxyR (84). H. pylori, like C. jejuni, does not encode OxyR, and we are left to ponder the potential for the existence of previously unidentified oxidative stress regulators encoded by its genome.

Conclusion

H. pylori is a microaerophile that colonizes the inflamed gastric mucosa of humans. These two facts suggest the presence of a network of systems needed to manage both the oxygen to which H. pylori is exposed and the oxidative stress induced by endogenous and exogenous processes. That oxygen and TOS are constant companions of H. pylori in vivo is reflected in the enzymes expressed to manage them and the regulatory and repair mechanisms developed by the bacterium to cope with this type of stress. Nonetheless, our understanding of how H. pylori evades and avoids toxic oxygen effects is far from complete; and despite the importance of the topic, the management of oxygen and oxidative stress in H. pylori is a relatively neglected subject area.

References

1.
Allan E. P. Mullany, Tabaqchali S. Construction and characterisation of a Helicobacter pylori clpB mutant and role of the gene in stress response. J. Bacteriol. 1998;180:426–429. [PMC free article: PMC106902] [PubMed: 9440536]
2.
Alm R., Ling L., Moir D., King B., Brown E., Doig P., Smith D., Noonan B., Guild B., DeJonge B., Carmel G., Tummino P., Caruso A., Uria-Nickelsen M., Mills D., Ives C., Gibson R., Merberg D., Mills S., JIang Q., Taylor D., Vovis G., Trust T. Genomic sequence comparison of two unrelated isolates of the human gastric pathogen Helicobacter pylori. Nature. 1999;397:176–180. [PubMed: 9923682]
3.
Arnér E. S. J., Bjornstedt M., Holmgren A. 1-chloro-2,4-dinitrobenzene is an irreversible inhibitor of human thioredoxin reductase—loss of thioredoxin disulfide reductase activity is accompanied by a large increase in NADPH oxidase activity. J. Biol. Chem. 1995;270:3479–3482. [PubMed: 7876079]
4.
Bagchi D., Bhattacharya G., Stohs S. J. Production of reactive oxygen species by gastric cells in association with Helicobacter pylori. Free Radical Res. 1996;24:439–450. [PubMed: 8804987]
5.
Baillon M. L., van Vliet A. H., Ketley J. M., Constantinidou C., Penn C. W. An iron-regulated alkyl hydroperoxide reductase (AhpC) confers aerotolerance and oxidative stress resistance to the microaerophilic pathogen Campylobacter jejuni. J. Bacteriol. 1999;181:4798–4804. [PMC free article: PMC93964] [PubMed: 10438747]
6.
Battistoni A., Donnarumma G., Greco R., Valenti P., Rotilio G. Overexpression of a hydrogen peroxide-resistant periplasmic Cu,Zn superoxide dismutase protects Escherichia coli from macrophage killing. Biochem. Biophys. Res. Commun. 1998;243:804–807. [PubMed: 9501009]
7.
Beier D., Spohn G., Rappuoli R., Scarlato V. Identification and characterization of an operon of Helicobacter pylori that is involved in motility and stress adaptation. J. Bacteriol. 1997;179:4676–4683. [PMC free article: PMC179311] [PubMed: 9244252]
8.
Benhamida A., Man W. K., Mcneil N., Spencer J. Histamine, xanthine oxidase generated oxygen derived free radicals and Helicobacter pylori in gastroduodenal inflammation and ulceration. Inflam. Res. 1998;47:193–199. [PubMed: 9628263]
9.
Bereswill S., Lichte F., Greiner S., Waidner B., Fassbinder F., Kist M. The ferric uptake regulator (Fur) homologue of Helicobacter pylori: functional analysis of the coding gene and controlled production of the recombinant protein in Escherichia coli. Med. Microbiol. Immunol. 1999;188:31–40. [PubMed: 10691091]
10.
Bereswill S., Neuner O., Strobel S., Kist M. Identification and molecular analysis of superoxide dismutase isoforms in Helicobacter pylori. FEMS Microbiol. Lett. 2000;183:241–245. [PubMed: 10675591]
11.
Bicout D., Field M., Gouet P., Jouve H. Simulations of electron transfer in the NADPH-bound catalase from Proteus mirabilis PR. Biochim. Biophys. Acta. 1995;1252:172–176. [PubMed: 7548161]
12.
Broide E., Klinowski E., Varsano R., Eshchar J., Herbert M., Scapa E. Superoxide dismutase activity in Helicobacter pylori-positive antral gastritis in children. J. Pediatr. Gastroenterol. Nutr. 1996;23:609–613. [PubMed: 8985854]
13.
Brown D. M., Upcroft J. A., Upcroft P. Cysteine is the major low molecular weight thiol in Giardia duodenalis. Mol. Biochem. Parasitol. 1993;61:155–158. [PubMed: 8259129]
14.
Brown D. M., Upcroft J. A., Upcroft P. A thioredoxin reductase-class of disulphide reductase in the protozoan parasite Giardia duodenalis. Mol. Biochem. Parasitol. 1996;83:211–220. [PubMed: 9027754]
15.
Bruchhaus I., Richter S., Tannich E. Removal of hydrogen peroxide by the 29 kDa protein of Entamoeba histolytica. Biochem. J. 1997;326:785–789. [PMC free article: PMC1218733] [PubMed: 9307028]
16.
Cadenas E. Biochemistry of oxygen toxicity. Annu. Rev. Biochem. 1989;58:79–110. [PubMed: 2673022]
17.
Calzi M. L., Poole L. B. Requirement for the two AhpF cystine disulfide centers in catalysis of peroxide reduction by alkyl hydroperoxide reductase. Biochemistry. 1997;36:13357–13364. [PubMed: 9341228]
18.
Cantoni O., Brandi G., Albano A., Cattabeni F. Action of cystine in the cytotoxic response of Escherichia coli cells exposed to hydrogen peroxide. Free Radical Res. 1995;22:275–283. [PubMed: 7757202]
19.
Carlberg I., Mannervik B. Oxidase activity of glutathione reductase effected by 2,4,6-trinitrobenzenesulfonate. FEBS Lett. 1980;115:265–268. [PubMed: 7398886]
20.
Carlberg I., Sahiman L., Mannervik B. The effect of 2,4,6-trinitrobenzenesulfonate on mercuric reductase, glutathione reductase and lipoamide dehydrogenase. FEBS Lett. 1985;180:102–106. [PubMed: 3917936]
21.
Cha M. K., Kim H. K., Kim I. H. Thioredoxin-linked "thiol peroxidase" from the periplasmic space of Escherichia coli. J. Biol. Chem. 1995;270:28635–28641. [PubMed: 7499381]
22.
Cha M. K., Kim H. K., Kim I. H. Mutation and mutagenesis of thiol peroxidase of Escherichia coli and a new type of thiol peroxidase family. J. Bacteriol. 1996;178:5610–5614. [PMC free article: PMC178398] [PubMed: 8824604]
23.
Chance B., Sies H., Boveris A. Hydroperoxide metabolism in mammalian organs. Physiol. Rev. 1979;59:527–605. [PubMed: 37532]
24.
Crabtree J. Immune and inflammatory responses to Helicobacter pylori infection. Scand. J. Gastroenterol. 1996;31:3–10. [PubMed: 8722376]
25.
Davies G. R., Banatvala N., Collins C. E., Sheaff M. T., Abdi Y., Clements L., Rampton D. S. Relationship between infective load of Helicobacter pylori and reactive oxygen metabolite production in antral mucosa. Scand. J. Gastroenterol. 1994;29:419–424. [PubMed: 8036457]
26.
Davies G. R., Simmonds N. J., Stevens T. R. J., Grandison A., Blake D. R., Rampton D. S. Mucosal reactive oxygen metabolite production in duodenal ulcer disease. Gut. 1992;33:1467–1472. [PMC free article: PMC1379529] [PubMed: 1452069]
27.
Degroote M. A., Ochsner U. A., Shiloh M. U., Nathan C., McCord J. M., Dinauer M. C., Libby S. J., Vazqueztorres A., Xu Y. S., Fang F. C. Periplasmic superoxide dismutase protects salmonella from product of phagocyte NADpH-oxidase and nitric oxide synthase. Proc. Natl. Acad. Sci. USA. 1997;94:13997–14001. [PMC free article: PMC28421] [PubMed: 9391141]
28.
Demple B. Redox signalling and gene control in the Escherichia coli soxrs oxidative stress regulon—a review. Gene. 1996;179:53–57. [PubMed: 8955629]
29.
Dhandayuthapani S., Mudd M., Deretic V. Interactions of oxyr with the promoter region of the oxyr and ahpc genes from Mycobacterium leprae and Mycobacterium tuberculosis. J. Bacteriol. 1997;179:2401–2409. [PMC free article: PMC178979] [PubMed: 9079928]
30.
Ellis J. E., Yarlett N., Cole D., Humphreys M. J., Lloyd D. Antioxidant defences in the microaerophilic protozoan Trichomonas vaginalis: comparison of metronidazole-resistant and sensitive strains. Microbiology. 1994;140:2489–2494. [PubMed: 7952198]
31.
Fahey R. C., Newton G. L., Arrick B., Overdankbogart T., Aley S. B. Entamoeba histolytica: a eukaryote without glutathione metabolism. Science. 1984;224:70–72. [PubMed: 6322306]
32.
Farr S., Touati D., Kogoma T. Effects of oxygen stress on membrane functions in Escherichia coli: role of HP1 catalase. J. Bacteriol. 1988;170:1837–1842. [PMC free article: PMC211039] [PubMed: 2832383]
33.
Farrant J. L., Sansone A., Canvin J. R., Pallen M. J., Langford P. R., Wallis T. S., Dougan G., Kroll J. S. Bacterial copper and zinc-cofactored superoxide dismutase contributes to the pathogenesis of systemic salmonellosis. Mol. Microbiol. 1997;25:785–796. [PubMed: 9379906]
34.
Fita I., Rossman M. The NADPH binding site of beef liver catalase. Proc. Natl. Acad. Sci. USA. 1985;82:1604–1608. [PMC free article: PMC397320] [PubMed: 3856839]
35.
Forest K. T., Langford P. R., Kroll J. S., Getzoff E. D. Cu,Zn superoxide dismutase structure from a microbial pathogen establishes a class with a conserved dimer interface. J. Mol. Biol. 2000;296:145–153. [PubMed: 10656823]
36.
Goldberg I., Hochman A. Three different types of catalase in Klebsiella pneumoniae. Arch. Biochem. Biophys. 1989;268:124–128. [PubMed: 2643382]
37.
Gort A. S., Imlay J. A. Balance between endogenous superoxide stress and antioxidant defenses. J. Bacteriol. 1998;180:1402–1410. [PMC free article: PMC107037] [PubMed: 9515906]
38.
Gotz J. M., Thio J. L., Verspaget H. W., Offerhaus G. J. A., Biemond I., Lamers C. B. H. W., Veenendaal R. A. Treatment of Helicobacter pylori infection favourably affects gastric mucosal superoxide dismutases. Gut. 1997;40:591–596. [PMC free article: PMC1027159] [PubMed: 9203935]
39.
Gotz J. M., Vankan C. I., Verspaget H. W., Biemond I., Lamers C. B. H. W., Veenendaal R. A. Gastric mucosal superoxide dismutases in Helicobacter pylori infection. Gut. 1996;38:502–506. [PMC free article: PMC1383104] [PubMed: 8707077]
40.
Graeffwohlleben H., Killat S., Banemann A., Guiso N., Gross R. Cloning and characterization of an MN-containing superoxide dismutase (Soda) of Bordetella pertussis. J. Bacteriol. 1997;179:2194–2201. [PMC free article: PMC178955] [PubMed: 9079904]
41.
Harris A., Hazell S. Evidence supporting post translational modification of the Helicobacter pylori catalase. XIIth International Workshop on Gastroduodenal Pathology and Helicobacter pylori. Helsinki, Finland 2–4 September 1999. Gut. 1999;45(Suppl. 111):A11.
42.
Hazell, S. L. 1990. Urease and catalase as virulence factors of Helicobacter pylori, p. 3–13. In H. Menge, M. Gregor, G. N. J. Tytgat, B. I. Marshall, and C. I. A. M. McNulty (ed.), Helicobacter pylori 1990. Springer-Verlag, Berlin, Germany.
43.
Hazell S., Evans D., Graham D. Helicobacter pylori catalase. J. Gen. Microbiol. 1991;137:57–61. [PubMed: 2045782]
44.
Hillar A., Nicholls P., Switala J., Loewen P. NADPH binding and control of catalase compound II formation: comparison of bovine, yeast and Escherichia coli enzymes. Biochem. J. 1994;300:531–539. [PMC free article: PMC1138194] [PubMed: 8002960]
45.
Hochman A., Shemesh A. Purification and characterisation of a catalase-peroxidase from the photosynthetic bacterium Rhodopseudomonas capsulata. J. Biol. Chem. 1987;262:6871–6876. [PubMed: 3571290]
46.
Juhnke H., Krems B., Kotter P., Entian K. D. Mutants that show increased sensitivity to hydrogen peroxide reveal an important role for the pentose phosphate pathway in protection of yeast against oxidative stress. Mol. Gen. Genet. 1996;252:456–464. [PubMed: 8879247]
47.
Keilin D., Hartree E. Properties of azide-catalase. Biochemistry. 1945;39:148–157. [PMC free article: PMC1258190] [PubMed: 16747875]
48.
Kim H. K., Kim S. J., Lee J. W., Lee J. W., Cha M. K., Kim I. H. Identification of promoter in the 5′-flanking region of the E. coli thioredoxin-linked thiol peroxidase gene: evidence for the existence of oxygen-related transcriptional regulatory protein. Biochem. Biophys. Res. Commun. 1996;221:641–646. [PubMed: 8630014]
49.
Kirkman H., Gaetani G. Catalase: a tetrameric enzyme with four tightly bound molecules of NADPH. Proc. Natl. Acad. Sci. USA. 1984;81:4343–4347. [PMC free article: PMC345585] [PubMed: 6589599]
50.
Kowalczykowski S. C., Dixon D. A., Eggleston A. K., Lander S. D., Rebrauer W. M. Biochemistry of homologous recombination in Escherichia coli. Microbiol. Rev. 1994;58:401–465. [PMC free article: PMC372975] [PubMed: 7968921]
51.
Ksenzenko M. Y., Vygodina T. V., Berka V., Rauge E. K., Konstantinov A. A. Cytochrome oxidase-catalyzed superoxide generation from hydrogen peroxide. FEBS Lett. 1992;297:63–66. [PubMed: 1312951]
52.
Lin S., Cullen W. R., Thomas D. J. Methylarsenicals and arsinothiols are potent inhibitors of mouse liver thioredoxin reductase. Chem. Res. Toxicol. 1999;12:924–930. [PubMed: 10525267]
53.
Loewen, P. 1997. Bacterial catalases, p. 273–308. In J. G. Scandalios (ed.), Oxidative Stress and the Molecular Biology of Antioxidant Defenses. Cold Spring Harbor Press, Plainview, N.Y.
54.
Manos J., Kolesnikow T., Hazell S. An investigation of the molecular basis of the spontaneous occurrence of a catalase negative phenotype in Helicobacter pylori. Helicobacter. 1998;4:1–7. [PubMed: 9546115]
55.
Meir E., Yagil E. Further characterization of the two catalases in Escherichia coli. Curr. Microbiol. 1985;12:315–320.
56.
Mendz G. L., Hazell S. L. Evidence for a pentose phosphate pathway in Helicobacter pylori. FEMS Microbiol. Lett. 1991;84:331–336.
57.
Mori M., Suzuki H., Suzuki M., Kai A., Miura S., Ishii H. Catalase and superoxide dismutase secreted from Helicobacter pylori. Helicobacter. 1997;2:100–105. [PubMed: 9432326]
58.
Nadler V., Goldberg I., Hochman A. Comparative study of bacterial catalases. Biochim. Biophys. Acta. 1986;882:234–241.
59.
Nalini S., Ramakrishna B. S., Mohanty A., Balasubramanian K. A. Hydroxyl radical formation in human gastric juice. J. Gastroenterol. Hepatol. 1992;7:497–501. [PubMed: 1327262]
60.
Netto L. E. S., Chae H. Z., Kang S. W., Rhee S. G., Stadtman E. R. Removal of hydrogen peroxide by thiol-specific antioxidant enzyme (TSA) is involved with its antioxidant properties. TSA possesses thiol peroxidase activity. J. Biol. Chem. 1997;271:15315–15321. [PubMed: 8663080]
61.
Niimura Y., Poole L. B., Massey V. Amphibacillus xylanus NADH oxidase and Salmonella typhimurium alkyl-hydroperoxide reductase flavoprotein components show extremely high scavenging activity for both alkyl hydroperoxide and hydrogen peroxide in the presence of S. typhimurium alkyl-hydroperoxide reductase 22 kDa protein component. J. Biol. Chem. 1995;270:25645–25650. [PubMed: 7592740]
62.
Niimura Y., Massey V. Reaction mechanism of Amphibacillus xylanus NADH oxidase alkyl-hydroperoxide reductase flavoprotein. J. Biol. Chem. 1996;271:30459–30464. [PubMed: 8940011]
63.
Nishiyama J., Mizuno M., Nasu J., Kiso T., Uesu T., Maga T., Okada H., Tomoda J., Yamada G., Tsuji T. Immunoelectron microscopic localization of copper-zinc superoxide dismutase in human gastric mucosa. Acta Histochem. Cytochem. 1996;29:215–220.
64.
Nordberg J., Zhong L., Holmgren A., Arner E. S. J. Mammalian thioredoxin reductase is irreversibly inhibited by dinitrohalobenzenes by alkylation of both the redox active selenocysteine and its neighboring cysteine residue. J. Biol. Chem. 1998;273:10835–10842. [PubMed: 9556556]
65.
Odenbreit S., Wieland B., Haas R. Cloning and genetic characterisation of Helicobacter pylori catalase and construction of a catalase deficient mutant. J. Bacteriol. 1996;178:6960–6967. [PMC free article: PMC178599] [PubMed: 8955320]
66.
Pandolfi P. P., Sonati F., Rivi R., Mason P., Grosveld F., Luzzatto L. Targeted disruption of the housekeeping gene encoding glucose 6-phosphate dehydrogenase (G6PD)—G6PD is dispensable for pentose synthesis but essential for defence against oxidative stress. EMBO J. 1995;14:5209–5215. [PMC free article: PMC394630] [PubMed: 7489710]
67.
Pesci E., Pickett C. Genetic organization and enzymatic activity of a superoxide dismutase from the microaerophilic human pathogen, Helicobacter pylori. Gene. 1994;143:111–116. [PubMed: 7515365]
68.
Phadnis S., Parlow M., Levy M., Ilver D., Caulkins C., Connors J., Dunn B. Surface localisation of Helicobacter pylori urease and a heat shock protein homolog requires bacterial autolysis. Infect. Immun. 1996;64:905–912. [PMC free article: PMC173855] [PubMed: 8641799]
69.
Poole L. B. Flavin-dependent alkyl hydroperoxide reductase from Salmonella typhimurium 2. Cystine disulfides involved in catalysis of peroxide reduction. Biochemistry. 1996;35:65–75. [PubMed: 8555199]
70.
Poole L. B., Ellis H. R. Flavin-dependent alkyl hydroperoxide reductase from Salmonella typhimurium 1. Purification and enzymatic activities of overexpressed AHPF and AHPC proteins. Biochemistry. 1996;35:56–64. [PubMed: 8555198]
71.
Radcliff F. J., Hazell S. L., Kolesnikow T., Doidge C., Lee A. Catalase, a novel antigen for Helicobacter pylori vaccination. Infect. Immun. 1997;65:4668–4674. [PMC free article: PMC175669] [PubMed: 9353048]
72.
Scrutton N. S., Berry A., Perham P. N. Redesign of the coenzyme specificities of a dehydrogenase by protein engineering. Nature. 1990;343:38–43. [PubMed: 2296288]
73.
Slekar K. H., Kosman D. J., Culotta V. C. The yeast copper zinc superoxide dismutase and the pentose phosphate pathway play overlapping roles in oxidative stress protection. J. Biol. Chem. 1996;271:28831–28836. [PubMed: 8910528]
74.
Smith M. A., Edwards D. I. Oxygen scavenging, NADH oxidase and metronidazole resistance in Helicobacter pylori. J. Antimicrob. Chemother. 1997;39:347–353. [PubMed: 9096184]
75.
Smith M. A., Edwards D. I. Redox potential and oxygen concentration as factors in the susceptibility of Helicobacter pylori to nitroheterocyclic drugs. J. Antimicrob. Chemother. 1995;35:751–764. [PubMed: 7559187]
76.
Smith N. C., Bryant C., Boreham P. F. L. Possible roles for pyruvate: ferredoxin oxidoreductase and thiol-dependent peroxidase and reductase activities in resistance to nitroheterocyclic drugs in Giardia intestinalis. Int. J. Parasitol. 1988;18:991–997. [PubMed: 3225121]
77.
Spiegelhalder C., Gerstenecker B., Kersten A., Schiltz E., Kist M. Purification of Helicobacter pylori superoxide dismutase and cloning and sequencing of the gene. Infect. Immun. 1993;61:5315–5325. [PMC free article: PMC281317] [PubMed: 8225605]
78.
Storz G., Tartaglia L. A., Farr S. B., Ames B. N. Bacterial defenses against oxidative stress. Trends Genet. 1990;6:363–368. [PubMed: 1965068]
79.
Suerbaum S., Thilberg J.-M., Kansau I., Ferrero R. L., Labigne A. Helicobacter pylori hspA-hspB heat shock gene cluster: nucleotide sequence, expression, putative function and immunogenicity. Mol. Microbiol. 1994;14:959–974. [PubMed: 7715457]
80.
Teshima S., Tsunawaki S., Rokutan K. Helicobacter pylori lipopolysaccharide enhances the expression of NADPH oxidase components in cultured guinea pig gastric mucosal cells. FEBS Lett. 1999;452:243–246. [PubMed: 10386599]
81.
Thompson S. A., Blaser M. J. Isolation of the Helicobacter pylori reca gene and involvement of the reca region in resistance to low pH. Infect. Immun. 1995;63:2185–2193. [PMC free article: PMC173284] [PubMed: 7768597]
82.
Tomb J.-F., White O., Kerlavage A., Clayton R., Sutton G., Fleischmann R., Ketchum K., Klenk H., Gill S., Dougherty B., Nelson K., Quackbush J., Zhou L., Kirkness E., Peterson S., Loftus B., Richardson D., Dodson R., Khalek H., Gludek A., McKenny K., Fitzegerald L., Lee N., Adams M., Hickey E., Berg D., Gocayne J., Utterback T., Peterson J., Kelley J., Cotton M., Weldman J., Fujii C., Bowman C., Watthey L., Wallin E., Hayes W., Borodovsky M., Karp P., Smith H., Fraser C., Venter J. The complete genome sequence of the gastric pathogen Helicobacter pylori. Nature. 1997;388:539–547. [PubMed: 9252185]
83.
Touati D. Iron and oxidative stress in bacteria. Arch. Biochem. Biophys. 2000;373:1–6. [PubMed: 10620317]
84.
van Vliet A. H. M., Baillon M. L. A., Penn C. W., Ketley J. M. Campylobacter jejuni contains two fur homologs: characterization of iron-responsive regulation of peroxide stress defense genes by the PerR repressor. J. Bacteriol. 1999;181:6371–6376. [PMC free article: PMC103772] [PubMed: 10515927]
85.
van Vliet A. H. M., Wooldridge K. G., Ketley J. M. Iron-responsive gene regulation in a Campylobacter jejuni fur mutant. J. Bacteriol. 1998;180:5291–5298. [PMC free article: PMC107575] [PubMed: 9765558]
86.
Vattanaviboon P., Varaluksit T., Mongkolsuk S. Modulation of peroxide stress response by thiol reagents and the role of a redox sensor-transcription regulator, OxyR in mediating the response in Xanthomonas. FEMS Microbiol. Lett. 1999;176:471–476.
87.
Viscogliosi E., Delgadoviscogliosi P., Gerbod D., Dauchez M., Gratepanche S., Alix A. J. P., Dive D. Cloning and expression of an iron-containing superoxide dismutase in the parasitic protist, Trichomonas vaginalis. FEMS Microbiol. Lett. 1998;161:115–123. [PubMed: 9561738]
88.
Wan X. Y., Zhou Y., Yan Z. Y., Wang W. L., Hou Y. D., Jin D. Y. Scavengase p20: a novel family of bacterial antioxidant enzymes. FEBS Lett. 1997;407:32–36. [PubMed: 9141476]
89.
Westblom T., Phadnis S., Langenberg W., Yoneda K., Madan E., Midkiff B. Catalase negative mutants of Helicobacter pylori. Eur. J. Clin. Microbiol. Infect. Dis. 1992;11:522–526. [PubMed: 1526235]
90.
Windle H. J., Fox A., Eidhin D. N., Kelleher D. The thioredoxin system of Helicobacter pylori. J. Biol. Chem. 2000;275:5081–5089. [PubMed: 10671551]
91.
Zhang Z., Hillas P. J., de Montellano P. R. O. Reduction of peroxides and dinitrobenzenes by Mycobacterium tuberculosis thioredoxin and thioredoxin reductase. Arch. Biochem. Biophys. 1999;363:19–26. [PubMed: 10049495]
92.
Zhou Y., Wan X. Y., Wang H. L., Yan Z. Y., Hou Y. D., Jin D. Y. Bacterial scavengase p20 is structurally and functionally related to peroxiredoxins. Biochem. Biophys. Res. Commun. 1997;233:848–852. [PubMed: 9168946]
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