No retraction, correction or concern found when last checked on Oct 26, 2026.

The Dual Role of Myeloperoxidase in Immune Response

Jürgen Arnhold

International Journal of Molecular Sciences · 2020 · Review · CC BY

Abstract

The heme protein myeloperoxidase (MPO) is a major constituent of neutrophils. As a key mediator of the innate immune system, neutrophils are rapidly recruited to inflammatory sites, where they recognize, phagocytose, and inactivate foreign microorganisms. In the newly formed phagosomes, MPO is involved in the creation and maintenance of an alkaline milieu, which is optimal in combatting microbes. Myeloperoxidase is also a key component in neutrophil extracellular traps. These helpful properties are contrasted by the release of MPO and other neutrophil constituents from necrotic cells or as a result of frustrated phagocytosis. Although MPO is inactivated by the plasma protein ceruloplasmin, it can interact with negatively charged components of serum and the extracellular matrix. In cardiovascular diseases and many other disease scenarios, active MPO and MPO-modified targets are present in atherosclerotic lesions and other disease-specific locations. This implies an involvement of neutrophils, MPO, and other neutrophil products in pathogenesis mechanisms. This review critically reflects on the beneficial and harmful functions of MPO against the background of immune response.

Full text

From PubMed Central, U.S. National Library of Medicine, as current on Oct 26, 2026. Paragraph numbers are ours, so an answer can point at one.

1. Immune Response and Tissue Destruction

1In humans and higher animals, protection against different threats that affect the homeostasis of host’s tissues is ensured by a coordinated action of the immune system in close association with activation of components of the acute phase, complement, coagulation, and contact systems. An immune response starts usually with the recruitment and activation of innate immune cells such as neutrophils, monocytes and others, and can be followed by the additional activation of dendritic cells and lymphocytes. Importantly, the immune system is involved to resist the invasion of viruses, bacteria, fungi and other pathogens, as well as to remove and replace damaged material.

2With these activities, immune response is primarily directed to diminish the degree of tissue destruction and to restore homeostatic conditions. Otherwise, immune cells themselves release numerous hazardous substances during their activation. These agents are predominantly used to inactivate, kill, and destroy pathogenic microorganisms. However, they can also damage cell material, cause necrotic cell death, and induce the release of antigens in the host. Altogether, the immune system plays a dual role with respect to cell and tissue damage. On the one hand, it protects cells and tissues from many destroying threats. On the other hand, it can provoke significant harm to host tissues.

3This general dual activity of the immune system is also reflected in the functional responses of many cells activated after their recruitment to inflammatory loci. Polymorphonuclear leukocytes (PMNs), also known as neutrophils, are key mediator cells of innate immunity. These cells are rapidly recruited to inflammatory sites. The infiltration of PMNs into infected and injured tissue is mediated by adhesion molecules, cytokines, chemotactic agents, and components of the extracellular matrix. In this way, through endothelium and adjacent tissue, neutrophils are stepwise activated and can phagocytose foreign microorganisms at the destination site. With the release of proteases, hydrolases, bactericidal proteins and others, and the generation of reactive species, PMNs contribute to the inactivation and killing of bacteria and fungi. Otherwise, these neutrophil-derived agents are also known to be able to damage intact cells and tissues.

4The heme protein myeloperoxidase (MPO) is a major active component in activated human neutrophils. It is also present in monocytes to a lesser extent, but usually lost during the maturation of these cells to macrophages. Some reports describe MPO-rich macrophage subpopulations in atherosclerotic plaques, circulating blood, and multiple sclerotic lesions. In macrophages, MPO can also result from phagocytosed apoptotic and necrotic PMNs or internalization of MPO-containing extracellular traps.

5In this review, the major focus is on the characteristics of neutrophil-derived MPO. The main properties of MPO and physiological consequences of MPO activation are summarized during immune response. This concerns the role of MPO in neutrophil functions during inactivation and killing of ingested microbes and fungi as well as the potential contribution of released MPO in the modification of proteins in disease progression. In sum, this heme protein plays also a dual role in immune response with both helpful and devastating properties. A hypothesis about the switch from the protecting to harmful functions of MPO, neutrophils, and general immune functions is presented too.

2.1. Selected Structural Properties

6To discuss the role of MPO in immune response, it is first necessary to consider major characteristics of this enzyme. As there are numerous comprehensive reviews about structural, biochemical, and redox properties of MPO, I will here mention only the main issues.

7Human myeloperoxidase is a cationic, dimeric protein with a ferric protoporphyrin IX (heme) in each subunit. In MPO, both heme groups are structurally and functionally identical. Each heme is associated with the surrounding apoprotein by three covalent bonds, hence the heme is slightly bow-shaped along the axis from pyrrole ring A to ring C, with a shift in the central iron ion by 0.2 Å to the proximal side. The three apoprotein-heme linkages determine the extraordinary biochemical and redox properties of this heme protein.

8In each subunit of MPO, the heme is deeply buried into the protein bulk and connected with the surface by a narrow substrate channel. At the entrance to the distal cavity, there is a conserved hydrophobic region among mammalian heme peroxidases. Aromatic molecules and some other substrates can get close enough to the heme to allow an electron to transfer between the heme and substrates.

2.2. Heme States and Redox Properties of Myeloperoxidase

9In resting MPO, the heme iron is in the ferric state (Fe3+). During activation, other heme states can result known as Compound I, Compound II, and Compound III (Table 1). Compounds I and II contain ferryl iron with an attached oxygen. In Compound I of MPO, the porphyrin ring is additionally modified to a π-cation radical (•+Por). In Compound III, a superoxide anion radical (O2•−) is attached to ferric heme iron. This compound can also be presented as a resonance structure where dioxygen is bound to ferrous heme iron.

10The oxidation states of Compounds I and II are higher by two or one units, respectively, in comparison to resting MPO (Table 1). Standard reduction potentials (E’°) for the interconversion of MPO states (Table 2) have been determined. In biological systems, the conversion of resting MPO into Compound I, which plays a central role in reactions of the halogenation and peroxidase cycles (see below), is possible by the concurrent reduction of hydrogen peroxide to water.

2.3. Reaction Cycles of Myeloperoxidase

11Halogenation cycle: An overview of the major reaction cycles of MPO is given in Figure 1. Compound I of MPO, which is formed as a result of a reaction between resting MPO and H2O2, is a short-lived, very reactive state. It interacts rapidly by abstracting two electrons with (pseudo)halides and oxidizes them to hypohalous acids or hypothiocyanite. In this reaction, resting MPO is formed again. The reaction sequence resting MPO→Compound I→resting MPO comprises the halogenation cycle (see Figure 1). The standard reduction potential of the couple Compound I/resting MPO is known to be 1.16 V at pH 7. The reactivity of Compound I with (pseudo)halides decreases in the order SCN− > I− > Br− > Cl−. This reactivity is also higher in slightly acidic media in contrast to neutral pH values.

12On the basis of second-order rate constants and (pseudo)halide ion concentrations, it has been calculated that Cl− and SCN− are equally oxidized by activated MPO in a milieu that corresponds to pH and ion composition with 0.1 M Cl− and 100 µM SCN− of blood. In secretions characterized by high-micromolar to low-millimolar SCN− concentrations and the presence of the heme protein lactoperoxidase, MPO from recruited neutrophils and lactoperoxidase can together contribute to the prevailing formation of hypothiocyanite.

13Hypochlorous acid as a myeloperoxidase product: In numerous reports, the formation of HOCl and chlorinated products by MPO was measured in the presence of taurine at pH 7.4. Other investigations did not detect chlorinated products in the absence of taurine at this pH. Moreover, investigation of pH dependencies of the reduction potentials of involved redox reactions revealed that the probability of HOCl formation is very low at neutral pH values. Interaction between Compound I and chloride yields a reversible high-spin complex. Taurine is known to react with this complex to yield taurine chloramine and resting MPO without the formation of free HOCl.

14In the cytoplasm of PMNs, the taurine concentration is 22–26 mM. In plasma, taurine concentration is much lower, with about 38–40 µM. In resting PMNs, both MPO and taurine are well separated. The formation of taurine chloramine inside neutrophils should be possible after permeabilization of azurophilic granules or after leakage of phagosomes. It remains unknown whether MPO and taurine can interact together already immediately after the initiation of phagosome formation.

15Numerous substrates are targets for free HOCl, and similarly for free HOBr. The reactivity of OSCN−/HOSCN is more restricted and mainly concerns the oxidation of sulfhydryls. As a result, HOSCN is much better able to penetrate into intact cells than HOCl and HOBr, and to thus exhibit a cytotoxic or bactericidal activity by affecting intracellular glutathione and other critical sulfhydryls. Moreover, tyrosine residues of albumin are brominated by the MPO-H2O2-halide system in the presence of physiological concentrations of chloride and bromide at pH values higher than 7.

16Peroxidase cycle: Compound I is also able to oxidize numerous small substrates in a one-electron reaction under the formation of a substrate radical and reduction to Compound II. Important substrates are urate, tyrosine, tryptophan, sulfhydryls, nitric oxide, polyphenols, nitrite, xenobiotics, hydrogen peroxide, xenobiotics, and others.

17The reduction of Compound II to resting MPO is more restricted due to a lower standard reduction potential for the couple Compound II/resting MPO (0.97 V at pH 7) in contrast to the couple Compound I/Compound II (1.35 V at pH 7)). In tissues, only few substrates are known able to reduce Compound II at a sufficient rate. Among these substrates are polyphenols (such as (−)-epicatechin, luteolin, quercetin, (±)-eriodyctiol, and (+)-taxifolin, urate, ascorbic acid, tyrosine, serotonin, and superoxide anion radicals. The highest rate for Compound II reduction by far was determined for quercetin, followed by (−)-epicatechin. Compound II is unable to oxidize halides.

18The reaction sequence resting MPO→Compound I→Compound II→resting MPO comprises the peroxidase cycle (see Figure 1). In the absence of substrates able to reduce Compound II, the latter compound can accumulate. In summary, the reactions of the halogenating and peroxidase cycles of MPO are driven by substrate availability, reaction rates, and redox properties.

19Other reaction cycles: Compound III is formed in a reaction of ferric MPO with a superoxide anion radical with a rate constant of 2 × 106 M−1s−1 at pH 7.4. This rate constant is one order of magnitude higher than the rate constant for spontaneous dismutation of O2•− at this pH. In a reaction of Compound III with a further O2•−, which precedes with a rate of 1.3 × 105 M−1s−1 at pH 7.4, resting MPO is recovered under the formation of H2O2 and O2. Compound III of MPO is not involved in the oxidation of halides and small substrates.

20With further increasing pH values, all O2•−-dependent reactions of MPO become more likely, as the rate constant of spontaneous dismutation of O2•− decreases steadily with increasing pH. These values are 6 × 104 M−1s−1 at pH 8, 6 × 103 M−1s−1 at pH 9, and 6 × 102 M−1s−1 at pH 10. Thus, MPO may exhibit a weak superoxide dismutase activity at alkaline pH values. In this O2•−-driven reaction sequence, only resting MPO and Compound III are involved (see Figure 1).

21At inflammatory sites, a very rapid reaction between NO and O2•− may occur under the formation of peroxynitrite. This reactive species is known to convert resting MPO directly to Compound II, whereby MPO may act as a sink for peroxynitrite. The additional presence of substrates like O2•− able to reduce Compound II at a sufficient rate can enhance the removal of peroxynitrite as a result of permanent switching between resting MPO and Compound II (see Figure 1).

3.1. Recruitment of PMNs to Inflamed Sites

22Neutrophils circulate in peripheral blood in huge amounts. With their receptors, they are able to sense any changes in properties of the vessel wall that allows these cells to adhere firmly to the endothelium and to permeate into inflamed regions. During and after diapedesis, this directed movement is driven by chemical gradients of cytokines and chemotactic agents such as interleukin-8, interferon γ, complement factors C3a and C5a, fMet-Leu-Phe, and leukotriene B4.

23In resting polymorphonuclear leukocytes, myeloperoxidase is stored in the so-called azurophilic granules. Other types of granules of these cells are secretory, tertiary, and specific granules. These granule types differ in their internal and membranous composition of active agents and dependence on cytoplasmic calcium levels to induce degranulation. During the recruitment of neutrophils to inflamed tissues, these cells are step-by-step activated with the release of granule contents into the cell environment or into newly formed phagosomes containing microbes of fungi. The released granule components from secretory, tertiary and, to some extent, specific granules help to digest the surrounding connective tissue to facilitate tissue invasion of these cells that allow a directed movement of invading cells to the inflamed area, where foreign microorganisms are phagocytosed.

3.2. Important Components of Azurophilic and Specific Granules of Neutrophils

24Components of specific and especially azurophilic granules are mostly involved in deactivation and killing of ingested microbes and fungi. They discharge their contents into the newly formed phagosomes. Azurophilic granules deliver, besides myeloperoxidase, serine proteases, bactericidal/permeability-increasing protein, defensins, lysozyme, azurocidin, and others. The main components of specific granules are lysozyme, lactoferrin, some serine proteinases, histaminase, type IV collagenases, gelatinase, and others. In the formed phagosomes, the ingested microbes and fungi are surrounded by a small fluid volume containing highly concentrated material discharged from fused granules.

25Granule components exhibit different functions in the formed phagosomes. Major functions of these components in the early phase of phagocytosis are listed in Figure 2. Some of them are known to interact with surface structures of ingested microorganisms, and thus contribute to the inactivation and killing of pathogens. Prominent examples are defensins, bactericidal/permeability-increasing protein, and azurocidin. Lactoferrin exerts also an antibacterial activity by interaction with outer bacterial membranes. Additionally, this protein sequesters free iron ions. Another group of agents has a pH optimum around 8–9 such as the serine proteases elastase, cathepsin G, and proteinase 3 as well as the hydrolase lysozyme. Neutrophil collagenase exhibits maximal activities between pH 6 and 9.5. The pH-dependence of enzymatic activity of PMN gelatinase is more bell-shaped, with a maximum between pH 7.5 and 8. As shown before, myeloperoxidase needs to be activated by reactive species such as H2O2, or O2•−, or peroxynitrite. This enzyme is principally active under alkaline, neutral, and acidic conditions. However, the chlorination and peroxidase activity of MPO is optimal at pH 5–6.

3.3. Conditions of Phagosomal Digestion

26Components of fused granules participate in the creation of a special milieu in phagosomes, which facilitates the inactivation of microorganisms. At the beginning of phagocytosis, NADPH oxidase is assembled from cytoplasmic and membranous components. This transmembrane protein generates a large amount of superoxide anion radicals (O2•−) by the oxidation of cytoplasmic NADPH, transfer of electrons through the phagosomal membrane, and one-electron reduction of O2 in the phagosomal space. Dismutation of O2•− yields hydrogen peroxide and dioxygen. The enhanced formation of O2•−, opening of ion channels, and altered ion fluxes through the phagosomal membrane contribute to a rapid increase in pH in newly formed phagosomes.

27Originally, it was assumed that the phagosomal pH increases rapidly to about 7.8–8.0 (during the first three minutes), and decreases gradually to 7.0 (after 10–15 min) and later to 6.0 (after about 1 h). The application of a better suitable pH indicator and the removal of azide from the incubation medium revealed a much higher and more long-lasting pH increase to a mean value of 9.0 (in few cases to pH 10.2) in human PMNs incubated with deactivated Candida albicans. Concomitantly with the rise in phagosomal pH in human PMNs, the cytoplasmic pH drops slightly from 7.56 to a medium value of 7.3.

28The significance of NADPH oxidase in an increase in phagosomal pH is demonstrated by the use of the NADPH oxidase inhibitor diphenylene iodonium (DPI) or by the application of NADPH oxidase-deficient neutrophils. Under these conditions, the phagosomal pH drops down to about 6.3 and slightly lower.

29In newly formed phagosomes, an alkaline milieu is important for the rapid and efficient inactivation of phagocytosed microorganisms. Numerous bactericidal constituents of azurophilic and specific granules have a pH optimum around 8–9, such as elastase, cathepsin G, proteinase 3, and lysozyme. Neutrophil collagenase and gelatinase also exhibit significant activities in this pH region (Figure 2). Thus, a cocktail of aggressive proteins, bactericidal proteins and reactive species is present in the formed phagosomes. The concerted action of these agents promotes the deactivation, killing, and digestion of the phagocytosed microorganisms.

3.4. Potential Role of Myeloperoxidase in Phagosomes

30In azurophilic granules of resting neutrophils, MPO as well as other cationic protein components are probably inactivated by sequestration with negatively charged proteoglycans and the presence of low pH. These conditions are changed when azurophilic granules discharge their content into the newly formed phagosomes of activated PMNs.

31Immediately after phagosome formation, the pH rises to alkaline values as a result of NADPH oxidase activation and compensatory ion fluxes. The consumption of H+ ions in reactions of the formed O2•− is responsible for this pH increase. During the early phase of phagocytosis, discharged MPO can likely act as a weak superoxide dismutase, considerably enhancing the removal of O2•−. In one dismutase cycle, two protons are consumed.

32Moreover, the high yield and stability of O2•− promotes also its reaction with nitrogen monoxide (NO), an agent that is freely diffusible. The reaction product of O2•− and NO is peroxynitrite (ONOO−). The protonated form of ONOO−, peroxynitrous acid ONOOH, is known to convert resting MPO directly into Compound II. As O2•− efficiently recovers resting MPO from Compound II, peroxynitrite removal by MPO can drive a permanent cycling between resting MPO and Compound II. Two protons are consumed during one cycle. The production of peroxynitrite also contributes to microbicidal activity in phagosomes.

33Superoxide anion radicals also interact with sulfur–iron-clusters under the release of Fe2+. This iron can be scavenged by lactoferrin.

34At alkaline pH values, MPO is far from the optimum for efficient halogenating and peroxidase activities. Under these conditions, MPO is unable to oxidize chloride at sufficient rate. However, it cannot be excluded that hydrogen peroxide favors Compound I formation and the reaction of Compound I with traces of SCN− under formation of −OSCN in the halogenation cycle or the oxidation of O2•− to O2 in the peroxidase cycle.

35Later, the phagosomal pH decreases step by step, reaching neutral values. It remains unsolved whether some phagosomes become leaky during phagocytosis with changes of pH to cytoplasmic level. Under these conditions, the dominance of O2•− is diminished, and the halogenating and peroxidase activity of MPO activity rises slowly. It remains unsolved whether acidic pH values can be achieved in phagosomes of neutrophils.

36We are far from a thorough understanding of the role of MPO during the early phase of phagocytosis in PMNs. The main reason for this is the lack of sufficient data about MPO reactions in the pH region 8–10, typical of early phagosomal pH values. It is quite evident that under these conditions, most reactions of halogenation and peroxidase cycles do not work or only work to a limited degree. Otherwise, superoxide anion radicals are stable enough due to the low rate of spontaneous dismutation and can drive now O2•−-dependent reactions of MPO. An enhanced catalase activity of MPO was also proposed in the pH region 9–10. However, a mechanism for this reactivity was not given. Compound III is indeed involved in a catalase-like removal of H2O2 by the sequential conversion: Compound III→ferrous MPO→Compound II→Compound III. As the H2O2-driven conversion of Compound II to Compound III is the rate-determining step, this cycle can explain the observed accumulation of Compound II at alkaline pH.

37The application of different inhibitors revealed cooperative effects of superoxide anion radicals and a myeloperoxidase-dependent pathway in the killing of Staphylococcus aureus by neutrophils. Despite the strong alkalization of early phagosomes in human neutrophils, the authors excluded a sufficient participation of MPO in these reactions, as the MPO inhibitors 4-aminobenzoic acid hydrazide (4-ABAH) and KCN failed to affect the phagosomal pH. The inhibitor 4-ABAH is known to interact with Compound I, which is apparently not involved in O2•−-driven MPO reactions under alkaline conditions. Hydrocyanic acid (HCN) forms a complex with resting MPO, whereby HCN is deprotonated by the interaction with distal histidine. This binding is optimal between pH values 5 and 8. At pH > 8, the apparent second-order rate constant for the complex formation between MPO and HCN decreases with increasing pH due to the pKa value of 9.2 for HCN. Considering the additional presence of O2•−, which can compete with HCN for ferric MPO, it is questionable whether KCN is able to inhibit MPO at alkaline pH values in the phagosome. Azide, another MPO inhibitor, caused a significant decrease in phagosomal pH indicating an involvement of MPO in phagosome alkalinization.

38In vitro experiments demonstrate the killing of microbes by the MPO-H2O2-halide system and inhibition of killing by MPO inhibitors or by the application of neutrophils from MPO-deficient individuals. From these data, it was concluded that halogenating MPO is critical for the inactivation of microbes. These experiments are usually performed at a low protein load, at neutral pH values, and in the absence of competing substrates, conditions that are far from the actual situation in phagosomes. Product analysis of phagocytosed material of PMNs revealed that most other released granule components were halogenated but not constituents from ingested microbes. Under these conditions, bacteria are apparently not killed by halogenated MPO products.

39Other data support an involvement of the halogenating activity of MPO in the termination of PMN serine proteases in phagosomes and protection of pericellular tissues from unwanted reactions. For example, the MPO product HOCl is known to inactivate elastase, cathepsin G, proteinase 3, and matrix metalloproteinase 9.

40In summary, in the early phase of phagocytosis in neutrophils, MPO apparently helps to intensify and prolong the duration of an alkaline milieu by activities mainly based on the consumption of O2•−. Later, MPO can probably contribute with its halogenation and peroxidase activity to the termination of phagocytotic activities and protection of surrounding tissues from uncontrolled proteolysis.

3.5. Redundancy in Deactivation and Killing Pathways and MPO Deficiency

41In activated neutrophils, a broad range of active agents ensures protection against different types of microbes and fungi. There are multiple deactivation and killing pathways and a redundancy exists between these pathways.

42For example, in humans, total and subtotal MPO-deficiency occurs with an abundance of 1 case on 2000–4000 individuals. As opposed to patients with an NADPH oxidase deficiency, which develop chronic granulomatous disease and suffer from multiple inflammatory events, MPO-deficient persons live normally. Only in few cases were persistent infections with Candida albicans reported.

43Unlike humans, serious complications were reported in investigations of different disease models using MPO-knockout mice. A careful analysis revealed a protective role of MPO in pathologies characterized by the infiltration of T-cells. MPO oxidants are involved in the suppression of lymphocyte functions. Otherwise, in disease models, where an innate immune response dominates exclusively, MPO knockout protects tissue from damage.

44In MPO-knockout mice, no MPO protein is present. Human MPO deficiency is mainly assessed by diminished or missing peroxidase activity, a property that is more pronounced under slightly acidic conditions and not in the early phase of phagocytosis. It remains unsolved whether MPO from MPO-deficient patients exhibits a superoxide dismutase-like activity, which can be involved in the alkalization of newly formed phagosomes.

45Moreover, human and murine neutrophils differ considerably in their properties. Mouse neutrophils do not contain any defensins, small cationic proteins that induce defects in microbial membranes. In azurophilic granules of human PMNs, four types of defensins are present. Other differences concern the lower expression of bactericidal/permeability-increasing protein, lysozyme, and β-glucuronidase in mouse neutrophils. The same holds for MPO, with a level of 10–20% in murine compared to human cells. A further difference concerns the degree and duration of the pH increase during the early phase in phagosomes. In murine neutrophils incubated with Candida albicans, the pH reaches with 8.5 a lower mean maximum value and decreases faster than in human cells. It remains unknown to what extent the difference in MPO expression in both species contributes to the more pronounced increase in pH in phagosomes of human neutrophils. A higher amount of MPO in human cells can more strongly enhance the dismutation of O2•− and can utilize more H2O2.

46These data support a higher degree of redundancy in deactivation and killing pathways in human neutrophils than in murine ones.

3.6. Cell Death of Neutrophils and Formation of Extracellular Traps

47Activated neutrophils are known to undergo apoptosis. Typical features of apoptotic cells are the appearance of phosphatidylserine epitopes on the cell surface by concurrent intactness of the plasma membrane. In necrotic cells, the permeability of the plasma membrane increases. At inflammatory sites, apoptotic and necrotic PMNs are recognized and ingested by macrophages that are attracted to these sites, usually time-delayed to PMNs. In both apoptotic and necrotic PMNs, catalytically active myeloperoxidase has been found attached to phosphatidylserine epitopes.

48In dying PMNs, so-called neutrophil extracellular traps (NETs) are formed and released. These traps represent a network of extracellular DNA-derived fibers, to which MPO, elastase, cathepsin G, gelatinase, lactoferrin, calprotectin and some others are attached. Extruded traps can bind and kill microbes independent of phagocytosis. It is discussed that NETs are important in host defense against fungal pathogens that are difficult to phagocytose. In blood, NETs are able to promote coagulation, vascular occlusion, thrombosis, sequestration of circulating tumor cells, and metastasis. In addition, both pro- and anti-inflammatory activities of NETs are reported. NETs are internalized and degraded by macrophages.

49Myeloperoxidase is required for NETs formation, as PMNs from patients completely deficient in MPO cannot form traps. Hydrogen peroxide is known to enhance the enzymatic activity of MPO in traps and the ability of NETs to kill microbes.

3.7. Frustrated Phagocytosis

50Besides necrotic cell death, aggressive agents from activated PMNs can also be released into the surrounding medium as a result of an incomplete, the so-called frustrated, phagocytosis. In this case, parts of specific and azurophilic granules fuse with the plasma membrane and discharge their content into external space. This may occur in premature or overstimulated neutrophils, at extended inflammatory loci, or as a result of a high load of pathogens. In this way, MPO, serine proteases, and other granule constituents are released from neutrophils, and can interact with host cells and tissues.

3.8. Degradation of Ingested Material by Macrophages

51At inflammatory sites, tissue-resident and monocyte-derived macrophages engulf and digest waste material and undergoing cells including apoptotic and necrotic neutrophils. It has been assumed that macrophages can modulate their activity depending on the dominance of apoptotic and necrotic cell material. An inflammation persists as long as constituents released from necrotic cells contribute to further cell and tissue damage. Under these conditions, activated macrophages mainly release proinflammatory cytokines. For example, some components from necrotic PMNs such as elastase, cathepsin G, and heat shock proteins are known to modify surface molecules of adjacent macrophages, and thus induce pro-inflammatory pathways in the latter ones. A switch to the resolution of inflammation and the release of mainly anti-inflammatory cytokines occurs when macrophages digest predominantly apoptotic cell material.

52An important aspect of the interaction between apoptotic PMNs and macrophages is the rapid recognition of apoptotic cells by macrophages. Their delayed phagocytosis by macrophages can result in secondary necrotic processes.

53Interestingly, in classically activated macrophages (type 1 macrophages) from human origin, the phagosomal pH rises to about 8.5 within five minutes, displays some oscillations of alkalinization, and remains nearly unchanged or returns to neutral values during the next 25 min. This pH increase depends on NADPH oxidase activity, like in phagocytosing neutrophils. Contrary to this, alternatively activated macrophages (type 2 macrophages) monotonically acidify their phagosomes to pH 5.0 within 10 min or to pH 5.5 within 30 min without any participation of NADPH oxidase. Intriguingly, in murine type 1 macrophages, which express unlike human cells nitric oxide synthase, the phagosomes are also acidified. Apparently, the very rapid reaction between NO and O2•− diminishes the availability of O2•− in these cells and limits, thus, the consumption of H+ by O2•−-driven reactions.

54The presence of MPO on surface epitopes of non-vital neutrophils and other cells implies the question of whether attached MPO plays a role following phagocytosis of undergoing cells by macrophages. At present, a clear answer on this subject cannot be given. Probably, MPO can play a similar role in alkalinization of the phagosome in classically activated macrophages, as assumed in phagocytosing neutrophils. The strong pH decrease in phagosomes of alternatively activated macrophages would, however, favor better conditions for the chlorinating and peroxidase activity of MPO.

4.1. The Fate of Myeloperoxidase at Inflammatory Sites

55Myeloperoxidase can be released from PMNs at inflammatory sites as a result of incomplete phagocytosis, cell necrosis, or as component of NETs. The cationic nature of MPO favors an interaction with numerous negatively charged serum proteins, endothelial cells, and components of the extracellular matrix. The activation of MPO by hydrogen peroxide causes halogenation and oxidation reactions in critical residues of the attached targets. On the basis of these interrelations, an involvement of MPO in several pathologies is discussed.

4.2. Important Binding Sites for Myeloperoxidase

56Myeloperoxidase is known to interact with numerous plasma proteins. Critical residues in albumin, α1-antiproteinase (also known as α1-antitrypsin), apolipoprotein A1, and soluble plasma fibronectin are modified by the interaction of MPO with these targets.

57Myeloperoxidase binds to heparin/heparan-containing epitopes of endothelial membranes. Complexes between albumin and MPO are also known to interact with the endothelium. Further, MPO is internalized by endothelial cells, rapidly transcytoses the intact endothelium, and associates closely with fibronectin. In an inflammatory model, MPO diminishes the bioavailability of NO and impairs the NO-dependent vessel relaxation.

58In the extracellular matrix and coronary smooth muscle cells, MPO is known to bind to collagen IV and fibronectin. Other targets of MPO are perlecan, laminin, and glycocalyx glycosaminoglycans.

4.3. The Protective Role of Ceruloplasmin

59However, the incubation of purified MPO with plasma revealed ceruloplasmin as the major binding target for MPO. In these experiments, co-elution between MPO and complement C3 was also observed. All other reported in vitro associations of MPO with serum proteins have likely only minor physiological significance.

60The copper-containing ceruloplasmin is a serum protein with multiple anti-inflammatory and antioxidant functions. It oxidizes Fe2+ and Cu+. Ceruloplasmin forms high-affinity complexes with myeloperoxidase and other cationic proteins of neutrophils such as lactoferrin and serine proteases. Its complex with lactoferrin facilitates the binding of the resulting Fe3+ and the transfer to the iron-binding protein transferrin. A superoxide dismutase activity of ceruloplasmin is also reported.

61The complex formation between MPO and ceruloplasmin strongly inhibits the halogenating and peroxidase activity of MPO. In the complex between MPO and ceruloplasmin, MPO is converted to Compound II and held in this inactive form until ceruloplasmin dissociates from MPO. Inhibition of MPO activities is favored by a loop of the ceruloplasmin chain that penetrates into the substrate channel and blocks the access to the active site of MPO. Partial proteolysis of ceruloplasmin by elastase, plasmin, or trypsin dampens this inhibition. Antibodies against MPO can prevent the binding between MPO and ceruloplasmin by steric hindrance. Hence, this interaction with antibodies favors the presence of active MPO in patients with renal vasculitis.

4.4. Myeloperoxidase in Atherosclerotic Plaques

62Atherosclerotic plaques limit vessel lumen, disturb blood circulation, and cause serious health problems for the affected patients. In plaque formation, lipoproteins, endothelial cells, immune cells, smooth muscle cells, and others are involved. Oxidative modifications to lipoproteins, extracellular matrix components, and cell surface elements also contribute to the formation of atherosclerotic lesions. Despite intense research, it remains puzzling how atherosclerosis is initiated in vivo and how MPO participates in this pathogenesis.

63In atherosclerotic lesions, active MPO was detected. Chlorinated and nitrated products such as 3-chlorotyrosine, 3-chlorouracil and 3-nitrotyrosine were found in plaque proteins as well. While chlorinated products are specific to MPO and associated with the MPO-mediated formation of HOCl, nitration can also result from peroxynitrite-driven reactions. In atherosclerotic lesions, numerous components of the extracellular matrix are oxidatively modified, such as fibronectin, laminin, and others. Again, the involvement of MPO is discussed. Moreover, there is an increased affinity of MPO to oxidized material in the extracellular matrix. This positive feedback can further promote oxidative matrix modifications in developing atherosclerotic lesions.

64It remains unclear whether the interaction of MPO with lipoproteins and endothelium components is the initial reason for plaque formation, or whether the presence of MPO and MPO products in plaques results from subsequent processes like the infiltration of PMNs into inflamed areas.

65In addition, several further MPO-related mechanisms are assumed to play a potential role in plaque development. Myeloperoxidase-derived HOCl is known to oxidize low-density lipoproteins and convert them into a pro-atherogenic species. The interaction of MPO with apolipoprotein A1, the major protein in high-density lipoproteins (HDLs), limits the ability of HDLs to remove cholesterol from lipid-laden cells. The formation of cyanate as a result of SCN− or cyanide oxidation by MPO leads to the carbamylation of lysine residues in proteins, a process that has been associated with foam cell formation. Myeloperoxidase is involved by several routes in protein carbamylation in atherosclerotic lesions. Enhanced MPO plasma levels probably decrease the bioavailability of nitric oxide, and can thus promote endothelial dysfunction. Another consequence of MPO activity is the enhanced activity of matrix metalloproteinases. It should be noted that there are other pathways independent of MPO leading to LDL oxidation, diminished bioavailability of NO, disturbed endothelial dysfunctions, or protein carbamylation.

4.5. Myeloperoxidase and Cardiovascular Diseases

66There are several reports specifying a relationship between enhanced MPO plasma levels and the development of cardio-vascular problems. Again, the question arises of whether the enhanced MPO level is the origin of these problems or the consequence of the additional recruitment of neutrophils to inflamed endothelium in these patients.

67Rupture of atherosclerotic plaques is a major reason for acute cardiovascular disease. Among critical factors in plaque formation, the infiltration of neutrophils and macrophages contributes to weakening of the fibrous cap, a process that precedes plaque rupture. Determination of MPO plasma levels has been used as a marker for the predictive outcome of cardiovascular problems. The majority of studies demonstrated a relationship between elevated plasma MPO levels and the presence of coronary artery disease.

4.6. Vasculitis Induced by Antineutrophil Cytoplasmic Antibodies

68Antineutrophil cytoplasmic antibodies (ANCA) are involved in ANCA-associated vasculitis, which represents a small vessel vasculitis. There are mainly antibodies against myeloperoxidase and, to a lesser extent, against proteinase 3. Both proteins are present in a high load in neutrophils and are known to be attached to the surface of dying neutrophils. It cannot be excluded that these proteins are also be found on surface areas of other undergoing cells at inflammatory sites and, hence, are presented as antigens by dendritic cells.

69The development of MPO-ANCA and other ANCA is poorly understood. Among the postulated mechanisms of ANCA generation are defective apoptosis of neutrophils and the hypothesis of molecular mimicry.

70Once formed, ANCA can recognize over-activated neutrophils, which are attached to inflamed vessel wall areas. These attacks further promote the inflammatory process, leading to a necrotizing vasculitis. Glomerulonephritis, vasculitis of the upper and lower respiratory tract, and diabetic retinopathy are prominent forms of ANCA-associated vasculitis.

71In the pathogenesis of different types of vasculitis, a relationship between MPO and α1-antiproteinase is discussed. Myeloperoxidase products oxidize critical methionine residues in this antiproteinase. As a result, the ability of α1-antiproteinase is limited to inactivate elastase and proteinase 3 at inflammatory sites and a partial protease-mediated hydrolysis of ceruloplasmin may occur. Partially proteolyzed ceruloplasmin is unable to inhibit MPO. Collectively, the imbalance between the protease–antiproteinase activities can favor a prolonged activity of myeloperoxidase and proteinase 3 and the formation of antibodies against both agents.

4.7. Involvement of Myeloperoxidase in Other Disease Scenarios

72In many diseases, the infiltration of neutrophils into inflamed tissue regions is closely associated with increased MPO levels. Active MPO was detected in tissue sections of patients with pancreatitis, in periodontitis, and in synovial fluid of patients with rheumatoid arthritis. The involvement of MPO is also discussed in the pathogenesis of Alzheimer’s disease, Parkinson’s disease, and multiple sclerosis. Further examples of the potential involvement of MPO in disease pathogenesis are obesity, sinusitis, cystic fibrosis, inflammatory bowel disease, renal, and liver diseases.

5.1. Chronic Inflammatory Processes

73Many disease scenarios are accompanied by chronic inflammatory processes, in which the inflammation is only insufficiently terminated. In inflamed tissues, there is a mixture of unperturbed tissue regions with necrotic areas, and regions where a de novo reconstitution of tissues occurs. Generally, it is impossible to give a clear description of chronically affected tissues as the appearance of these regions varies enormously. There are also flashes of the active inflammation with silent phases.

74In the termination of inflammation, a switch occurs from a pro-inflammatory phase to resolution of inflammation followed by restoration of tissue homeostasis. In chronic inflammations, damage-associated molecular patterns (DAMPs) released from necrotic cells can repeatedly foment the inflammatory process. A crucial aspect for chronicity is the balance between the damage to host’s tissues by agents from activated immune cells and undergoing tissue cells and the ability of host’s tissues to resist and inactivate these destructive agents. There is a wide range of variability of protective mechanisms from one patient to another. The low-tissue capacity of antagonizing agents generally prolongs inflammation, favors its chronic conversion, and provides the basis for disease progression. Thus, the preponderance of damaging over protective mechanisms disturbs the termination of inflammation and favors long-lasting inflammatory events.

5.2. Protection of Surrounding Media Against Damaging Agents

75In human blood and tissues, numerous mechanisms and strategies exist to protect biological material at inflammatory sites from damage caused by destroying agents released from activated neutrophils, other immune cells, and damaged tissue cells. Known examples for the latter agents are free hemoglobin, and free myoglobin, which are released from red blood cells during intravascular hemolysis, and muscle cells as a result of rhabdomyolysis, respectively. Both free hemoglobin and free myoglobin are the source of the very cytotoxic agent ferric protoporphyrin IX (free heme).

76Intact cells and tissues are generally protected by the presence of molecules, which deactivate cytotoxic agents by binding, inactivation, or degradation. Examples of these protective interactions are given in Table 3. It should be noted that these mechanisms are mostly directed against frequently occurring damaging agents.

77Some of the listed protecting species are acute phase proteins (such as ceruloplasmin, α1-antiproteinase, α1-antichymotrypsin, and haptoglobin), which can be up-regulated to a certain degree during the course of inflammation. Otherwise, at prolonged and severe inflammatory events, a decrease in or exhaustion of distinct acute phase proteins and other protective agents can be observed.

78Released myeloperoxidase is inactivated by complex formation with ceruloplasmin, which is a late acute-phase protein and reaches its maximal plasma level after most other acute-phase proteins. The question arises of whether this interaction with ceruloplasmin is sufficient to inactivate all released MPO under severe inflammatory conditions.

79As shown in Table 3, protective mechanisms are very broad-ranged and comprise numerous anti-proteinases, control over superoxide anion radicals, hydrogen peroxide, and free metal ions, the presence of lipid- and water-soluble antioxidants and agents, to detoxify free heme and free heme proteins. These immediately acting, ready-to-use protective mechanisms are completed by inducible processes that enhance the protective power of cells, and by formation of antibodies against antigens.

80Of the antagonizing agents against neutrophil products, α1-antiproteinase deserves special attention. This acute phase protein is produced in the liver and protects the lungs and other tissues from neutrophil elastase. Individuals with α1-antiproteinase deficiency may develop chronic obstructive pulmonary disease, chronic liver disease, skin inflammations, ANCA-related vasculitis, glomerulonephritis, and Bowel disease. The protease-anti-proteinase imbalance can also affect the ability of ceruloplasmin to bind and inactivate MPO.

5.3. Immunosuppression

81During the resolution of inflammation, transient immunosuppression is important to allow final apoptosis of immune cells, to restore the normal tissue homeostasis, to terminate all inflammatory processes, and to induce, if necessary, de novo formation of tissues. Many chronic disease processes are accompanied by pronounced immunocompromised states of patients. This mostly concerns persons of advanced age, who suffer from different comorbidities. However, long-lasting immunosuppression depresses general immune functions, and favors the appearance of opportunistic infections.

5.4. Sepsis

82Strong deviations of normal tissue homeostasis can result in sepsis. According to the last consensus definition in 2016, sepsis is, at present, defined as life-threatening organ dysfunction caused by a dysregulated host response to infection. Septic shock is determined as a subset of sepsis, in which the underlying circulatory and cellular/metabolic abnormalities are profound enough to substantially increase mortality. Sepsis is characterized by malfunction of one or several organs such as liver, kidneys, or lungs. It occurs most frequently in immunocompromised individuals.

83Immunosuppression impacts PMN functions in septic patients. These cells show a delayed apoptosis and diminished chemotactic mobility. With these properties, PMNs can contribute to tissue damage distant from inflammatory and infection sites. In addition, an increased percentage of immature neutrophils is found in septic patients.

84In sepsis, tissue-damaging processes can vary widely and depend largely on the state of individual protective mechanisms. Besides invading microorganisms, products of neutrophils including MPO, and products of other immune and tissue cells, can be involved in cell and tissue damage in septic patients. Furthermore, antigens and DAMPs released form necrotic tissue cells also play a role in the recruitment and activation of immune cells. The considerable decline or exhaustion of some protective mechanisms also worsens the recovery of unperturbed tissue homeostasis.

85In septic shock patients, plasma levels of MPO–DNA complexes, which result from neutrophil extracellular traps, are significantly enhanced in contrast to healthy volunteers and closely related to the severity of organ dysfunction. Patients with sepsis or with septic shock show higher plasma levels of MPO in comparison to patients with systemic inflammatory response syndrome without infection. In addition, an association between higher MPO levels and increased mortality was found. Enhanced MPO values were also detected in other studies with septic patients.

6. Conclusions

86The fate of myeloperoxidase reflects the general role of neutrophils in immune response. On the one hand, neutrophils and neutrophil constituents are highly essential to successfully combat foreign bacteria and other pathogens. In the phagosomes of PMNs, myeloperoxidase is involved in the creation and maintenance of an alkaline milieu, which is optimal for the activity of serine proteases and other granule components in the deactivation and killing of microbes. Halogenation and peroxidative activities of MPO are apparently involved in later phases of phagocytosis of pathogens and can play a role in the control and termination of phagocytic activities in neutrophils, and maybe also in macrophages, after the ingestion of undergoing PMNs. In addition, MPO is a mandatory element for the formation of neutrophil extracellular traps.

87On the other hand, the release of PMNs constituents during frustrated phagocytosis or from necrotic PMNs can damage intact host tissues. Myeloperoxidase as a major constituent of PMNs contributing to this damage by its close association with negatively charged components of plasma and extracellular matrix and by the chemical modification of these targets. The acute phase protein ceruloplasmin binds and inactivates MPO. The ability of MPO to damage molecules and tissues can be enhanced at inflammatory sites by the massive release of MPO from PMNs and the imbalance between proteases and anti-proteinases. In several disease scenarios and sepsis, the involvement of MPO in the pathological process is discussed. It remains unknown whether MPO contributes to basic mechanisms of disease induction or whether the increased MPO level in affected tissues results from downstream effects due to the recruitment of neutrophils to inflammatory sites.

88In sum, MPO exhibits, like neutrophils and many other components of the immune system, both a protective and harmful role in the maintenance and disturbance of tissue homeostasis.

References 294

1.
Arnhold J. Immune response and tissue damage. Cell and Tissue Destruction. Mechanisms, Protection, Disorders. 2020;155-204.
Not in our library
2.
Arnhold J. Acute-phase proteins and additional protective systems. Cell and Tissue Destruction. mechanisms, Protection, Disorders. 2020;205-228.
Not in our library
3.
Muller W.A. Leukocyte-endothelial interactions in the inflammatory response. Lab. Investig. 2002;82:521-533.
Not in our library10.1038/labinvest.3780446
4.
Taylor K.R., Gallo R.I. Glycosaminoglycans and their proteoglycans: Host-associated molecular pattern for initiation and modulation of inflammation. FASEB J. 2002;20:9-22.
Not in our library10.1096/fj.05-4682rev
5.
Allen R.C., Stevens D.L. The circulating phagocyte reflects the in vivo state of immune response. Curr. Opin. Infect. Dis. 2006;5:389-398.
Not in our library10.1097/00001432-199206000-00013
6.
Klebanoff S.J. Myeloperoxidase: Friend and foe. J. Leukoc. Biol. 2005;77:598-625.
Not in our library10.1189/jlb.1204697
7.
Arnhold J., Flemmig J. Human myeloperoxidase in innate and acquired immunity. Arch. Biochem. Biophys. 2010;500:92-106.
Not in our library10.1016/j.abb.2010.04.008
8.
Bos A., Wever R., Roos D. Characterization and quantification of the peroxidase in human monocytes. Biochim. Biophys. Acta. 1978;525:37-44.
Not in our library10.1016/0005-2744(78)90197-3
9.
van der Veen B.S., de Winther M.P., Heeringa P. Myeloperoxidase: Molecular mechanisms and their relevance to human health and disease. Antioxid. Redox Signal. 2009;11:2899-2937.
Not in our library10.1089/ars.2009.2538
10.
Owen C.A., Campbell M.A., Boukedes S.S., Stockley R.A., Campbell E.J. A discrete subpopulation of human monocytes expresses a neutrophil-like proinflammatory (P) phenotype. Am. J. Physiol. 1994;267:L775-L785.
Not in our library10.1152/ajplung.1994.267.6.l775
11.
Nagra R.M., Becher B., Trottellotte W.W., Antel J.P., Gold D., Paladino T., Smith R.A., Nelson J.R., Reynolds W.F. Immunohistochemical and genetic evidence of myeloperoxidase involvement in multiple sclerosis. J. Neuroimmunol. 1997;78:97-107.
Not in our library10.1016/s0165-5728(97)00089-1
12.
Sugiyama S., Okada Y., Suhkova G.K., Virmani R., Heinecke J.W., Libby P. Macrophage myeloperoxidase regulation by granulocyte macrophage colony-stimulating factor in human atherosclerosis and implications in acute coronary syndromes. Am. J. Pathol. 2001;158:879-891.
Not in our library10.1016/s0002-9440(10)64036-9
13.
Neutrophil-Mediated Regulation of Innate and Adaptive Immunity: The Role of Myeloperoxidase · 2016
No notice
14.
Shepherd V.L., Hoidal J.R. Clearance of neutrophil-derived myeloperoxidase by the macrophage mannose receptor. Am. J. Respir. Cell Mol. Biol. 1990;2:335-340.
Not in our library10.1165/ajrcmb/2.4.335
15.
Lazzaretto B., Fadeel B. Intra- and extracellular degradation of neutrophil extracellular traps by macrophages and dendritic cells. J. Immunol. 2019;203:2276-2290.
Not in our library10.4049/jimmunol.1800159
16.
Furtmüller P.G., Zederbauer M., Jantschko W., Helm J., Bogner M., Jakopitsch C., Obinger C. Active site structure and catalytic mechanisms of human peroxidases. Arch. Biochem. Biophys. 2006;445:199-213.
Not in our library10.1016/j.abb.2005.09.017
17.
Klebanoff S.J., Kettle A.J., Rosen H., Winterbourn C.C., Nauseef W.M. Myeloperoxidase: A front-line defender against phagocytosed microorganisms. J. Leukoc. Biol. 2013;93:185-198.
Not in our library10.1189/jlb.0712349
18.
Andrews P.C., Parnes C., Krinsky N.I. Comparison of myeloperoxidase and hemi-myeloperoxidase with respect to catalysis, regulation, and bactericidal activity. Arch. Biochem. Biophys. 1984;228:439-442.
Not in our library10.1016/0003-9861(84)90008-0
19.
Fiedler T.J., Davey C.A., Fenna R.E. X-ray crystal structure and characterization of halide binding sites of human myeloperoxidase at 1.8 Å resolution. J. Biol. Chem. 2000;275:11964-11971.
Not in our library10.1074/jbc.275.16.11964
20.
Dolphin D., Forman A., Borg D.C., Fajer J., Felton R.H. Compound I of catalase and horseradish peroxidase: π-cation radicals. Proc. Natl. Acad. Sci. USA. 1971;68:614-618.
Not in our library10.1073/pnas.68.3.614
21.
Odajima T., Yamazaki I. Myeloperoxidase of the leukocytes of normal blood: III. The reaction of ferric myeloperoxidase with superoxide anion. Biochim. Biophys. Acta. 1972;284:355-359.
Not in our library10.1016/0005-2744(72)90130-1
22.
Arnhold J., Furtmüller P.G., Obinger C. Redox properties of myeloperoxidase. Redox Rep. 2003;8:179-186.
Not in our library10.1179/135100003225002664
23.
Arnhold J., Monzani E., Furtmüller P.G., Zederbauer M., Casella L., Obinger C. Kinetics and thermodynamics of halide and nitrite oxidation by mammalian heme peroxidases. Eur. J. Inorg. Chem. 2006;3801-3811.
Not in our library10.1002/ejic.200600436
24.
Arnhold J., Furtmüller P.G., Regelsberger G., Obinger C. Redox properties of the couple compound I/native enzyme of myeloperoxidase and eosinophil peroxidase. Eur. J. Biochem. 2001;268:5142-5148.
Not in our library10.1046/j.0014-2956.2001.02449.x
25.
Furtmüller P.G., Arnhold J., Jantschko W., Pichler H., Obinger C. Redox properties of the couples compound I/compound II and compound II/native enzyme of human myeloperoxidase. Biochem. Biophys. Res. Commun. 2003;301:551-557.
Not in our library10.1016/s0006-291x(02)03075-9
26.
Furtmüller P.G., Burner U., Obinger C. Reaction of myeloperoxidase compound I with chloride, bromide, iodide, and thiocyanate. Biochemistry. 1998;37:17923-17930.
Not in our library10.1021/bi9818772
27.
van Dalen C.J., Whitehouse M.W., Winterbourn C.C., Kettle A.J. Thiocyanate and chloride as competing substrates for myeloperoxidase. Biochem. J. 1997;327:487-492.
Not in our library10.1042/bj3270487
28.
Tenovuo J., Makinen K.K. Concentration of thiocyanate and ionizable iodine in saliva of smokers and nonsmokers. J. Dent. Res. 1976;55:661-663.
Not in our library10.1177/00220345760550042001
29.
Schultz C.P., Ahmed M.K., Dawes C., Mantsch H.H. Thiocyanate levels in human saliva: Quantitation by Fourier transform infrared spectroscopy. Anal. Biochem. 1996;240:7-12.
Not in our library10.1006/abio.1996.0323
30.
Chandler J.D., Day B.J. Thiocyanate: A potential useful therapeutic agent with host defense and antioxidant properties. Biochem. Pharmacol. 2012;84:1381-1387.
Not in our library10.1016/j.bcp.2012.07.029
31.
Chandler J.D., Day B.J. Biochemical mechanisms and therapeutic potential of pseudohalide thiocyanate in human health. Free Radic. Res. 2015;49:695-710.
Not in our library10.3109/10715762.2014.1003372
32.
Flemmig J., Gau J., Schlorke D., Arnhold J. Lactoperoxidase as potential drug target. Expert Opin. Ther. Targets. 2016;20:447-461.
Not in our library10.1517/14728222.2016.1112378
33.
Bakkenist A.R.J., de Boer J.E.G., Plat H., Wever R. The halide complexes of myeloperoxidase and the mechanism of halogenation reactions. Biochim. Biophys. Acta. 1980;613:337-348.
Not in our library10.1016/0005-2744(80)90088-1
34.
Kettle A.J., Winterbourn C.C. Superoxide modulates the activity of myeloperoxidase and optimizes the production of hypochlorous acid. Biochem. J. 1988;252:529-536.
Not in our library10.1042/bj2520529
35.
Zuurbier K.W.M., Bakkenist A.R.J., Wever R., Muijsers A.O. The chlorinating activity of myeloperoxidase: High initial activity at neutral pH and activation by electron donors. Biochim. Biophys. Acta. 1990;1037:140-146.
Not in our library10.1016/0167-4838(90)90159-d
36.
Kettle A.J., Winterbourn C.C. Assays for the chlorination activity of myeloperoxidase. Meth. Enzymol. 1994;233:502-512.
Not in our library10.1016/s0076-6879(94)33056-5
37.
Kettle A.J., Winterbourn C.C. Myeloperoxidase: A key regulator of neutrophil oxidant production. Redox Rep. 1997;3:3-15.
Not in our library10.1080/13510002.1997.11747085
38.
Jerlich A., Horakova L., Fabjan J.S., Giessauf A., Jürgens G., Schaur J.R. Correlation of low-density lipoprotein modification by myeloperoxidase with hypochlorous acid formation. Int. J. Clin. Lab. Res. 1999;29:155-161.
Not in our library10.1007/s005990050083
39.
Panasenko O.M., Spalteholz H., Schiller J., Arnhold J. Myeloperoxidase-induced formation of chlorohydrins and lysophospholipids from unsaturated phosphatidylcholines. Free Radic. Biol. Med. 2003;34:553-562.
Not in our library10.1016/s0891-5849(02)01358-8
40.
Spalteholz H., Panasenko O.M., Arnhold J. Formation of reactive halide species by myeloperoxidase and eosinophil peroxidase. Arch. Biochem. Biophys. 2006;445:225-234.
Not in our library10.1016/j.abb.2005.06.025
41.
Furtmüller P.G., Obinger C., Hsuanyu Y., Dunford H.B. Mechanism of reaction of myeloperoxidase with hydrogen peroxide and chloride ion. Eur. J. Biochem. 2000;267:5858-5864.
Not in our library10.1046/j.1432-1327.2000.01491.x
42.
Marquez L.A., Dunford H.B. Chlorination of taurine by myeloperoxidase: Kinetic evidence for an enzyme-bound intermediate. J. Biol. Chem. 1994;269:7950-7956.
Not in our library
43.
Ramos D.R., Victoria Garcia M., Canle L.M., Santaballa J., Furtmüller P.G., Obinger C. Myeloperoxidase-catalyzed taurine chlorination: Initial versus equilibrium rate. Arch. Biochem. Biophys. 2007;466:221-233.
Not in our library10.1016/j.abb.2007.07.024
44.
Thomas E.L., Grisham M.B., Melton D.F., Jefferson M.M. Evidence for the role of taurine in the in vitro oxidative toxicity of neutrophils toward erythrocytes. J. Biol. Chem. 1985;260:3321-3329.
Not in our library
45.
Learn D.B., Fried V.A., Thomas E.L. Taurine and hypotaurine content of human leukocytes. J. Leukoc. Biol. 1990;48:174-182.
Not in our library10.1002/jlb.48.2.174
46.
Blomgran R., Zheng L., Stendahl O. Cathepsin-cleaved Bid promotes apoptosis in human neutrophils via oxidative stress-induced lysosomal membrane permeabilization. J. Leukoc. Biol. 2007;81:1213-1223.
Not in our library10.1189/jlb.0506359
47.
Kanayama A., Miyamoto Y. Apoptosis triggered by phagocytosis-related oxidative stress through FLIPs down-regulation and JNK activation. J. Leukoc. Biol. 2007;82:1344-1352.
Not in our library10.1189/jlb.0407259
48.
Pattison D.I., Davies M.J. Absolute rate constants for the reaction of hypochlorous acid with protein side chains and peptide bonds. Chem. Res. Toxicol. 2001;14:453-464.
Not in our library10.1021/tx0155451
49.
Hawkins C.L., Pattison D.I., Davies M.J. Hypochlorite-induced oxidation of amino acids, peptides, and proteins. Amino Acids. 2003;25:259-274.
Not in our library10.1007/s00726-003-0016-x
50.
Pattison D.I., Davies M.J. Kinetic analysis of the reaction of hypobromous acid with protein components: Implication for cellular damage and the use of 3-bromotyrosine as a marker of oxidative stress. Biochemistry. 2004;43:4799-4809.
Not in our library10.1021/bi035946a
51.
Skaff O., Pattison D.I., Davies M.J. Hypothiocyanous acid reactivity with low-molecular-mass and protein thiols: Absolute rate constants and assessment of biological relevance. Biochem. J. 2009;442:111-117.
Not in our library10.1042/bj20090276
52.
Hawkins C.L. The role of hypothiocyanous acid (HOSCN) in biological systems. Free Radic. Res. 2009;43:1147-1158.
Not in our library10.3109/10715760903214462
53.
Barrett T.J., Hawkins C.L. Hypothiocyanous acid: Benign or deadly? Chem. Res. Toxicol. 2012;25:263-273.
Not in our library10.1021/tx200219s
54.
Salavej P., Spalteholz H., Arnhold J. Modification of amino acid residues in human serum albumin by myeloperoxidase. Free Radic. Biol. Med. 2006;40:516-525.
Not in our library10.1016/j.freeradbiomed.2005.09.007
55.
Senthilmohan R., Kettle A.J. Bromination and chlorination reactions of myeloperoxidase at physiological concentrations of bromide and chloride. Arch. Biochem. Biophys. 2006;445:235-244.
Not in our library10.1016/j.abb.2005.07.005
56.
Marquez L.A., Dunford H.B. Kinetic of oxidation of tyrosine and dityrosine by myeloperoxidase compounds I and II. J. Biol. Chem. 1995;270:30434-30440.
Not in our library10.1074/jbc.270.51.30434
57.
Burner U., Jantschko W., Obinger C. Kinetic of oxidation of aliphatic and aromatic thiols by myeloperoxidase compounds I and II. FEBS Lett. 1999;43:290-296.
Not in our library10.1016/s0014-5793(98)01727-x
58.
Burner U., Furtmüller P.G., Kettle A.J., Koppenol W.H., Obinger C. Mechanism of reaction of myeloperoxidase with nitrite. J. Biol. Chem. 2000;275:20597-20601.
Not in our library10.1074/jbc.m000181200
59.
Jantschko W., Furtmüller P.G., Allegra M., Livrea M.A., Jakopitsch C., Regelsberger G., Obinger C. Redox intermediates of plant and mammalian peroxidases: A comparative transient-kinetic study of their reactivity toward indole derivatives. Arch. Biochem. Biophys. 2002;398:12-22.
Not in our library10.1006/abbi.2001.2674
60.
Meotti F.C., Jameson G.N.L., Turner R., Harwood T.D., Stockwell S., Rees M.D., Thomas S.R., Kettle A.J. Urate as a physiological substrate for myeloperoxidase. Implications for hyperuricemia and inflammation. J. Biol. Chem. 2011;286:12901-12911.
Not in our library10.1074/jbc.m110.172460
61.
Spalteholz H., Furtmüller P.G., Jakopitsch C., Obinger C., Schewe T., Sies H., Arnhold J. Kinetic evidence for rapid oxidation of (−)-epicatechin by human myeloperoxidase. Biochem. Biophys. Res. Commun. 2008;371:810-813.
Not in our library10.1016/j.bbrc.2008.04.139
62.
Kirchner T., Flemmig J., Furtmüller P.G., Obinger C., Arnhold J. (−)-Epicatechin enhances the chlorinating activity of human myeloperoxidase. Arch. Biochem. Biophys. 2010;495:21-27.
Not in our library10.1016/j.abb.2009.12.013
63.
Gau J., Furtmüller P.G., Obinger C., Prévost M., van Antwerpen P., Arnhold J., Flemmig J. Flavonoids as promoters of the (pseudo)halogenating activity of lactoperoxidase and myeloperoxidase. Free Radic. Biol. Med. 2016;97:307-319.
Not in our library10.1016/j.freeradbiomed.2016.06.026
64.
Bolscher B.G.J.M., Zoutberg G.R., Cuperus R.A., Wever R. Vitamin C stimulates the chlorinating activity of human myeloperoxidase. Biochim. Biophys. Acta. 1984;784:189-191.
Not in our library10.1016/0167-4838(84)90127-4
65.
Dunford H.B., Hsuanyu Y. Kinetics of oxidation of serotonin by myeloperoxidase compounds I and II. Biochem. Cell Biol. 1999;77:449-457.
Not in our library10.1139/o99-052
66.
Kettle A.J., Anderson R.F., Hampton M.B., Winterbourn C.C. Reactions of superoxide with myeloperoxidase. Biochemistry. 2007;46:4888-4897.
Not in our library10.1021/bi602587k
67.
Kettle A.J., Winterbourn C.C. Mechanism of inhibition of myeloperoxidase by anti-inflammatory drugs. Biochem. Pharmacol. 1991;41:1485-1492.
Not in our library10.1016/0006-2952(91)90565-m
68.
Nève J., Parij N., Moguilevsky N. Inhibition of the myeloperoxidase chlorinating activity by non-steroidal anti-inflammatory drugs investigated with a human recombinant enzyme. Eur. J. Pharmacol. 2001;417:37-43.
Not in our library10.1016/s0014-2999(01)00895-0
69.
Kettle A.J., Sangster D.F., Gebicki J.M., Winterbourn C.C. A pulse radiolysis investigation of the reactions of myeloperoxidase with superoxide and hydrogen peroxide. Biochim. Biophys. Acta. 1988;956:58-62.
Not in our library10.1016/0167-4838(88)90297-x
70.
Bielski B.H.J., Cabelli D.E., Arudi R.L. Reactivity of HO2/O2− radicals in aqueous solution. J. Phys. Chem. Ref. Data. 1985;14:1041-1100.
Not in our library10.1063/1.555739
71.
Huie R.E., Padmaja S. Reactions of NO and O2−. Free Radic. Res. Commun. 1993;18:195-199.
Not in our library10.3109/10715769309145868
72.
Kissner R., Nauser T., Bugnon P., Lye P.G., Koppenol W.H. Formation and properties of peroxynitrite as studied by laser flash photolysis, high-pressure stopped-flow technique, and pulse radiolysis. Chem. Res. Toxicol. 1997;10:1285-1292.
Not in our library10.1021/tx970160x
73.
Furtmüller P.G., Jantschko W., Zederbauer M., Schwanninger M., Jakopitsch C., Herold S., Koppenol W.H., Obinger C. Peroxynitrite efficiently mediates the interconversions of redox intermediates of myeloperoxidase. Biochem. Biophys. Res. Commun. 2005;337:944-954.
Not in our library10.1016/j.bbrc.2005.09.138
74.
Koyani C.N., Flemmig J., Malle E., Arnhold J. Myeloperoxidase scavenges peroxynitrite: A novel anti-inflammatory action of the heme enzyme. Arch. Biochem. Biophys. 2015;571:1-9.
Not in our library10.1016/j.abb.2015.02.028
75.
Fernandez H.N., Henson P.M., Otani A., Hugli T.E. Chemotactic response of human C3a and C5a anaphylatoxins. I. Evaluation of C3a and C5a leukotaxis in vitro and under simulated in vivo conditions. J. Immunol. 1978;120:109-115.
Not in our library
76.
Witko-Sarsat V., Rieu P., Descamps-Latscha B., Lesavre P., Halbwachs-Mecarelli L. Neutrophils: Molecules, functions, and pathophysiological aspects. Lab. Investig. 2000;80:617-653.
Not in our library10.1038/labinvest.3780067
77.
Mukaida N. Pathophysiological roles of interleukin 8/CXCL8 in pulmonary diseases. Am. J. Physiol. Lung Cell Mol. Physiol. 2003;284:L566-L577.
Not in our library10.1152/ajplung.00233.2002
78.
Phillipson M., Kubes P. The neutrophil in vascular inflammation. Nat. Med. 2011;17:1381-1390.
Not in our library10.1038/nm.2514
79.
Schlorke D., Thomas L., Samsonov S.A., Huster D., Arnhold J., Pichert A. The influence of glycosaminoglycans on IL-8-mediated functions of neutrophils. Carbohydr. Res. 2012;356:196-203.
Not in our library10.1016/j.carres.2012.02.025
80.
Sengeløv H., Kjeldson L., Borregaard N. Control of exocytosis in early neutrophil activation. J. Immunol. 1993;150:1535-1543.
Not in our library
81.
Sengeløv H., Follin P., Kjeldson L., Lollike K., Dahlgren C., Borregaard N. Mobilization of granules and secretory vesicles during in vivo exudation of human neutrophils. J. Immunol. 1995;154:4157-4165.
Not in our library
82.
Edwards S.W. Biochemistry and Physiology of the Neutrophil. 1994.
Not in our library
83.
Segal A.W. How neutrophils kill microbes. Annu. Rev. Immunol. 2005;23:197-223.
Not in our library10.1146/annurev.immunol.23.021704.115653
84.
Elsbach P. The bactericidal/permeability-increasing protein (BPI) in antibacterial host defense. J. Leukoc. Biol. 1998;64:14-18.
Not in our library10.1002/jlb.64.1.14
85.
Soehnlein O., Lindbom L. Neutrophil-derived azurocidin alarms the immune system. J. Leukoc. Biol. 2009;85:344-351.
Not in our library10.1189/jlb.0808495
86.
Lehrer R.I., Lu W. α-Defensins in human innate immunity. Immunol. Rev. 2012;245:84-112.
Not in our library10.1111/j.1600-065x.2011.01082.x
87.
Odell E.W., Sarra R., Foxworthy M., Chapple D.S., Evans R.W. Antibacterial activities of peptides homologous to a loop region in lactoferrin. FEBS Lett. 1996;382:175-178.
Not in our library10.1016/0014-5793(96)00168-8
88.
Famaud S., Evans R.W. Lactoferrin—A multifunctional protein with antimicrobial properties. Mol. Immunol. 2003;40:395-405.
Not in our library10.1016/s0161-5890(03)00152-4
89.
Wardlaw A.C. The complement-dependent bacteriolytic activity of normal human serum. I. The effect of pH and ionic strength and the role of lysozyme. J. Exp. Med. 1962;115:1231-1249.
Not in our library10.1084/jem.115.6.1231
90.
Korkmaz B., Horwitz M.S., Jenne D.E., Gauthier F. Neutrophil elastase, proteinase 3, and cathepsin G as therapeutic targets in human diseases. Pharmacol. Rev. 2010;62:726-759.
Not in our library10.1124/pr.110.002733
91.
Fasciglione G.F., Marini S., D’Alessio S., Politi V., Coletta M. pH- and temperature-dependence of functional modulation in metalloproteinases. A comparison between neutrophil collagenase and gelatinases A and B. Biophys. J. 2000;79:2138-2149.
Not in our library10.1016/s0006-3495(00)76461-7
92.
Babior B.M. NADPH oxidase. Curr. Opin. Immunol. 2004;16:42-47.
Not in our library10.1016/j.coi.2003.12.001
93.
Foote J.R., Behe P., Frampton M., Lewine A.P., Segal A.W. An exploration of charge compensating ion channels across the phagocytic vacuole of neutrophils. Front. Pharmacol. 2017;8:94.
Not in our library10.3389/fphar.2017.00094
94.
Segal A.W., Geisow M., Garcia R., Harper A., Miller R. The respiratory burst of phagocytic cells is associated with a rise in vacuolar pH. Nature. 1981;290:406-409.
Not in our library10.1038/290406a0
95.
Cech P., Lehrer R.I. Phagolysosomal pH of human neutrophils. Blood. 1984;63:88-95.
Not in our library10.1182/blood.v63.1.88.88
96.
Levine A.P., Duchen M.R., de Villiers S., Rich P.R., Segal A.W. Alkalinity of neutrophil phagocytic vacuoles is modulated by HVCN1 and has consequences for myeloperoxidase activity. PLoS ONE. 2015;10:e0125906.
Not in our library10.1371/journal.pone.0125906
97.
Variations in the Phagosomal Environment of Human Neutrophils and Mononuclear Phagocyte Subsets · 2019
Not checked
98.
Murata K. Acidic glycosaminoglycans in human platelets and leukocytes: The isolation and enzymatic characterization of chondroitin-4-sulfate. Clin. Chim. Acta. 1974;57:115-124.
Not in our library10.1016/0009-8981(74)90418-5
99.
Kolset S.O., Gallaher J.T. Proteoglycans in haemopoietic cells. Biochim. Biophys. Acta. 1990;1032:191-211.
Not in our library10.1016/0304-419x(90)90004-k
100.
Radi R. Oxygen radicals, nitric oxide, and peroxynitrite: Redox pathway in molecular medicine. Proc. Natl. Acad. Sci. USA. 2018;115:5839-5848.
Not in our library10.1073/pnas.1804932115
101.
Flint D.H., Tominello J.F., Emptage M.H. The investigation of Fe-S cluster containing hydrolyases by superoxide. J. Biol. Chem. 1993;268:22369-22376.
Not in our library
102.
Gardner P.R. Aconitase: Sensitive target and measure of superoxide. Meth. Enzymol. 2002;349:9-23.
Not in our library10.1016/s0076-6879(02)49317-2
103.
Jantschko W., Furtmüller P.G., Zederbauer M., Lanz M., Jakopitsch C., Obinger C. Direct conversion of ferrous myeloperoxidase to compound II by hydrogen peroxide: An anaerobic stopped-flow study. Biochem. Biophys. Res. Commun. 2003;312:292-298.
Not in our library10.1016/j.bbrc.2003.10.117
104.
Hoogland H., Dekker H.L., van Riel C., van Kuilenberg A., Muijsers A.O., Wever R. A steady-state study on the formation of compounds II and III of myeloperoxidase. Biochim. Biophys. Acta. 1988;955:337-345.
Not in our library10.1016/0167-4838(88)90213-0
105.
Hampton M.B., Kettle A.J., Winterbourn C.C. Involvement of superoxide and myeloperoxidase in oxygen-dependent killing of Staphylococcus aureus by neutrophils. Infect. Immun. 1996;64:3512-3517.
Not in our library10.1128/iai.64.9.3512-3517.1996
106.
Kettle A.J., Geyde C.A., Winterbourn C.C. Mechanism of inactivation of myeloperoxidase by 4-aminobenzoic acid hydrazide. Biochem. J. 1997;321:503-508.
Not in our library10.1042/bj3210503
107.
Bolscher B.G.J.M., Wever R. A kinetic study of the reaction between human myeloperoxidase, hydroperoxides and cyanide. Inhibition by chloride and thiocyanate. Biochim. Biophys. Acta. 1984;788:1-10.
Not in our library10.1016/0167-4838(84)90290-5
108.
Lehrer R.I. Inhibition by sulfonamides of the candidacidal activity of human neutrophils. J. Clin. Investig. 1971;50:2498-2505.
Not in our library10.1172/jci106750
109.
Koch C. Effect of sodium azide upon normal and pathological granulocyte function. Acta Pathol. Microbiol. Scand. B. 1974;82:136-142.
Not in our library10.1111/j.1699-0463.1974.tb02304.x
110.
Yamamoto K., Miyoshi-Koshio T., Utsuki Y., Mizuno S., Suzuki K. Virucidal activity and viral protein modification by myeloperoxidase: A candidate for defense factor of human polymorphonuclear leukocytes against influenza virus infection. J. Infect. Dis. 1991;164:8-14.
Not in our library10.1093/infdis/164.1.8
111.
Aratani Y., Koyama H., Nyui S.-I., Suzuki K., Kura F., Maeda N. Severe impairment in early host defense against Candida albicans in mice deficient in myeloperoxidase. Infect. Immun. 1999;67:1828-1836.
Not in our library10.1128/iai.67.4.1828-1836.1999
112.
Chapman A.L., Hampton M.B., Senthilmohan R., Winterbourn C.C., Kettle A.J. Chlorination of bacterial and neutrophil proteins during phagocytosis and killing of Staphylococcus aureus. J. Biol. Chem. 2002;277:9757-9762.
Not in our library10.1074/jbc.m106134200
113.
Reeves E.P., Nagl M., Godovac-Zimmermann J., Segal A.W. Reassessment of the microbicidal activity of reactive oxygen species and hypochlorous acid with reference to the phagocytic vacuole of the neutrophil granulocyte. J. Med. Microbiol. 2003;53:643-651.
Not in our library10.1099/jmm.0.05181-0
114.
Hirche T.O., Gaut J.P., Heinecke J.W., Belaaouaj A. Myeloperoxidase plays critical roles in killing Klebsiella pneumoniae and inactivating elastase: Effects of host defense. J. Immunol. 2005;174:1557-1565.
Not in our library10.4049/jimmunol.174.3.1557
115.
Parry M.F., Root R.K., Metcalf J.A., Delaney K.K., Kaplow L.S., Richar W.J. Myeloperoxidase deficiency: Prevalence and clinical significance. Ann. Intern. Med. 1981;85:293-301.
Not in our library10.7326/0003-4819-95-3-293
116.
Kutter D. Prevalence of myeloperoxidase deficiency: Population studies using Bayer-Technicon automated hematology. J. Mol. Med. 1998;76:669-675.
Not in our library10.1007/s001090050266
117.
Trasher A.J., Keep N.H., Wientjes F., Segal A.W. Chronic granulomatous disease. Biochim. Biophys. Acta. 1994;1227:1-24.
Not in our library10.1016/0925-4439(94)90100-7
118.
Lehrer R.I., Cline M.J. Leukocyte myeloperoxidase deficiency and disseminated candidiasis: The role of myeloperoxidase in resistance to Candida infection. J. Clin. Investig. 1969;48:1478-1488.
Not in our library10.1172/jci106114
119.
Cech P., Stadler H., Widmann J.J., Rohner A., Miescher P.A. Leukocyte myeloperoxidase deficiency and diabetes mellitus associated with Candida albicans liver abscess. Am. J. Med. 1979;66:149-153.
Not in our library10.1016/0002-9343(79)90507-2
120.
Nguyen C., Katner H.P. Myeloperoxidase deficiency manifesting as pustular candida dermatitis. Clin. Infect. Dis. 1997;24:258-260.
Not in our library10.1093/clinids/24.2.258
121.
Gaut J.P., Yeh G.C., Tran H.D., Byun J., Henderson J.P., Richter G.M., Brennan M.-L., Lusis A.J., Belaaouiaj A., Hotchkiss R.S., et al. Neutrophils employ the myeloperoxidase system to generate antimicrobial brominating and chlorinating oxidants during sepsis. Proc. Natl. Acad. Sci. USA. 2001;98:11961-11966.
Not in our library10.1073/pnas.211190298
122.
Brennan M.-L., Gaur A., Pahuja A., Lusis A.J., Reynolds W.F. Mice lacking myeloperoxidase are more susceptible to experimental encephalomyelitis. J. Neuroimmunol. 2001;112:97-105.
Not in our library10.1016/s0165-5728(00)00392-1
123.
Aratani Y., Kura F., Watanabe H., Akagawa H., Takano Y., Ishida-Okawara A., Suzuki K., Maeda N., Koyama H. Contribution of the myeloperoxidase-dependent oxidative systems to host defence against Cryptococcus neoformans. J. Med. Microbiol. 2006;55:1291-1299.
Not in our library10.1099/jmm.0.46620-0
124.
Zschaler J., Schlorke D., Arnhold J. Differences in innate immune response between man and mouse. Crit. Rev. Immunol. 2014;34:433-454.
Not in our library10.1615/critrevimmunol.2014011600
125.
Eisenhauer P.B., Lehrer R.I. Mouse neutrophils lack defensins. Infect. Immun. 1992;60:3446-3447.
Not in our library10.1128/iai.60.8.3446-3447.1992
126.
Lehrer R.I., Lichtenstein A.K., Ganz T. Defensins: Antimicrobial and cytotoxic peptides of mammalian cells. Annu. Rev. Immunol. 1993;11:105-128.
Not in our library10.1146/annurev.iy.11.040193.000541
127.
Rausch P.G., Moore T.G. Granule enzymes of polymorphonuclear neutrophils: A phylogentic comparison. Blood. 1975;46:913-919.
Not in our library10.1182/blood.v46.6.913.913
128.
Noguchi N., Nakano K., Aratani Y., Koyama H., Kodama T., Niki E. Role of myeloperoxidase in the neutrophil-induced oxidation of low density lipoprotein as studied by myeloperoxidase-knockout mouse. J. Biochem. 2000;127:971-976.
Not in our library10.1093/oxfordjournals.jbchem.a022713
129.
Lennartsson A., Pieters K., Vidovic K., Gullberg U. A murine antibacterial ortholog to human bactericidal/permeability-increasing protein (BPI) is expressed in testis, epididymis, and bone marrow. J. Leukoc. Biol. 2005;77:369-377.
Not in our library10.1189/jlb.0304159
130.
Simon H.-U. Neutrophil apoptosis pathways and their modification in inflammation. Immunol. Rev. 2003;193:101-110.
Not in our library10.1034/j.1600-065x.2003.00038.x
131.
Walker A., Ward C., Taylor E.I., Dransfield I., Hart S.P., Haslett C., Rossi A.G. Regulation of neutrophil apoptosis and removal of apoptotic cells. Curr. Drug Targets—Inflamm. Allergy. 2005;4:447-454.
Not in our library10.2174/1568010054526278
132.
Nathan C. Points of control in inflammation. Nature. 2002;420:846-852.
Not in our library10.1038/nature01320
133.
Barton G.M. A calculated response: Control of inflammation by the innate immune system. J. Clin. Investig. 2008;118:413-420.
Not in our library10.1172/jci34431
134.
Flemmig J., Leßig J., Reibetanz U., Dautel P., Arnhold J. Non-vital polymorphonuclear leukocytes express myeloperoxidase on their surface. Cell. Physiol. Biochem. 2008;21:287-296.
Not in our library10.1159/000129387
135.
Vandivier R.W., Fadok V.A., Hoffmann P.R., Bratton D.L., Penvari C., Brown K.K., Brain J.D., Accurso F.J., Henson P.M. Elastase-mediated phosphatidylserine receptor cleavage impairs apoptotic cell clearance in cystic fibrosis and bronchiectasis. J. Clin. Investig. 2002;109:661-670.
Not in our library10.1172/jci0213572
136.
Wang R., Town T., Gokarn V., Flavell R.A., Chandawarkar R.Y. HSP70 enhances macrophage phagocytosis by interaction with lipid raft-associated TLR-7 and upregulating p38 MAPK and PI3K pathways. J. Surg. Res. 2006;136:58-69.
Not in our library10.1016/j.jss.2006.06.003
137.
Vandivier R.W., Henson P.M., Douglas I.S. Burying the death: The impact of failed apoptosis removal (efferocytosis) on chronic inflammatory lung disease. Chest. 2006;129:1673-1682.
Not in our library10.1378/chest.129.6.1673
138.
Fadok V.A., Bratton D.L., Konowal A., Freed P.W., Westcott J.Y., Henson P.M. Macrophages that have ingested apoptotic cells in vitro inhibit proinflammatory cytokine production hrough autocrine/paracrine mechanisms involving TGF-beta, PGE2 and PAF. J. Clin. Investig. 1998;101:890-898.
Not in our library10.1172/jci1112
139.
Fadeel B. Programmed cell clearance. Cell. Mol. Life Sci. 2003;60:2575-2585.
Not in our library10.1007/s00018-003-3145-1
140.
Krysko D.V., D’Herde K., Vandenabeele P. Clearance of apoptotic and necrotic cells and its immunological consequences. Apoptosis. 2006;11:1673-1682.
Not in our library10.1007/s10495-006-9527-8
141.
Erwig L.-P., Henson P.M. Immunological consequences of apoptotic cell phagocytosis. Am. J. Pathol. 2007;171:2-8.
Not in our library10.2353/ajpath.2007.070135
142.
Keel M., Ungethüm U., Steckholzer U., Niederer E., Hartung T., Trentz O., Ertel W. Interleukin-10 counterregulates proinflammatory cytokine-induced inhibition of neutrophil apoptosis during severe sepsis. Blood. 1997;90:3356-3363.
Not in our library10.1182/blood.v90.9.3356
143.
Haslett C. Granulocyte apoptosis and its role in the resolution and control of lung inflammation. Am. J. Respir. Crit. Care Med. 1999;160:S5-S11.
Not in our library10.1164/ajrccm.160.supplement_1.4
144.
Hirahashi J., Mekala D., van Ziffie J., Xiao L., Saffaripour S., Wagner D.D., Shapiro S.D., Lowell C., Mayadas T.N. Mac-1 signalling via SRc-family and Syk kinases results in elastase-dependent vasculopathy. Immunity. 2006;25:271-283.
Not in our library10.1016/j.immuni.2006.05.014
145.
Fadok V.A., Bratton D.L., Guthrie L., Henson P.M. Differential effects of apoptotic versus lysed cells on macrophage production of cytokines: Role of proteases. J. Immunol. 2001;166:8647-8654.
Not in our library10.4049/jimmunol.166.11.6847
146.
Brinkmann V., Reichard U., Goosmann C., Fauler B., Uhlemann Y., Weiss D.S., Weinrauch Y., Zychlinsky A. Neutrophil extracellular traps kill bacteria. Science. 2004;303:1532-1535.
Not in our library10.1126/science.1092385
147.
Neutrophil Extracellular Traps Contain Calprotectin, a Cytosolic Protein Complex Involved in Host Defense against Candida albicans · 2009
No notice
148.
Papayannopoulos Y., Zychlinsky A. NETs: A new strategy for using old weapons. Trends Immunol. 2009;30:513-521.
Not in our library10.1016/j.it.2009.07.011
149.
Urban C.F., Reichard U., Brinkmann V., Zychlinsky A. Neutrophil extracellular traps capture and kill Candida albicans yeast and hyphal forms. Cell. Microbiol. 2006;8:668-676.
Not in our library10.1111/j.1462-5822.2005.00659.x
150.
Fuchs T.A., Brill A., Duerschmied D., Schatzberg D., Monestier M., Uhlemann Y., Myers D.D., Wroblewski S.K., Wakefield T.W., Hartwig J.H., et al. Extracellular DNA traps promote thrombosis. Proc. Natl. Acad. Sci. USA. 2010;107:15880-15885.
Not in our library10.1073/pnas.1005743107
151.
Martinod K., Demers M., Fuchs T.A., Wong S.L., Brill A., Gallant M., Hu J., Wang Y., Wagner D.D. Neutrophil histone modification by peptidylarginine deiminase 4 is critical for deep vein thrombosis in mice. Proc. Natl. Acad. Sci. USA. 2013;110:8674-8679.
Not in our library10.1073/pnas.1301059110
152.
Cools-Lartigue J., Spicer J., McDonald B., Gowing S., Chow S., Giannias B., Bourdeau F., Kubes P., Ferri L. Neutrophil extracellular traps sequester circulating tumor cells and promote metastasis. J. Clin. Investig. 2013;123:3446-3458.
Not in our library10.1172/jci67484
153.
Metzler K.D., Fuchs T.A., Nauseef W.M., Reumaux D., Roesler J., Schulze I., Wahn V., Papyannopoulos V., Zychlinsky A. Myeloperoxidase is required for neutrophil extracellular trap formation: Implications for innate immunity. Blood. 2011;117:953-959.
Not in our library10.1182/blood-2010-06-290171
154.
Parker H., Albrett A.M., Kettle A.J., Winterbourn C.C. Myeloperoxidase associated with neutrophil extracellular traps is active and mediates bacterial killing in the presence of hydrogen peroxide. J. Leukoc. Biol. 2012;91:369-376.
Not in our library10.1189/jlb.0711387
155.
Sheppard F.R., Kelher M.R., Moore E.E., McLaughlin N.J., Banerjee A., Silliman C.C. Structural organization of the neutrophil NADPH oxidase: Phosphorylation and translocation during priming and activation. J. Leukoc. Biol. 2005;78:1025-1042.
Not in our library10.1189/jlb.0804442
156.
Canton J., Khezri R., Glogauer M., Grinstein S. Contrasting phagosome pH regulation and maturation in human M1 and M2 macrophages. Mol. Biol. Cell. 2014;25:3330-3341.
Not in our library10.1091/mbc.e14-05-0967
157.
Mantovani A., Sica A., Sozzani S., Allavena P., Vecchi A., Locati M. The chemokine system in diverse forms of macrophage activation and polarization. Trends Immunol. 2004;25:677-686.
Not in our library10.1016/j.it.2004.09.015
158.
Schneemann M., Schoeden G. Macrophage biology and immunology: Man is not a mouse. J. Leukoc. Biol. 2007;81:579-580.
Not in our library10.1189/jlb.1106702
159.
Yates R.M., Hermetter A., Taylor G.A., Russell D.G. Macrophage activation downregulates the degradative capacity of the phagosome. Traffic. 2007;8:241-250.
Not in our library10.1111/j.1600-0854.2006.00528.x
160.
Balce D.R., Li B., Allan E.R., Rybicka J.M., Krohn R.M., Yates R.M. Alternative activation of macrophages by IL-4 enhances the proteolytic capacity of their phagosomes through synergistic mechanisms. Blood. 2011;118:4199-4208.
Not in our library10.1182/blood-2011-01-328906
161.
Lefkowitz D.L., Mone J., Lefkowitz S.S. Myeloperoxidase: The good, the bad, and the ugly. Crit. Immunol. Rev. 2010;6:123-129.
Not in our library10.2174/157339510791111691
162.
Myeloperoxidase as an Active Disease Biomarker: Recent Biochemical and Pathological Perspectives · 2018
No notice
163.
Willard B.B., Ruse C.I., Keightley J.A., Bond M., Kinter M. Site-specific quantitation of protein nitration using liquid chromatography/tandem mass spectrometry. Anal. Chem. 2003;75:2370-2376.
Not in our library10.1021/ac034033j
164.
Tiruppathi C., Naqvi T., Wu Y., Vogel S.M., Minshall R.D., Malik A.B. Albumin mediates the transcystosis of myeloperoxidase by means of caveolae in endothelial cells. Proc. Natl. Acad. Sci. USA. 2004;101:7699-7704.
Not in our library10.1073/pnas.0401712101
165.
Capeillere-Blandin C., Gausson V., Descamps-Latscha B., Witko-Sarsat V. Biochemical and spectrophotometric significance of advanced oxidized protein products. Biochim. Biophys. Acta. 2004;1689:91-102.
Not in our library10.1016/j.bbadis.2004.02.008
166.
Matheson M.R., Wong P.S., Travis J. Enzymatic inactivation of human alpha-1 proteinase inhibitor by neutrophil myeloperoxidase. Biochem. Biophys. Res. Commun. 1979;88:402-409.
Not in our library10.1016/0006-291x(79)92062-x
167.
Patterson S.D. Mammalian α1-antitrypsins: Comparative biochemistry and genetics of the major plasma serpins. Comp. Biochem. Physiol. B. 1991;100:439-454.
Not in our library10.1016/0305-0491(91)90202-o
168.
Hiemstra P.S., van Wetering S., Stolk J. Neutrophil serine proteinases and defensins in chronic obstructive pulmonary disease: Effects on pulmonary epithelium. Eur. Respir. J. 1988;12:1200-1208.
Not in our library10.1183/09031936.98.12051200
169.
Bouriche H., Salavei P., Lessig J., Arnhold J. Differential effects of flavonols on inactivation of α1-antitrypsin induced by hypohalous acids and the myeloperoxidase-hydrogen peroxide-halide systems. Arch. Biochem. Biophys. 2007;459:137-143.
Not in our library10.1016/j.abb.2006.10.030
170.
Zheng L., Nukuna B., Brennan M.-L., Sun M., Goormastic M., Settle M., Schmitt X., Fu L., Thomson P.L., Ischiripoulos H., et al. Apolipoprotein A-1 is a selective target for myeloperoxidase-catalyzed oxidation and functional impairment in subjects with cardiovascular disease. J. Clin. Investig. 2004;114:529-541.
Not in our library10.1172/jci200421109
171.
Malle E., Marsche G., Panzenboeck U., Sattler W. Myeloperoxidase-mediated oxidation of high-density lipoproteins: Fingerprints of newly recognized potential proatherogenic lipoproteins. Arch. Biochem. Biophys. 2006;45:245-255.
Not in our library10.1016/j.abb.2005.08.008
172.
Nybo T., Cai H., Chuang C.Y., Gamon L.F., Rogowska-Wrzesinska A., Davies M.J. Chlorination and oxidation of human plasma fibronectin by myeloperoxidase-derived oxidants, and its consequences for smooth muscle cell function. Redox Biol. 2018;19:388-400.
Not in our library10.1016/j.redox.2018.09.005
173.
Myeloperoxidase-derived damage to human plasma fibronectin: Modulation by protein binding and thiocyanate ions (SCN−) · 2020
Not checked
174.
Daphna E.M., Michaela S., Eynat P., Irit A., Rimon S. Association of myeloperoxidase with heparin: Oxidative inactivation of proteins on the surface of endothelial cells by the bound enzyme. Mol. Cell. Biochem. 1998;183:55-61.
Not in our library10.1023/a:1006848730927
175.
Baldus S., Eiserich J.P., Mani A., Castrom L., Figueroa M., Chumley P., Ma W., Tousson A., White R., Bullard D.C., et al. Endothelial transcytosis of myeloperoxidase confers specificity to vascular ECM proteins as targets for tyrosine nitration. J. Clin. Investig. 2001;108:1759-1770.
Not in our library10.1172/jci200112617
176.
Eiserich J.P., Baldus S., Brennan M.-L., Ma W., Zhang C., Tousson A., Castro L., Lusis A.J., Nauseef W.M., White C.R., et al. Myeloperoxidase, a leukocyte-derived vascular NO oxidase. Science. 2002;196:2391-2394.
Not in our library10.1126/science.1106830
177.
Kubala L., Kolářová H., Vitećek J., Kremserová S., Klinke A., Lau D., Chapman A.L.P., Baldus S., Eiserich J.P. The potentiation of myeloperoxidase activity by the glycosaminoglycan-dependent binding of myeloperoxidase to proteins of the extracellular matrix. Biochim. Biophys. Acta. 2013;1830:4524-4536.
Not in our library10.1016/j.bbagen.2013.05.024
178.
Cai H., Chuang C.Y., Hawkins C.L., Davies M.J. Binding of myeloperoxidase to the extracellular matrix of smooth muscle cells and subsequent matrix modification. Sci. Rep. 2020;10:666.
Not in our library10.1038/s41598-019-57299-6
179.
Rees M.D., Whitelock J.M., Malle E., Chuang C.Y., Iozzo R.V., Nilasaroya A., Davies M.J. Myeloperoxidase-derived oxidants selectively disrupt the protein core of the heparan sulfate proteoglycan perlecan. Matrix Biol. 2010;29:63-73.
Not in our library10.1016/j.matbio.2009.09.005
180.
Nybo T., Dieterich S., Gamon L.F., Chuang C.Y., Hammer A., Hoefler G., Malle E., Rogowska-Wrzesinska A., Davies M.J. Chlorination and oxidation of the extracellular matrix protein laminin and basement membrane extracts by hypochlorous acid and myeloperoxidase. Redox Biol. 2019;20:496-513.
Not in our library10.1016/j.redox.2018.10.022
181.
Manchanda K., Kolářová H., Kerkenpaß C., Mollenhauer M., Vitećek J., Rudolph V., Kubala L., Baldus S., Adam M., Klinke A. MPO (myeloperoxidase) reduces endothelial glycocalyx thickness dependent on its cationic charge. Arterioscler. Thromb. Vasc. Biol. 2018;38:1859-1867.
Not in our library10.1161/atvbaha.118.311143
182.
Sokolov A.V., Ageeva K.V., Cherkalina O.S., Pulina M.O., Zakharova E.T., Prozorovskii V.N., Aksenov D.V., Vasilyev V.B., Panasenko O.M. Identification and properties of complexes formed by myeloperoxidase with lipoproteins and ceruloplasmin. Chem. Phys. Lipids. 2010;163:347-353.
Not in our library10.1016/j.chemphyslip.2010.02.002
183.
Chapman A.L.P., Mocatta T.J., Shiva S., Seidel A., Chen B., Khalilova I., Paumann-Page M.E., Jameson G.N.L., Winterbourn C.C., Kettle A.J. Ceruloplasmin is an endogenous inhibitor of myeloperoxidase. J. Biol. Chem. 2013;288:6464-6477.
Not in our library10.1074/jbc.m112.418970
184.
Stoj C., Kosman D.J. Cuprous oxidase activity of yeast Fet3p and human ceruloplasmin: Implication for function. FEBS Lett. 2003;554:422-426.
Not in our library10.1016/s0014-5793(03)01218-3
185.
Sokolov A.V., Pulina M.O., Ageeva K.V., Ayrapetov M.I., Berlov M.I., Volgin G.N., Markov A.G., Yablonsky P.K., Kolodkin N.I., Zakharova E.T., et al. Interaction of ceruloplasmin, lactoferrin, and myeloperoxidase. Biochemistry (Moscow). 2007;72:409-415.
Not in our library10.1134/s0006297907040074
186.
Sokolov A.V., Pulina M.O., Ageeva K.V., Runova O.I., Zakharova E.T., Vasilyev V.B. Identification of leukocyte cationic proteins that interact with ceruloplasmin. Biochemistry (Moscow). 2007;72:872-877.
Not in our library10.1134/s0006297907080093
187.
Vasilyev V.B., Kachurin A.M., Soronka N.V. Dismutation of superoxide anion radicals by ceruloplasmin. Details of the mechanism. Biokhimija. 1988;83:2051-2058.
Not in our library
188.
Segelmark M., Persson B., Hellmark T., Wieslander J. Binding and inhibition of myeloperoxidase (MPO): A major function of ceruloplasmin? Clin. Exp. Immunol. 1997;108:167-174.
Not in our library10.1046/j.1365-2249.1997.d01-992.x
189.
Park Y.S., Suzuki K., Mumby S., Taniguchi N., Gutteridge J.M.C. Antioxidant binding of caeruloplasmin to myeloperoxidase: Myeloperoxidase is inhibited, but oxidase, peroxidase, and immunoreactive properties of caeruloplasmin remain intact. Free Radic. Res. 2000;33:261-265.
Not in our library10.1080/10715760000301421
190.
Sokolov A.V., Ageeva K.V., Pulina M.O., Cherkalina O.S., Samygina V.R., Vlasova I.I., Panasenko O.M., Zakharova E.T., Vasilyev V.B. Ceruloplasmin and myeloperoxidase in complex affect the enzymatic properties of each other. Free Radic. Res. 2008;42:989-998.
Not in our library10.1080/10715760802566574
191.
Samygina V.R., Sokolov A.V., Bourenkov G., Petoukhov M.V., Pulina M.O., Zakharova E.T., Vasilyev V.B., Bartunik H., Svergun D.I. Ceruloplasmin: Macromolecular assemblies with iron-containing acute phase proteins. PLoS ONE. 2013;8:e67145.
Not in our library10.1371/journal.pone.0067145
192.
Sokolov A.V., Kostevich V.A., Zakharova E.T., Samygina V.R., Panasenko O.M., Vasilyev V.B. Interaction of ceruloplasmin with eosinophil peroxidase as compared to its interplay with myeloperoxidase: Reciprocal effect on enzymatic properties. Free Radic. Res. 2015;49:800-811.
Not in our library10.3109/10715762.2015.1005615
193.
Griffin S.V., Chapman P.T., Lianos E.A., Lockwood C.M. The inhibition of myeloperoxidase by ceruloplasmin can be reversed by anti-myeloperoxidase antibodies. Kidney Int. 1999;55:917-925.
Not in our library10.1046/j.1523-1755.1999.055003917.x
194.
Lusis A.J. Atherosclerosis. Nature. 2000;407:233-241.
Not in our library10.1038/35025203
195.
Wick G., Knoflach M., Xu Q. Autoimmune and inflammatory mechanisms in atherosclerosis. Annu. Rev. Atheroscl. 2004;22:361-403.
Not in our library10.1146/annurev.immunol.22.012703.104644
196.
Tabas I., Garcia-Gardeña G., Owens G.K. Recent insights into the cellular biology of atherosclerosis. J. Cell Biol. 2015;209:13-22.
Not in our library10.1083/jcb.201412052
197.
Pathophysiology and treatment of atherosclerosis · 2017
No notice
198.
Gisterå A., Hansson G. The immunology of atherosclerosis. Nat. Rev. Nephrol. 2017;13:368-380.
Not in our library10.1038/nrneph.2017.51
199.
Steinberg D., Parthasarathy S., Carew T.E., Khoo J.C., Witztum J.L. Beyond cholesterol: Modification of low-density lipoprotein that increases its atherogenicity. N. Engl. J. Med. 1989;320:915-924.
Not in our library10.1056/nejm198904063201407
200.
Teng N., Maghzal G., Talib J., Rashid I., Lau A.K., Stocker R. The roles of myeloperoxidase in coronary artery disease and its potential implication in plaque rupture. Redox Rep. 2017;22:51-73.
Not in our library10.1080/13510002.2016.1256119
201.
Daugherty A., Rateri D.L., Dunn J.L., Heinecke J.W. Myeloperoxidase, a catalyst for lipoprotein oxidation, is expressed in human atherosclerotic lesions. J. Clin. Investig. 1994;94:437-444.
Not in our library10.1172/jci117342
202.
Malle E., Waeg G., Schreiber R., Gröne E.F., Sattler W., Gröne H.J. Immunological evidence for the myeloperoxidase/H2O2/halide system in human atherosclerotic lesions. Eur. J. Biochem. 2000;267:4495-4503.
Not in our library10.1046/j.1432-1327.2000.01498.x
203.
Hazen S.L., Heinecke J.W. 3-Chlorotyrosine, a specific marker of myeloperoxidase-catalyzed oxidation, is markedly elevated in low density lipoprotein isolated from human atherosclerotic intima. J. Clin. Investig. 1997;99:2075-2081.
Not in our library10.1172/jci119379
204.
Takeshita J., Byun J., Nhan T.Q., Pritchard D.K., Pennathur S., Schwartz S.M., Chait A., Heinecke J.W. Myeloperoxidase generates 5-chlorouracil in human atherosclerotic tissue: A potential pathway for somatic mutagenesis by macrophages. J. Biol. Chem. 2005;281:3096-3104.
Not in our library10.1074/jbc.m509236200
205.
Hazell L.J., Stocker R. Oxidation of low-density lipoprotein with hypochlorite causes transformation of the lipoprotein into a high-uptake form for macrophages. Biochem. J. 1993;290:165-172.
Not in our library10.1042/bj2900165
206.
Yang C.Y., Gu Z.W., Yang M., Lin S.N., Garcia-Prats A.J., Rogers L.K., Welty S.E., Smith C.V. Selective modification of apoB-100 in the oxidation of low density lipoprotein by myeloperoxidase in vitro. J. Lipid Res. 1999;40:686-698.
Not in our library
207.
Smith J.D. Dysfunctional HDL as a diagnostic and therapeutic target. Arterioscler. Thromb. Vasc. Biol. 2010;30:151-155.
Not in our library10.1161/atvbaha.108.179226
208.
Fisher E.A., Feig J.E., Hewing B., Hazen S.L., Smith J.D. High-density lipoprotein function, dysfunction, and reverse cholesterol transport. Arterioscler. Thromb. Vasc. Biol. 2012;32:2813-2820.
Not in our library10.1161/atvbaha.112.300133
209.
Wang Z., Nicholls S.J., Rodriguez E.R., Kummu O., Hörkkö S., Barnard J., Reynolds W.F., Topol E., DiDonato J.A., Hazen S.L. Protein carbamylation links inflammation, smoking, uremia and atherogenesis. Nat. Med. 2007;113:1176-1184.
Not in our library10.1038/nm1637
210.
Delporte C., Boudjeltia K.Z., Furtmüller P.G., Maki R.A., Dieu M., Noyon C., Soudi M., Dufour D., Coremans C., Nuyens V., et al. Myeloperoxidase-catalyzed oxidation of cyanide to cyanate: A potential carbamylation route involved in the formation of atherosclerotic plaques? J. Biol. Chem. 2018;293:6374-6386.
Not in our library10.1074/jbc.m117.801076
211.
Zhang C., Reiter C., Eiserich J.P., Boersma B., Parks D.A., Beckman J.S., Barnes S., Kirk M., Baldus S., Darley-Usmar V.M., et al. L-Arginine chlorination products inhibit endothelial nitric oxide production. J. Biol. Chem. 2001;276:27159-27165.
Not in our library10.1074/jbc.m100191200
212.
Koeth R.A., Haselden V., Tang W.H. Myeloperoxidase in cardiovascular disease. Adv. Clin. Chem. 2013;62:1-32.
Not in our library10.1016/b978-0-12-800096-0.00001-9
213.
Wang Y., Rosen H., Madtes D.K., Shao B., Martin T.R., Heinecke J.W., Fu X. Myeloperoxidase inactivates TIMP-1 by oxidizing its N-terminal cysteine residue: An oxidative mechanism for regulating proteolysis during inflammation. J. Biol. Chem. 2007;282:31826-31834.
Not in our library10.1074/jbc.m704894200
214.
Ehrenfeld P., Matus C.E., Pavicic F., Toledo C., Nualart F., Gonzalez C.B., Burgos R.A., Bhoola K.D., Figueroa C.D. Kinin B1 receptor activation turns on exocytosis of matrixmetalloprotease-0 and myeloperoxidase in human neutrophils: Involvement of mitogen-activated protein kinase family. J. Leukoc. Biol. 2009;86:1179-1189.
Not in our library10.1189/jlb.0109012
215.
Baldus S., Heeschen C., Meinertz T., Zeiher A.M., Eiserich J.P., Münzel T., Simoons M.L., Hamm C.W. Myeloperoxidase serum levels predict risk in patients with acute coronary syndromes. Circulation. 2003;108:1440-1445.
Not in our library10.1161/01.cir.0000090690.67322.51
216.
Tang W.H.W., Katz R., Brennan M.-L., Tracy R.P., Aviles R.J., Psaty B.M., Hazen S.L. Usefulness of myeloperoxidase levels in healthy elderly subjects to predict risk of developing heart failure. Am. J. Cardiol. 2009;103:1269-1274.
Not in our library10.1016/j.amjcard.2009.01.026
217.
Nussbaum C., Klinke A., Adam M., Baldus S., Sperandio M. Myeloperoxidase: A leukocyte-derived protagonist of inflammation and cardiovascular disease. Antioxid. Redox Signal. 2013;18:692-713.
Not in our library10.1089/ars.2012.4783
218.
Narula J., Nakano M., Virmani R., Kolodgie F.D., Petersen R., Newcomb R., Malik S., Fuster V., Finn A.V. Histopathologic characteristics of atherosclerotic coronary disease and implications of the findings for the invasive and noninvasive detection of vulnerable plaques. J. Am. Cell Cardiol. 2013;61:1041-1051.
Not in our library10.1016/j.jacc.2012.10.054
219.
Jennette J.C., Falk R.J., Hu P., Xiao H. Pathogenesis of anti-neutrophil cytoplasmic autoantibodies associated small vessel vasculitis. Annu. Rev. Pathol. Mech. Dis. 2017;12:139-160.
Not in our library10.1146/annurev-pathol-011811-132453
220.
Crisford H., Sapey E., Stockley R.A. Proteinase 3; a potential target in chronic obstructive pulmonary disease and other chronic inflammatory diseases. Respir. Res. 2018;19:180.
Not in our library10.1186/s12931-018-0883-z
221.
Gómez-Puerta J.A., Bosch X. Anti-neutrophil cytoplasmic antibody pathogenesis in small-vessel vasculitis. Am. J. Pathol. 2009;175:1790-1798.
Not in our library10.2353/ajpath.2009.090533
222.
Yang J.J., Tuttle R.H., Hogan S.L., Taylor J.G., Philipps B.D., Falk R.J., Jennette J.C. Target antigens for anti-neutrophil cytoplasmic autoantibodies (ANCA) are on the surface of primed and apoptotic but not unstimulated neutrophils. Clin. Exp. Immunol. 2000;121:165-172.
Not in our library10.1046/j.1365-2249.2000.01228.x
223.
Kain R., Exner M., Brandes R., Ziebermayr R., Cunningham D., Alderson D.A., Davidovits A., Raab I., Jahn R., Ashour O., et al. Molecular mimicry in pauci-immune focal necrotizing glomerulonephritis. Nat. Med. 2008;14:1088-1096.
Not in our library10.1038/nm.1874
224.
Jennette J.C., Nachman P.H. ANCA glomerulonephritis and vasculitis. Clin. J. Am. Soc. Nephrol. 2017;12:1680-1691.
Not in our library10.2215/cjn.02500317
225.
Greenan K., Vassallo D., Chinnadurai R., Ritchie J., Shepard K., Green D., Ponnusamy A., Sinha S. Respiratory manifestations of ANCA-associated vasculitis. Clin. Respir. J. 2018;12:57-61.
Not in our library10.1111/crj.12478
226.
Zhang W., Liu H., Al-Shabrawey M., Caldwell R.W., Caldwell R.B. Inflammation and diabetic retinal microvascular complications. J. Cardiovasc. Dis. Res. 2011;2:96-103.
Not in our library10.4103/0975-3583.83035
227.
Baskin E., Bakkaloglu A., Besbas N., Hascelik G., Saatci U., Gök F., Ozen S. Ceruloplasmin levels in antineutrophil cytoplasmic antibody-positive patients. Pediatr. Nephrol. 2002;17:917-919.
Not in our library10.1007/s00467-002-0969-0
228.
Chooklin S., Pereyaslov A., Bihalskyy I. Pathogenic role of myeloperoxidase in acute pancreatitis. Hepatobiliary Pancreat. Dis. Int. 2009;8:627-631.
Not in our library
229.
Klangprapan S., Chaiyarit P., Hormdee D., Kampichai A., Khampitak T., Daduang J., Tavichakorntrakool R., Panijpan B., Boonsiri P. Salivary myeloperoxidase, assessed by 3,3’-diaminobenzidine colorimetry, can differentiate periodontal patients from nonperiodontal subjects. Enzyme Res. 2016;2016:7517928.
Not in our library10.1155/2016/7517928
230.
Edwards S.W., Hallett M.B. Seeing the wood for the trees: The forgotten role of neutrophils in rheumatoid arthritis. Immunol. Today. 1997;18:320-324.
Not in our library10.1016/s0167-5699(97)01087-6
231.
Green P.S., Mendez A.J., Jacob J.S., Crowley J.R., Growdon W., Hyman B.T., Heinecke J.W. Neuronal expression of myeloperoxidase is increased in Alzheimer’s disease. J. Neurochem. 2004;90:724-733.
Not in our library10.1111/j.1471-4159.2004.02527.x
232.
Maki R.A., Tyurin V.A., Lyon R.C., Hamilton R.L., DeKosky S.T., Kagan V.E., Reynolds W.F. Aberrant expression of myeloperoxidase in astrocytes promotes phospholipid oxidation and memory deficits in a mouse model of Alzheimer disease. J. Biol. Chem. 2009;284:3158-3169.
Not in our library10.1074/jbc.m807731200
233.
Choi D.-K., Pennathur S., Perier C., Tieu K., Teismann P., Wu D.-C., Jackson-Lewis V., Vila M., Vonsattel J.-P., Heinecke J.W., et al. Ablation of the inflammatory enzyme myeloperoxidase mitigates features of Parkinson’s disease in mice. J. Neurosci. 2005;25:6594-6600.
Not in our library10.1523/jneurosci.0970-05.2005
234.
Teismann P. Myeloperoxidase in the neurodegenerative process of Parkinson’s disease. Dtsch. Med. Wochenschr. 2014;139:99-102.
Not in our library10.1055/s-0033-1359907
235.
Gray E., Thomas T.L., Betmouni S., Scolding N., Love S. Elevated myeloperoxidase activity in white matter in multiple sclerosis. Neurosci. Lett. 2008;444:195-198.
Not in our library10.1016/j.neulet.2008.08.035
236.
Olza J., Aguilera C.M., Gil-Campos M., Leis R., Bueno G., Martinez-Jiménez M.D., Valle M., Cañete R., Tojo R., Moreno L.A., et al. Myeloperoxidase is an early biomarker of inflammation and cardiovascular risk in prepubertal obese children. Diabetes Care. 2012;35:2373-2376.
Not in our library10.2337/dc12-0614
237.
Demoly P., Crampette L., Mondain M., Enander I., Jones I., Bousquet J. Myeloperoxidase and interleukin-8 levels in chronic sinusitis. Clin. Exp. Allergy. 1997;27:672-675.
Not in our library10.1111/j.1365-2222.1997.tb01195.x
238.
Sagel S.D., Wagner B.D., Anthony M.M., Emmett P., Zemanick E.T. Sputum biomarkers of inflammation and lung function decline in children with cystic fibrosis. Am. J. Respir. Crit. Care Med. 2012;186:857-865.
Not in our library10.1164/rccm.201203-0507oc
239.
Hansberry D.R., Shah K., Agarwal P., Agarwal N. Fecal myeloperoxidase as a biomarker for inflammatory bowel disease. Cureus. 2017;9:e1004.
Not in our library10.7759/cureus.1004
240.
Charin B., Martin N.J.J., Dennis J.H., Witting P.K. Myeloperoxidase in the inflamed colon: A novel target for treating inflammatory bowel disease. Arch. Biochem. Biophys. 2018;645:61-71.
Not in our library10.1016/j.abb.2018.03.012
241.
Malle E., Woenckhaus C., Waeg G., Esterbauer H., Gröne E.F., Gröne H.-J. Immunological evidence for hypochlorite-modified proteins in human kidney. Am. J. Pathol. 1997;150:603-615.
Not in our library
242.
Malle E., Buch T., Gröne H.-J. Myeloperoxidase in kidney disease. Kidney Int. 2003;64:1956-1967.
Not in our library10.1046/j.1523-1755.2003.00336.x
243.
Hanumegowda U.M., Copple B.I., Shibuya M., Malle E., Ganey P.E., Roth R.A. Basement membrane and matrix metalloproteinases in monocrotaline-induced liver injury. Toxicol. Sci. 2003;76:237-246.
Not in our library10.1093/toxsci/kfg222
244.
Serhan C.N., Savill J. Resolution of inflammation: The beginning programs the end. Nat. Immunol. 2005;6:1191-1197.
Not in our library10.1038/ni1276
245.
Li M.O., Flavell R.A. Contextual regulation of inflammation: A duet by transforming growth factor-beta and interleukin-10. Immunity. 2008;28:468-476.
Not in our library10.1016/j.immuni.2008.03.003
246.
Yoshimura A., Wakabayashi Y., Mori T. Cellular and molecular basis for the regulation of inflammation by TGF-β. J. Biochem. 2010;147:781-792.
Not in our library10.1093/jb/mvq043
247.
Chandrasekharan J.A., Sharma-Walia N. Lipoxins: Nature’s way to resolve inflammation. J. Inflamm. Res. 2015;8:181-192.
Not in our library10.2147/jir.s90380
248.
Matzinger P. Tolerance, danger, and the extended family. Annu. Rev. Immunol. 1994;12:991-1045.
Not in our library10.1146/annurev.iy.12.040194.005015
249.
Rother R.P., Bell L., Hillman P., Gladwin M.T. The clinical sequelae of intravascular hemolysis and extracellular plasma hemoglobin. J. Am. Med. Assoc. 2005;293:1653-1662.
Not in our library10.1001/jama.293.13.1653
250.
Kato G.J., Steinberg M.H., Gladwin M.T. Intravascular hemolysis and the pathophysiology of sickle cell disease. J. Clin. Investig. 2017;127:750-760.
Not in our library10.1172/jci89741
251.
Sauret J.M., Marinides G., Wang G.K. Rhabdomyolysis. Am. Fam. Phys. 2002;65:907-912.
Not in our library
252.
Hunter J.D., Gregg K., Damani Z. Rhabdomyolysis. Cont. Ed. Anaesth. Crit. Care Pain. 2006;6:141-143.
Not in our library10.1093/bjaceaccp/mkl027
253.
Jeney V., Balla J., Yachie A., Varga Z., Vercelotti G.M., Eaton J.W., Balla G. Pro-oxidant and cytotoxic effects of circulating heme. Blood. 2002;100:879-887.
Not in our library10.1182/blood.v100.3.879
254.
Lin T., Sammy F., Yang H., Thundivalappil S., Hellman J., Tracey K.C., Warren H.S. Identification of hemopexin as an anti-inflammatory factor that inhibits synergy of hemoglobin with HMGB1 in sterile and infectious inflammation. J. Immunol. 2012;189:2017-2022.
Not in our library10.4049/jimmunol.1103623
255.
Schaer D.J., Buehler P.W., Alayash A.I., Belcher J.D., Vercelotti G.M. Hemolysis and free heme revisited: Exploring hemoglobin and hemin scavengers as a novel class of therapeutic proteins. Blood. 2013;121:1276-1284.
Not in our library10.1182/blood-2012-11-451229
256.
Flemmig J., Schlorke D., Kühne F.-W., Arnhold J. Inhibition of the heme-induced hemolysis of red blood cells by the chlorite-based drug WF10. Free Radic. Res. 2016;50:1386-1395.
Not in our library10.1080/10715762.2016.1252838
257.
Fridovich I. Superoxide radical and superoxide dismutases. Annu. Rev. Biochem. 1995;64:97-112.
Not in our library10.1146/annurev.bi.64.070195.000525
258.
Starkov A.A. The role of mitochondria in reactive oxygen species metabolism and signaling. Ann. N. Y. Acad. Sci. 2008;1147:37-52.
Not in our library10.1196/annals.1427.015
259.
Antonyuk S.V., Strange R.W., Marklund S.L., Hasnain S.S. The structure of human extracellular copper-zinc superoxide dismutase at 1.7 Å resolution: Insight into heparin and collagen binding. J. Mol. Biol. 2009;388:310-326.
Not in our library10.1016/j.jmb.2009.03.026
260.
Low F.M., Hampton M.B., Winterbourn C.C. Prx2 and peroxide metabolism in the erythrocyte. Antioxidants Redox Signal. 2008;10:1621-1630.
Not in our library10.1089/ars.2008.2081
261.
Goyal M.M., Basak A. Human catalase: Looking for complete identity. Prot. Cell. 2010;1:888-897.
Not in our library10.1007/s13238-010-0113-z
262.
Brigelius-Flohé R., Maiorino M. Glutathione peroxidases. Biochim. Biophys. Acta. 2013;1840:3289-3303.
Not in our library10.1016/j.bbagen.2012.11.020
263.
Ponka P. Cellular iron metabolism. Kidney Int. 1999;55:S2-S11.
Not in our library10.1046/j.1523-1755.1999.055suppl.69002.x
264.
Massover W.H. Ultrastructure of ferritin and apoferritin: A review. Micron. 1993;24:389-437.
Not in our library10.1016/0968-4328(93)90005-l
265.
Chiabrando D., Vinchi F., Florito V., Tolosano E., Veas F. Haptoglobin and hemopexin in heme detoxification and iron recycling. Acute Phase Proteins—Regulation and Functions of Acute Phase Proteins. 2011;261-288.
Not in our library
266.
Niki E. Antioxidants in relation to lipid peroxidation. Chem. Phys. Lipids. 1987;44:227-253.
Not in our library10.1016/0009-3084(87)90052-1
267.
Buettner G.R. The pecking order of free radicals and antioxidants: Lipid peroxidation, α-tocopherol, and ascorbate. Arch. Biochem. Biophys. 1993;300:535-543.
Not in our library10.1006/abbi.1993.1074
268.
Hochstein P., Hatch L., Sevanian A. Uric acid: Functions and determinations. Meth. Enzymol. 1984;105:162-166.
Not in our library10.1016/s0076-6879(84)05022-9
269.
Nyyssönen K., Porkkala-Sarataho E., Kaikkonen J., Salonen J.T. Ascorbate and urate are the strongest determinants of plasma antioxidative capacity and serum lipid resistance to oxidation in Finnish men. Atherosclerosis. 1997;130:223-233.
Not in our library10.1016/s0021-9150(96)06064-9
270.
Koj A., Gordon A.H., Koj A. Biological functions of acute-phase proteins. The Acute Phase Response to Injury and Infection. 1985;145-160.
Not in our library
271.
Baumann H., Gauldie J. The acute phase response. Immunol. Today. 1994;15:74-80.
Not in our library10.1016/0167-5699(94)90137-6
272.
Musci G. Structure/function relationships in ceruloplasmin. Adv. Exp. Med. Biol. 1999;448:175-182.
Not in our library10.1007/978-1-4615-4859-1_15
273.
Taggart C.C., Greene C.M., Carroll T.P., O’Neill S.J., McElvaney N.G. Elastolytic proteases. Inflammation regulation and dysregulation in chronic infective lung disease. Am. J. Respir. Crit. Care Med. 2005;171:1070-1076.
Not in our library10.1164/rccm.200407-881pp
274.
Brode S.K., Ling S.C., Chapman K.R. Alpha-1 antitrypsin deficiency: A commonly overlooked cause of lung disease. Can. Med. Assoc. J. 2012;184:1365-1371.
Not in our library10.1503/cmaj.111749
275.
Silverman E.K., Sandhaus R.A. Alpha1-antitrypsin deficiency. N. Engl. J. Med. 2009;360:2749-2757.
Not in our library10.1056/nejmcp0900449
276.
Sandhaus R.A., Turino G., Brantly M.L., Campos M., Cross C.E., Goodman K., Hogart D.K., Knight S.L., Stocks J.M. The diagnosis and management of alpha-1 antitrypsin deficiency in the adult. Chron. Obstr. Pulm. Dis. 2016;3:668-682.
Not in our library10.15326/jcopdf.3.3.2015.0182
277.
Townsend S.A., Edgar R.G., Kantas P.R., Newsome P.N., Turner A.M. Systematic review: The natural history of alpha-1 antitrypsin deficiency, and associated liver disease. Alim. Pharmacol. Ther. 2018;47:877-885.
Not in our library10.1111/apt.14537
278.
Li M.O., Wan Y.Y., Sanjabi S., Robertson A.K., Flavell R.A. Transforming growth factor-beta regulation of immune response. Annu. Rev. Immunol. 2006;24:99-146.
Not in our library10.1146/annurev.immunol.24.021605.090737
279.
Reber A.J., Chirkova T., Kim J.H., Cao W., Biber R., Shay D.K., Sambhara S. Immunosenescence and challenges of vaccination against influenza in the aging population. Aging Dis. 2012;3:68-90.
Not in our library
280.
Sepkowitz K.A. Opportunistic infections in patients with and patients without acquired immunodeficiency syndrome. Clin. Infect. Dis. 2002;34:1098-1107.
Not in our library10.1086/339548
281.
Kampitak T., Suwanpimolkul G., Browne S., Suankratay C. Anti-interferon-γ autoantibody and opportunistic infections: Case series and review of the literature. Infection. 2011;39:65-71.
Not in our library10.1007/s15010-010-0067-3
282.
Singer M., Deutschman C.S., Seymour C.W., Shankar-Hari M., Annane D., Bauer M., Bellamo R., Bernard G.R., Chiche J.-D., Coppersmith C.M., et al. The third international consensus definitions for sepsis and septic shock (sepsis-3). J. Am. Med. Assoc. 2016;23:801-810.
Not in our library10.1001/jama.2016.0287
283.
Podnos Y.D., Jiminez J.C., Wilson S.E. Intraabdominal sepsis in elderly persons. Clin. Infect. Dis. 2002;35:62-68.
Not in our library10.1086/340866
284.
Hospitalized cancer patients with severe sepsis: analysis of incidence, mortality, and associated costs of care · 2004
No notice
285.
Jiminez M.F., Watson R.W., Parodo J., Evans D., Foster D., Steinberg M., Rotstein O.D., Marshall J.C. Dysregulated expression of neutrophil apoptosis in the systemic inflammatory response syndrome. Arch. Surg. 1997;132:1263-1270.
Not in our library10.1001/archsurg.1997.01430360009002
286.
Demaret J., Venet F., Friggeri A., Cauzalis M.-A., Plassais J., Jallades L., Malcus C., Poitevin-Later F., Textoris J., Lepape A., et al. Marked alterations in neutrophil functions during sepsis-induced immunosuppression. J. Leukoc. Biol. 2015;98:1081-1090.
Not in our library10.1189/jlb.4a0415-168rr
287.
Brown K.A., Brain S.D., Pearson J.D., Edgeworth J.D., Lewis S.M., Treacher D.F. Neutrophils in development of multiorgan failure in sepsis. Lancet. 2006;368:157-169.
Not in our library10.1016/s0140-6736(06)69005-3
288.
Kovach M.A., Standiford T.J. The functions of neutrophils in sepsis. Curr. Opin. Infect. Dis. 2012;25:321-327.
Not in our library10.1097/qco.0b013e3283528c9b
289.
Crouser E., Exline M., Wewers M.D. Sepsis: Links between pathogen sensing and organ damage. Curr. Pharmaceut. Des. 2008;14:1840-1852.
Not in our library10.2174/138161208784980572
290.
Rittirsch D., Flierl M.A., Ward P.A. Harmful molecular mechanisms in sepsis. Nat. Rev. Immunol. 2008;8:776-787.
Not in our library10.1038/nri2402
291.
Maruchi Y., Tsuda M., Mori H., Takenaka N., Gocho T., Huq M.A., Takeyama N. Plasma myeloperoxidase-conjugated DNA level predicts outcomes and organ dysfunction in patients with septic shock. Crit. Care. 2018;22:176.
Not in our library10.1186/s13054-018-2109-7
292.
Myeloperoxidase can differentiate between sepsis and non-infectious SIRS and predicts mortality in intensive care patients with SIRS · 2017
Not checked
293.
Kothari N., Keshari R.S., Bogra J., Kohli M., Abbas H., Malik A., Dikshit M., Barthwal M.K. Increased myeloperoxidase enzyme activity in plasma is an indicator of inflammation and onset of sepsis. J. Crit. Care. 2011;26:435.e1-453.e7.
Not in our library10.1016/j.jcrc.2010.09.001
294.
Cha Y.S., Yoon J.M., Jung W.J., Kim Y.W., Kim T.H., Kim O.H., Cha K.C., Kim H., Hwang S.O., Lee K.H. Evaluation of usefulness of myeloperoxidase index (MPXI) for differential diagnosis of systemic inflammatory response syndrome (SIRS) in the emergency department. Emerg. Med. J. 2015;32:304-307.
Not in our library10.1136/emermed-2013-203137

Cited by papers we hold 1

Phospholipases and Reactive Oxygen Species Derived Lipid Biomarkers in Healthy and Diseased Humans and Animals – A Focus on Lysophosphatidylcholine · 2021
Not checked

Reads

Read by people on this page

None yet.

Checked or read by AI through naulon

None yet.

A person counts once a day per paper. We keep no cookie and no address: a reader is a hash that changes every day. Any month under five reads shows no number.

Are you an author?

Is this your paper?

Claim it to add your note and be paid when agents cite it. Free.