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Logo of nihpaAbout Author manuscriptsSubmit a manuscriptNIH Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
Curr Protoc Toxicol. Author manuscript; available in PMC Aug 1, 2012.
Published in final edited form as:
Curr Protoc Toxicol. Aug 2011; CHAPTER: Unit7.9.
doi:  10.1002/0471140856.tx0709s49
PMCID: PMC3156475
NIHMSID: NIHMS316456

Overview of Peroxiredoxins in oxidant defense and redox regulation

Abstract

Peroxiredoxins are important hydroperoxide detoxification enzymes, yet have only come to the fore in recent years relative to the other major players in peroxide detoxification, heme-containing catalases and peroxidases, and glutathione peroxidases. These cysteine-dependent peroxidases exhibit high reactivity with hydrogen peroxide, organic hydroperoxides and peroxynitrite and play major roles not only in peroxide defense, but also in regulating peroxide-mediated cell signaling. This overview focuses on important peroxiredoxin features that have emerged over the past several decades with an emphasis on catalytic mechanism, regulation and biological function.

Keywords: peroxidases, antioxidants, antioxidant enzymes, sulfenic acids, hydroperoxides, hydrogen peroxide, thiol peroxidase, Prx, PRDX, redox regulation

Introduction

Peroxiredoxins (Prxs) are a fascinating group of thiol-dependent peroxidases (EC 1.11.1.15) which detoxify H2O2, aliphatic and aromatic hydroperoxides, and peroxynitrite (Dubuisson, et al., 2004, Flohé, et al., 2010, Poole, 2007). They are ubiquitously expressed, with multiple isoforms present in most organisms (e.g., 3 isoforms in Escherichia coli, 5 in Saccharomyces cerevisiae, 6 in Homo sapiens and 9 in Arabidopsis thaliana) (Dietz, 2011, Knoops, et al., 2007). Many exhibit very fast rates of peroxide reduction on the order of 106 to 108 M−1 s−1 using a conserved active site architecture that is highly specialized for peroxide reduction, as their reactivity with other thiol reagents is only modest (Cox, et al., 2009, Dubuisson, et al., 2004, Horta, et al., 2010, Manta, et al., 2009, Nelson, et al., 2008, Parsonage, et al., 2010, Parsonage, et al., 2005, Peskin, et al., 2007, Stacey, et al., 2009, Trujillo, et al., 2007). While the pKa of the active site Cys is strongly influenced by electrostatic environment and is lowered in Prxs to promote thiolate formation (values around or below ~6), this feature is quite insufficient to explain the rapid catalytic rates (Flohé, et al., 2010, Manta, et al., 2009, Nelson, et al., 2008, Winterbourn, 2008). The high catalytic efficiency as well as general high abundance of Prx protein in cells makes them the predominant scavengers of peroxides in many circumstances (Adimora, et al., 2010, Winterbourn, 2008), and it is increasingly recognized that they play major roles in the detoxification of and defense against potentially damaging oxidants, the same oxidants that regulate and mediate cell signaling processes (Flohé, 2010, Fomenko, et al., 2008, Hall, et al., 2009, Poole and Nelson, 2008).

The structural context and biophysical properties that underlie Prx function are also fascinating aspects of this group of proteins. Prxs are built on a thioredoxin (Trx) scaffold, like glutathione peroxidases (Gpxs), and rely on an active site cysteine within a PxxxTxxC motif for catalysis. Structural and bioinformatic evidence supports the idea that the canonical CxxC redox motif in Trxs and glutaredoxins (Grxs) (with the first Cys acting as the nucleophile in thiol-disulfide interchange) has diverged to become both the TxxC motif of Prxs and the CxxT (or UxxT, with U = selenocysteine) of Gpxs (Fomenko and Gladyshev, 2003). This change results in the gain of peroxidase activity and the loss of the more general protein disulfide reductase functions. Intriguingly, one additional residue required for catalysis which lies outside the active site loop/helix region, an Arg residue contributing to the Prx active site from about 75 residues away in sequence, is in a position equivalent to that of the conserved cis-Pro in the reductases (Copley, et al., 2004, Su, et al., 2007).

Bioinformatics tools applied to gain insight into the key amino acids and interactions underlying catalytic function have also provided information about the subfamilies into which members of this broad and diverse Prx family can be subdivided (Copley, et al., 2004, Nelson, et al., 2011). Structural information is also increasingly available and has provided additional opportunities to investigate the important features of Prxs (Hall, et al., 2011).

This unit provides an overview of the important Prx features that have emerged over the past several decades with an emphasis on catalytic mechanism, regulation and biological function.

The common and distinct features of peroxide reduction and enzyme recycling among the Prx subfamilies

The Catalytic Cycle

Accumulating information in the early 90’s pointed to the essentiality of a single Cys residue for catalysis of peroxide reduction by Prxs. It was noted that a second Cys was often, but not always, present and conserved near the C-terminus, suggesting a mechanistic distinction between these “2-Cys” and “1-Cys” groups of Prx enzymes (Chae, et al., 1994). We now recognize the absolutely conserved Cys residue as the “peroxidatic” Cys (denoted Cp, or Sp for the sulfur atom) that attacks the hydroperoxyl substrate, forming the first product (water or alcohol in the case of H2O2 or larger ROOH substrates, respectively) and a sulfenic acid moiety on the active site Cp residue (Fig. 1). This chemistry matches the now well established mechanism of H2O2 reduction by the single-Cys containing NADH peroxidase flavoenzymes from gram positive lactic acid bacteria (Crane, et al., 1997, Poole and Claiborne, 1989, Yeh, et al., 1996).

Figure 1
The catalytic cycle of Prx. The three main chemical steps of catalysis are (1) peroxidation, forming the sulfenic acid at the peroxidatic cysteine (SpOH), (2) resolution which generates a disulfide bond, and (3) recycling by reduction to return the enzyme ...

In a minimal catalytic mechanism for Prxs, sulfenic acid is “captured” (or resolved) by a thiol group, generating H2O as the second product and a disulfide bond in the enzyme on the pathway to reductive recycling (Fig. 1). If the thiol group comes from the Prx, this Cys residue is called the “resolving” Cys or Cr; in the earliest studies of Prxs (Chae, et al., 1994, Chae, et al., 1994), this residue within the C-terminus was noted to come from another subunit of a dimer, generating an intersubunit disulfide bond with the Cp (two active sites and two disulfides per α2 dimer). In the subsequent years it has been increasingly recognized that the Cr can be contributed from a number of other positions within the same subunit, as well (sometimes designated the “atypical” 2-Cys Prxs) (Wood, et al., 2003). In the 1-Cys Prxs, the resolving thiol must come from a non-Prx molecule which may be another protein or even a small molecule reductant like glutathione. While this 1-Cys mechanism was first described for human PrxVI1, the first Prx to be structurally characterized (Choi, et al., 1998), it is now recognized to apply to Prxs within other subfamilies that lack a Cr, as well.

The most common reductant used for recycling by members of all subfamilies of Prxs is Trx (where both R” SH moieties of Fig. 1 come from a single Trx CxxC active site). Other reductases similar to Trx (e.g. tryparedoxin from kinetoplastids, the Grx-like Cp9 of Clostridium pasteurianum, the N-terminal domain of bacterial AhpF, and plant NTR) are also found to act as specialized Prx reductases in certain organisms (Dietz, 2011, Flohé, et al., 2010, Poole, et al., 2000). The nature of recycling of human PrxVI and many other 1-Cys Prx enzymes in vivo continues to be a matter of some debate; evidence has been reported for the involvement of glutathione and glutathione transferase pi (GSTπ), lipoic acid, ascorbate, cyclophilins, and reductases including Grxs and Trxs (Dietz, 2011, Flohé, et al., 2010).

Localized unfolding and refolding for catalysis

Resolution of the Prx sulfenic acid, regardless of mechanism, requires a localized unfolding of structures around the Cp (often accompanied by structural changes around the Cr, when present) that removes the SpOH from the “fully folded” (FF) active site and exposes it (in the “locally unfolded” or LU form) for disulfide bond formation with Cr or another resolving thiol group. This essential step occurs with varying efficiencies and can lead to the persistence of the SOH within the FF active site of some Prxs, providing an opportunity for oxidative regulation during the catalytic cycle (Fig. 1), an issue which is discussed in more detail below.

Prx subfamilies

Based on the variation in the location and even presence of Cr in different Prxs as well as notable structural distinctions between them, it is clear that different classes of proteins have arisen during Prx evolution, all of which maintain the common peroxide reduction chemistry and active site architecture, but vary in the details of their recycling pathways (Hall, et al., 2011). Moreover, features such as oligomeric state and propensity toward oxidative inactivation during turnover also vary among different Prxs. To better explore the distinctions between such subclasses and take advantage of the large amounts of genomic sequence information currently available, various bioinformatics approaches have been applied (Copley, et al., 2004, Dietz, 2011, Knoops, et al., 2007, Koua, et al., 2009). One recent study (Nelson, et al., 2011) utilized an approach known as functional site (or active site) profiling to identify >3500 Prx sequences from the Jan 2008 release of GenBank, each classified unambiguously into one of six Prx subfamilies that had been previously identified by other bioinformatic and structural analyses (Table 1). The strength of this approach is that classification is based not on full sequence alignments but on the conservation of conserved sequences proximal to the active site. The generation of large lists of Prx sequences with known subfamily membership, now available in a publically accessible database (http://csb.wfu.edu/prex/) (Soito, et al., 2011), allowed for detailed analyses of residue conservation, phylogenetic distribution and overall conservation patterns for the Cr of each subfamily, and some of the most relevant results from this work are described in the following paragraphs.

Table I
Summary of the structural and bioinformatic analyses of Prx subfamilies

The “typical 2-Cys” group of Prxs comprised of the founding members of the 2-Cys Prxs described above (denoted here as the Prx1/AhpC subfamily) is a much studied and broadly distributed group of proteins that form intersubunit disulfide bonds during catalysis. This group includes as members human PrxI through IV and bacterial “AhpC” proteins (named for their alkyl hydroperoxide reductase activity). Members of both the Prx1/AhpC subfamily and the Prx6 subfamily contain a C-terminal extension relative to the ancestral Trx fold and to other Prx subfamilies (Table 1), a feature which has led to their being grouped together by some bioinformatic analyses (Copley, et al., 2004, Karplus and Hall, 2007). Despite this, active site profiling clearly distinguishes the two subfamilies. For the 2-Cys proteins in the Prx1/AhpC subfamily, the Cr is found within this C-terminal extension; although Prx6 subfamily members do in a very few cases possess a putative Cr, the majority appear to lack a Cr (Table 1, see below). Members of the Prx1/AhpC subfamily form not only α2 dimers, brought together through interactions at the edges of their β sheets (the so-called B-type interface), but also higher order oligomeric structures [primarily (α2)5 decamers] built up from an alternative, A-type interface (Fig. 2) (Hall, et al., 2011). This A-type interface plays an important role in supporting catalysis in Prx1/AhpC proteins, as described in more detail below.

Figure 2
Oligomeric interfaces and quanternary structures of Prxs. Dimerization of Prx subunits can occur through two different types of interfaces, the A-type interface (as observed for Tpx and Prx5 subfamily members), or the B-type interface which brings the ...

Two thiol-dependent peroxidases described in the mid- to late 90’s, human PrxV and E. coli thiol peroxidase (Tpx), became founding members of two distinct Prx subfamilies (Baker and Poole, 2003, Cha, et al., 1995, Knoops, et al., 1999, Sarma, et al., 2005, Seo, et al., 2000). Members of these subfamilies are typically dimeric and form intrasubunit disulfide bonds (Table 1). Interestingly, the interface used for dimer formation in these enzymes is essentially the same A-type interface used to form higher order oligomeric structures in Prx1/AhpC proteins (Fig. 2).

Two other subfamilies have also emerged, one of which is relatively broadly distributed in biology, though lacking in animals (the BCP/PrxQ group), and the other (AhpE) which is a recent addition to the Prx family and is only narrowly represented among species related to that of the founding member, Mycobacterium tuberculosis (Table 1). Within the BCP/PrxQ subfamily of proteins, the low potential PrxQ proteins of plant chloroplasts have received much attention and likely have a function in photosynthesis within this organelle (Dietz, 2011). The representative from E. coli, BCP (which stands for bacterioferritin comigratory protein), seems to be less active as a peroxidase than most Prxs (Jeong, et al., 2000), though the functional importance of this protein has been demonstrated, at least in the human pathogen Helicobacter pylori, where it helps establish long term infections within the host’s gastric mucosa (Wang, et al., 2005). Members of this group have generally been thought to be monomeric (Nelson, et al., 2011) though recently added structural representatives show that some members can form dimers using the A-type interface used to form dimers in the Tpx and Prx5 subfamilies (Hall, et al., 2011). The few AhpE representatives do not clearly fit into any other Prx subfamilies (Hall, et al., 2011, Nelson, et al., 2011), and only one representative from M. tuberculosis has been structurally and kinetically characterized as of December 2010. This protein has a significant peroxynitrite reductase activity on the order of 107 M−1 s−1 that dominates somewhat over its peroxidase activity (Hugo, et al., 2009), and forms an A-type dimer (Li, et al., 2005). Although M. tuberculosis AhpE utilizes a 1-Cys mechanism, other members of this subfamily contain a potential Cr (Nelson, et al., 2011).

Location and prevalence of Cr in Prx subfamilies

For 2-Cys Prxs, the step of catalysis after peroxide reaction is resolution of the Cp-sulfenic acid by condensation with a thiol group provided by another Cys in the enzyme (Cr, or Sr in Fig. 1), forming either an intra- or intersubunit disulfide bond. As mentioned above, a remarkable feature of the Prxs is how many different sites have been shown to contain a functionally active Cr. In two of the subfamilies, Prx1/AhpC and Tpx, the distinct location of Cr is highly conserved (within the C-terminus and helix α3, respectively), with >96% of all members conforming to this pattern (Table 1) (Hall, et al., 2011, Nelson, et al., 2011). For Prx6 and AhpE subfamily members, the Cr may occur only rarely, with the majority of subfamily members functioning as 1-Cys Prxs. For BCP/PrxQ and Prx5 subfamilies, the presence and/or location of Cr is more variable. The Cr of human PrxV, within helix α5, is in fact relatively uncommon in this group of proteins, with only 17% of members (mostly metazoans) possessing it; another 16% of Prx5 members, all from bacteria, are fused to a Grx domain as observed for the canonical Grx/Prx hybrid protein from Haemophilus influenza (Nelson, et al., 2011). Among BCP/PrxQ proteins, a slight majority (55%) have Cr located ~1 ½ turns down the α2 helix from Cp; another subset of subfamily members (7%) are now recognized to use an alternate position for Cr, in helix α3 (as is usually observed for Tpx proteins). These observations support the hypothesis that the Cr has arisen independently multiple times during the evolutionary divergence of the Prxs and also support the notion that the BCP/PrxQ group of proteins are the most representative of an ancestral form of the Prxs (Copley, et al., 2004, Hall, et al., 2011).

Chemical and structural features promoting catalysis in peroxiredoxins

While the chemistry of a thiolate group attacking the −O–O− bond of a hydroperoxide substrate seems simple at first glance, the forces that drive and regulate catalysis are many and sometimes contradictory. For example, thiolates (R-S) are better nucleophiles than thiols (R-SH); the reactivity of the latter with hydroperoxides is vanishingly low. Therefore, a pKa for the Cp of one or several pH units below the expected, unperturbed pKa of around 8.5 would be considered supportive of catalysis through enhanced ratios of thiolate to thiol at physiological pHs. For Prxs, measured pKa values have ranged from 5.2 to 6.3 (Table 2) (Horta, et al., 2010, Hugo, et al., 2009, Manta, et al., 2009, Nelson, et al., 2008, Ogusucu, et al., 2007). The complication of lowering the active site pKa is, however, the fact that a low pKa thiolate is a worse nucleophile than a high pKa thiolate, so that stabilization of the negative-charged thiolate would seem to come at the expense of chemical reactivity. However, Prxs, which were once thought to exhibit rather poor activity, have more recently been noted for their quite high reactivities with hydroperoxide substrates, ranging upwards of 107 – 108 M−1 s−1, values which are within the realm of the highly reactive selenocysteine-containing glutathione peroxidases (Cox, et al., 2009, Horta, et al., 2010, Manta, et al., 2009, Parsonage, et al., 2005, Peskin, et al., 2007, Trujillo, et al., 2007). How then can the Prx active site impart such high reactivity to this sulfur-containing residue?

Table 2
Summary of measured pKa values for the Cp of Prxs

Part of the answer seems to lie in the observations that the charge stabilization and hydrogen-bonding features of the active site not only promote thiolate stability but also position and thus activate the peroxide to be attacked. The active site geometry is also set up for stabilization of the transition state wherein the “OA”, the terminal oxygen of the hydroperoxide, makes favorable interactions for weakening the bond to the “OB” of the peroxide and for forming a new bond to the Cp sulfur atom (Fig. 3). Based on a large scale analysis of over 70 Prx structures and emerging examples or mimics of substrate and product complexes, the OA can be envisioned as sliding along a hydrogen bond-guided “track” during catalysis, starting from a position covalently bonded to OB 1.5 Å apart, and ending up attached to the Sp of the Cys, ~3 Å from OB (Hall, et al., 2011, Hall, et al., 2010). This track is reasonably well defined structurally given the many ligand-containing Prx structures where one or two oxygen atoms occupy positions within the track that mimic OA and/or OB. In some cases where carboxylates and diols are bound, such as the high resolution benzoate and dithiothreitol (DTT) complexes with human PrxV (PDB identifiers 1hd2 and 3mng, respectively), two oxygens are bound along the track with atomic separations of ~ 2.4 Å, suggesting that these types of ligands act as transition-state analogs, sampling intermediate positions along the linear oxygen track created within the Prx active site (Hall, et al., 2010).

Figure 3
Interactions around the Prx active site with bound hydroperoxide substrate. The stabilizing hydrogen bonding interactions (dotted lines) between key atoms from the backbone and the four conserved residues, and with the ROOH substrate, are indicated. The ...

As our view of the Prx active site has sharpened, the roles for the highly conserved residues within the active site have become clearer based on these analyses: (1) the hydroxyl group of Thr (or in rare cases Ser) positions and activates the protonated OA atom of the incoming hydroperoxide substrate; (2) the Arg, through both charge and hydrogen-bonding interactions, positions and activates both the active site thiolate and the peroxide substrate; and (3) the Pro shields the active site from unwanted reactions and positions the backbone of the following two residues to provide additional hydrogen bonding interactions (Fig. 2) (Hall, et al., 2010).

Regulatory Oxidation and Other Post-translational Modifications

As Prxs were studied in increasing molecular detail, it became apparent that the active sites of some, particularly those from higher organisms, are surprisingly susceptible to oxidative inactivation by their own substrate (Baker and Poole, 2003, Rabilloud, et al., 2002, Yang, et al., 2002). The enzyme-associated product of this oxidation, cysteine sulfinic acid (R-SpO2H), is stabilized within the Prx active site (e.g., in human PrxII as captured by crystallography in 1qmv), although at least a few Prxs can undergo yet another hyperoxidation step to form the cysteine sulfonic acid (R-SpO3H) (Sarma, et al., 2005, Seo, et al., 2009) (Fig. 1). Using these species, a model could be generated whereby the oxygen of the sulfenic acid product from catalysis rotates so that another lone pair of electrons on the sulfur can again attack a hydroperoxide substrate within the fully folded enzyme active site; a further rotation would permit sulfonic acid formation in those enzymes for which the geometry is suitable (Sarma, et al., 2005). While both sulfinic and sulfonic acids were long believed to be irreversibly oxidized forms of sulfur within biological systems, new findings in the early 2000’s demonstrated the reversibility of sulfinic acid formation in a subset of Prxs that can be repaired by the enzyme sulfiredoxin (Srx) (Biteau, et al., 2003, Lowther and Haynes, 2011). The existence of a repair system that seems to be dedicated to Prx recovery from oxidative inactivation implies an important biological role for this inactivation; the nature of how this oxidative regulatory cycle adds to the functional properties of this family of proteins is a topic of much discussion (see below). As sulfinic acids other than the Cp sulfinic acid of some Prxs have not yet been demonstrated to serve as substrates of Srx proteins, this appears to be very specialized chemistry that may only pertain to this group of enzymes.

Oxidative inactivation during turnover at submillimolar levels of H2O2 is observed for a number of eukaryotic Prxs, but is rare among prokaryotic Prxs (Hall, et al., 2009, Wood, et al., 2003). A structural explanation for the sensitivity lies in the architecture of the protein surrounding the active site, at least in the case of the Prx1/AhpC group of Prxs which form intersubunit disulfide bonds during the catalytic cycle. In some of the sensitive Prx1-like proteins, the FF form of the active site is stabilized by interactions between the active site surrounding the Cp and two regions in the protein with conserved sequences. Stabilization of the FF form of the active site by these surrounding interactions causes the sulfenic acid formed upon peroxide reduction to persist longer within the active site, where a second peroxide can bind and further oxidize it. The first stabilizing interaction involves a proximal GGLG-containing segment following a 3/10 helix, from the same subunit as the Cp, and the second interaction is with the C-terminal tail extending past the Cr of a second subunit, including a conserved YF motif within the helix (present in Prx1-like proteins but not bacterial AhpCs). Several lines of evidence support the role of these two regions in hyperoxidation. For example, the shorter C-terminus and absence of the inserted GGLG in bacterial AhpC support a much more flexible active site compared with human PrxII, a sensitive Prx, and local unfolding of the active site is more favorable (Wood, et al., 2003). In this way, intersubunit disulfide bond formation is facilitated, locking in the LU conformation and removing the reactive sulfenic acid, thereby avoiding hyperoxidation. This emphasizes the important role for Cr in protection against oxidative inactivation (Ellis and Poole, 1997, Trujillo, et al., 2006). Where investigators have introduced single amino acid substitutions into sensitive Prxs (Koo, et al., 2002) or swapped C-terminal tails of sensitive and robust Prxs by mutagenesis (Sayed and Williams, 2004), predictable effects on hyperoxidation sensitivity were observed, consistent with a linkage between the propensity toward hyperoxidation during catalysis and C-terminal tail packing over the active site. Thus, while stabilization of the FF active site should promote the peroxidase reaction to a point, further stability promotes hyperoxidation, providing a “rheostat” for oxidative regulation through the Cp sulfenic acid formed during catalysis. Through evolution, this feature can therefore be “tuned” to fit the need corresponding to its biological function (Poole, et al., 2004). Interestingly, new findings suggest that further oxidation to form the sulfonic acid at Cp is also observed primarily for PrxI, but not PrxII (which becomes N-terminally acetylated in cells, unlike PrxI); this irreversible oxidation may be important as an additional regulatory step that prevents recovery through Srx-dependent repair (Seo, et al., 2009).

Other controls on catalytic efficiency and/or hyperoxidation sensitivity also involve posttranslational modifications that are probably only partially recognized at this point, though the list is steadily growing [reviewed in (Aran, et al., 2009)]. Briefly, cell cycle-dependent phosphorylation of Thr90 in PrxI was observed to occur during mitosis through action of CDK1, and this modification led to an overall decrease in activity (Chang, et al., 2002). A later paper identified a new site of phosphorylation on PrxI, Tyr194, as a modification occurring to membrane-associated PrxI, inactivating it under conditions where oxidative signaling may be occurring (see below). PrxI and II were also shown to accumulate as acetylated proteins in cells depleted of HDAC6, and this modification occurred at the penultimate residue, a Lys, within the C-terminal tail of these proteins (Parmigiani, et al., 2008). Acetylation reportedly enhances the activity of Prxs and increases their resistance to hyperoxidation (Parmigiani, et al., 2008), as does C-terminal truncation (Jara, et al., 2008, Koo, et al., 2002, Seo, et al., 2004), consistent with disruption of the packing of this region over the active site which in turn promotes reductive recycling (see next section).

Interplay between oligomerization and catalysis

As X-ray structures and biophysical characterizations of Prxs were emerging in the early 2000’s, it became clear that there was an unusual link between oligomeric state, redox status and catalysis by Prxs in the AhpC/Prx1 subfamily (i.e. typical 2-Cys Prxs) (Hall, et al., 2009, Wood, et al., 2002). While one might expect a disulfide bond to be stabilizing in terms of higher order structures, this disulfide between the Cp and Cr in the oxidized Prxs (across the B-type interface) instead destabilizes the decameric structures observed in the reduced and hyperoxidized enzymes (represented by the tryparedoxin peroxidase and human PrxII structures, 1uul and 1qmv, respectively) and predominantly yields dimers (as seen in rat PrxI, 1qq2). This transition was suggested initially based on differences between some of the early crystal structures, and was subsequently confirmed through analytical ultracentrifugation studies of bacterial AhpC (Wood, et al., 2002). Disulfide bond formation not only destabilizes decamers, it also results in mobilization of the extended C-termini. The disorder in this region of the protein is responsible for a lack of electron density within crystals of proteins in LU conformation (e.g., oxidized Prx1 and AhpC proteins, 1qq2 and 1yep) for ~20 or more C-terminal residues (Hirotsu, et al., 1999, Wood, et al., 2002). Not surprisingly, the Prx1/AhpC proteins which are hyperoxidized and cannot form the decamer-disrupting disulfide bond are also stabilized as decamers in solution.

The structural explanation for this link between active site conformation and oligomerization lies in the relative proximity of the active site to the decamer building (A-type) interface. Local unfolding of the active site, which gets “locked in” by disulfide bond formation, results in a loss in stability for the decamer, because the active site loop in the FF conformation acts to buttress the decamer-building interface. Similarly, one would expect that decamer formation and the buttressing effect of adjacent dimers would contribute to the stability of the FF form, potentially affecting catalytic activity. This hypothesis was tested through the introduction of decamer destabilizing mutations into Salmonella typhimurium AhpC. Disrupting mutations (e.g. T77I and T77D mutations) not only promoted dimer rather than decamer formation in both oxidized and reduced forms of AhpC, but also had the effect of decreasing the catalytic efficiency of H2O2 reduction by about 2 orders of magnitude, with most of the effect on Km for peroxide (Parsonage, et al., 2005). The enzymatic activity of the primarily dimeric mutants was, however, less sensitive toward hyperoxidation during turnover, as predicted based on greater flexibility around the active site that would be caused by the absence of an adjacent dimer. Computational work has also supported the importance of decamer formation for catalytic function and the electrostatic interactions around the active site that influence Cp pKa; decamer formation restricts the conformations available to some amino acid side chains which are expected to have direct or indirect influences on Cp reactivity in the 2-Cys Prxs of this group (Yuan, et al., 2010).

Peroxide defense and cell signaling control functions for Prxs

The biological role of E. coli and other bacterial AhpCs as oxidant defense enzymes is well established (Imlay, 2008) and protection of cellular macromolecules from oxidative damage is likely to be an important role for most if not all Prxs. The essentiality of Prxs in biology is highlighted by their ubiquity and abundance, as one or more representative(s) seem to be highly expressed in most organisms, with the exception of some spirochetes (i.e., those closely related to Borrelia burgdorferi) (Parsonage, et al., 2010, Winterbourn, 2008). However, the existence of multiple means to regulate the peroxidase activity of Prxs, particularly within higher organisms, indicates that this relatively simple “oxidant defense” view doesn’t tell the whole story. Indeed, it is becoming more and more clear in higher organisms that localized reactive oxygen species (ROS) production, and particularly H2O2, is intimately linked to many cell signaling pathways, and the enzymatic control of the levels of these oxidants must clearly be a key factor in their effectiveness as mediators of cell signaling processes (Flohé, 2010, Rhee, et al., 2005, Wood, et al., 2003). This highlights the inescapable overlap between the concepts of oxidative stress and redox signaling, which are often taken to represent different aspects of the redox “continuum”. Cancer cells, which often exhibit high internal oxidant levels (Szatrowski and Nathan, 1991), are also highly proliferative and in some respects resistant to the effects of oxidative damage. In fact, one route for bypassing oxidant-induced apoptosis is through the deletion of the gene for p53, a redox-sensitive transcriptional regulator that is commonly lacking in aggressive cancers (Fridman and Lowe, 2003). Cancer cells also typically exhibit an enhanced arsenal of antioxidant defense enzymes, many of which are among the repertoire under the control of Keap1/Nrf2 signaling (Kim, et al., 2008). A number of studies have linked high Prx levels with the cancerous phenotype (Lehtonen, et al., 2004, Quan, et al., 2006, Roumes, et al., 2010), with radiation resistance in cell lines (Lee, et al., 2008, Smith-Pearson, et al., 2008, Wang, et al., 2005, Zhang, et al., 2009), and with poor prognosis for chemotherapy (Iwao-Koizumi, et al., 2005, Kim, et al., 2008, Pak, et al., 2011). Nonetheless, while cancer cells may better cope with oxidants given elevated Prx expression, whole animal knockouts of PrxI, in particular, yield mice which are predisposed to development of lymphomas, sarcomas and carcinomas (Neumann, et al., 2003). Thus, Prx proteins may act in different ways, both as tumor suppressors, since their absence leads to cancer, and as tumor promoters, where they provide important defenses against oxidative damage within the tumor enivroment (Neumann and Fang, 2007).

Oxidant defense provided by Prx activity does have an impact on cell signaling events, as shown most clearly in unicellular organisms. Prokaryotic, peroxide-sensing transcription factors which orchestrate the expression of defense enzymes illustrate a direct linkage between oxidant levels and signaling, while the more complex peroxide-signaling pathways in eukaryotes are likely to be regulated by Prxs in more subtle ways. Two families of bacterial transcription factors with peroxide-reactive Cys residues analogous to Cp of Prxs, OxyR and OhrR, are directly regulated by their small molecule effectors, H2O2 and organic hydroperoxides, respectively, using chemistry that parallels the catalytic cycle of Prxs, but with a much slower reduction step (Antelmann and Helmann, 2010, Imlay, 2008). The regulatory action of Prxs in such model organisms as E. coli seems well accounted for by their capacity to reduce and remove these oxidants, at least in terms of AhpC activity (Yamamoto, et al., 2008).

Nonetheless, Prxs sensitive to overoxidation by certain substrates also exist within prokaryotes including E. coli [Tpx (Baker and Poole, 2003)] and cyanobacteria [Anabaena 2-Cys Prx (Pascual, et al., 2010)], suggesting more complexity to the roles of Prxs in these organisms, as well. In yeast, the transcription factors like Pap1 from Schizosaccharomyces pombe and Yap1 from Saccharomyces cerevisiae do not directly sense peroxides, but instead interact through thiol-disulfide exchange pathways with Prx and Gpx homologues to transmit the oxidation “signal” sensed by these peroxidases (Antelmann and Helmann, 2010, D’Autreaux and Toledano, 2007, Klomsiri, et al., 2010, Veal, et al., 2007). This ability to be reduced by alternative partners requires special features within the “sensor” peroxidases that allow for a kinetic pause in the catalytic cycle after sulfenic acid formation, thereby promoting disulfide bond formation with Cys residues from other proteins. In fact, recent studies of S. cerevisiae where all 8 thiol-dependent peroxidases (5 Prxs and 3 sulfur-containing Gpxs) were knocked out demonstrated a profound loss in sensitivity of transcriptional activity toward H2O2 in the absence of these proteins, highlighting their dual roles as defense enzymes and signal transduction intermediates (Fomenko, et al., 2011).

The roles played by Prx proteins in cell signaling processes in higher organisms are under intense investigation, and although few would argue that Prxs are not important regulators of cell signaling, the molecular details of how they exert their specific effects and when and where they are important players are the subject of a burgeoning literature (D’Autreaux and Toledano, 2007, Flohé, 2010, Forman, et al., 2010, Hall, et al., 2009, Winterbourn, 2008). For the purpose of this overview, some of the major points of discussion are summarized herein, leaving it to the interested reader to seek further depth to the arguments from the literature.

One major feature of Prx regulation that has gained considerable attention is sensitivity to hyperoxidation caused by turnover of these enzymes in the presence of relatively high levels of peroxide substrates, as discussed above. The idea that sensitivity toward hyperoxidation plays any role in biology is supported by several arguments: (i) sensitivity is an evolved feature which is readily dampened by mutation or modification, (ii) an ATP-dependent enzymatic repair system exists to reverse this inactivating modification, and (iii) hyperoxidation of Prxs can occur within various cell types after stimulation, being caused, for example, by treatment with tumor necrosis factor alpha, 6-hydroxydopamine or expression plasmids for elevating lipoxygenase and cyclooxygenase expression (Cordray, et al., 2007, Lee, et al., 2008, Rabilloud, et al., 2002). In 2003, Karplus and Poole proposed the floodgate hypothesis, suggesting that this sensitivity coevolved within abundant cellular Prx proteins in concert with the need for localized peroxide production to drive redox-dependent signaling processes (Wood, et al., 2003). By switching “off” the abundant and highly active Prxs around sites of rapid ROS production, like those with activated NADPH oxidase complexes, the oxidants produced could reach higher levels within these foci that could then permit the chemical oxidation of other protein targets involved in promoting or controlling signaling, processes which would normally be much too slow to compete with oxidation of the reactive Prxs. Alternative roles for hyperoxidation have also been proposed, including promotion of a chaperone-like function that accompanies aggregation beyond the level of decamer formation (Jang, et al., 2004, Moon, et al., 2005). The sulfinic (or sulfonic) acid form(s) could also (or instead) act as signals sensed in some way by the cell to elicit appropriate responses. Such a role was suggested by the appearance of aggregates of hyperoxidized PrxII protein within cells that correlated with cell cycle arrest caused by a low grade, continuous exposure to H2O2 (Phalen, et al., 2006). After removal of the oxidant, recovery of the non-aggregated forms of PrxII also correlated with the resumption of the normal cell cycle. While the concept that a loss in peroxidase activity caused by Prx hyperoxidation would be sufficient to promote redox-based cell signaling is undergoing considerable debate in the literature (Flohé, 2010, Forman, et al., 2010, Stone and Yang, 2006, Winterbourn, 2008), a new study from the Rhee group uses a parallel argument to describe the redox signaling promoting effects of membrane-localized Prx phosphorylation (Woo, et al., 2010). There is currently a large degree of interest in the idea that Prxs act as transducers of the peroxide signal through redox-regulated interactions with effector proteins (e.g., cAbl, cMyc, JNK1), or by oxidizing Trx proteins which can regulate signaling pathways (e.g., through Ask1 regulation) (Flohé, 2010, Forman, et al., 2010, Winterbourn, 2008). Both the floodgate model and the intermediary sensor role for Prxs are strongly supported by investigations into their roles in yeast, while these models in mammals remain much more speculative at this point (Hall, et al., 2009).

SUMMARY

Peroxiredoxins are abundant cellular antioxidant proteins that help to control intracellular peroxide and peroxynitrite levels. These proteins may also function in regulating hydrogen peroxide signaling in eukaryotes through an evolved sensitivity of some peroxiredoxins towards peroxide-mediated inactivation. The conserved active-site environment activates both the peroxidatic cysteine and the hydroperoxide substrate, supporting a high catalytic efficiency. Oligomeric state may also change with redox state, and can play a role in promoting catalysis.

ACKNOWLEDGEMENT

Peroxiredoxin research in the Poole and Karplus laboratories has been supported by funding from the National Institutes of Health (R01 GM050389). Bioinformatics research was also supported, in part, by a grant to Jacquelyn S. Fetrow from the National Science Foundation (MCB 0517343).

Footnotes

1Herein we will use the Roman numerals as specific mammalian Prx names, as they have been used before, and Arabic for the other species and the larger subclasses, but there is no clear concensus across the field on this issue.

INTERNET RESOURCES

http://www.csb.wfu.edu/prex/: PREX is a searchable database containing > 6,000 Prx protein sequences unambiguously classified into one of six distinct subclasses. Subfamily classifications use information around the active sites of structurally characterized subfamily members to search for sequences with conserved functionally-relevant motifs (Nelson, et al., 2011, Soito, et al., 2011).

LITERATURE CITED

  • Adimora NJ, Jones DP, Kemp ML. A model of redox kinetics implicates the thiol proteome in cellular hydrogen peroxide responses. Antioxid Redox Signal. 2010;13:731–743. [PMC free article] [PubMed]
  • Antelmann H, Helmann JD. Thiol-based Redox Switches and Gene Regulation. Antioxid Redox Signal. 2010 [PMC free article] [PubMed]
  • Aran M, Ferrero DS, Pagano E, Wolosiuk RA. Typical 2-Cys peroxiredoxins--modulation by covalent transformations and noncovalent interactions. Febs J. 2009;276:2478–2493. [PubMed]
  • Baker LM, Poole LB. Catalytic mechanism of thiol peroxidase from Escherichia coli. Sulfenic acid formation and overoxidation of essential CYS61. J Biol Chem. 2003;278:9203–9211. [PMC free article] [PubMed]
  • Biteau B, Labarre J, Toledano MB. ATP-dependent reduction of cysteine-sulphinic acid by S. cerevisiae sulphiredoxin. Nature. 2003;425:980–984. [PubMed]
  • Cha MK, Kim HK, Kim IH. Thioredoxin-linked “thiol peroxidase” from periplasmic space of Escherichia coli. J Biol Chem. 1995;270:28635–28641. [PubMed]
  • Chae HZ, Chung SJ, Rhee SG. Thioredoxin-dependent peroxide reductase from yeast. J Biol Chem. 1994;269:27670–27678. [PubMed]
  • Chae HZ, Robison K, Poole LB, Church G, Storz G, Rhee SG. Cloning and sequencing of thiol-specific antioxidant from mammalian brain: alkyl hydroperoxide reductase and thiol-specific antioxidant define a large family of antioxidant enzymes. Proc Natl Acad Sci USA. 1994;91:7017–7021. [PMC free article] [PubMed]
  • Chang TS, Jeong W, Choi SY, Yu S, Kang SW, Rhee SG. Regulation of peroxiredoxin I activity by Cdc2-mediated phosphorylation. J Biol Chem. 2002;277:25370–25376. [PubMed]
  • Choi HJ, Kang SW, Yang CH, Rhee SG, Ryu SE. Crystal structure of a novel human peroxidase enzyme at 2.0 Å resolution. Nat Struct Biol. 1998;5:400–406. [PubMed]
  • Copley SD, Novak WR, Babbitt PC. Divergence of function in the thioredoxin fold suprafamily: evidence for evolution of peroxiredoxins from a thioredoxin-like ancestor. Biochemistry. 2004;43:13981–13995. [PubMed]
  • Cordray P, Doyle K, Edes K, Moos PJ, Fitzpatrick FA. Oxidation of 2-Cys-peroxiredoxins by arachidonic acid peroxide metabolites of lipoxygenases and cyclooxygenase-2. J Biol Chem. 2007;282:32623–32629. [PubMed]
  • Cox AG, Peskin AV, Paton LN, Winterbourn CC, Hampton MB. Redox potential and peroxide reactivity of human peroxiredoxin 3. Biochemistry. 2009;48:6495–6501. [PubMed]
  • Crane EJ, 3rd, Vervoort J, Claiborne A. 13C NMR analysis of the cysteine-sulfenic acid redox center of enterococcal NADH peroxidase. Biochemistry. 1997;36:8611–8618. [PubMed]
  • D’Autreaux B, Toledano MB. ROS as signalling molecules: mechanisms that generate specificity in ROS homeostasis. Nat Rev Mol Cell Biol. 2007;8:813–824. [PubMed]
  • Dietz KJ. Peroxiredoxins in Plants and Cyanobacteria. Antioxid Redox Signal. 2011 in press. [PMC free article] [PubMed]
  • Dubuisson M, Vander Stricht D, Clippe A, Etienne F, Nauser T, Kissner R, Koppenol WH, Rees JF, Knoops B. Human peroxiredoxin 5 is a peroxynitrite reductase. FEBS Lett. 2004;571:161–165. [PubMed]
  • Ellis HR, Poole LB. Roles for the two cysteine residues of AhpC in catalysis of peroxide reduction by alkyl hydroperoxide reductase from Salmonella typhimurium. Biochemistry. 1997;36:13349–13356. [PubMed]
  • Flohé L. Changing paradigms in thiology from antioxidant defense toward redox regulation. Methods Enzymol. 2010;473:1–39. [PubMed]
  • Flohé L, Toppo S, Cozza G, Ursini F. A Comparison of Thiol Peroxidase Mechanisms. Antioxid Redox Signal. 2010 in press. [PubMed]
  • Fomenko DE, Gladyshev VN. Identity and functions of CxxC-derived motifs. Biochemistry. 2003;42:11214–11225. [PubMed]
  • Fomenko DE, Marino SM, Gladyshev VN. Functional diversity of cysteine residues in proteins and unique features of catalytic redox-active cysteines in thiol oxidoreductases. Molecules and cells. 2008;26:228–235. [PMC free article] [PubMed]
  • Fomenko DE, Koc A, Agisheva N, Jacobsen M, Kaya A, Malinouski M, Rutherford J, Siu K-L, Jin D-Y, Winge D, Gladyshev VN. Thiol Peroxidases Mediate Specific Genome-wide Regulation of Gene Expression in Response to Hydrogen Peroxide. Proc Natl Acad Sci U S A. 2011 in press. [PMC free article] [PubMed]
  • Forman HJ, Maiorino M, Ursini F. Signaling functions of reactive oxygen species. Biochemistry. 2010;49:835–842. [PubMed]
  • Fridman JS, Lowe SW. Control of apoptosis by p53. Oncogene. 2003;22:9030–9040. [PubMed]
  • Hall A, Karplus PA, Poole LB. Typical 2-Cys peroxiredoxins - structures, mechanisms and functions. Febs J. 2009;276:2469–2477. [PMC free article] [PubMed]
  • Hall A, Parsonage D, Poole LB, Karplus PA. Structural Evidence that Peroxiredoxin Catalytic Power Is Based on Transition-State Stabilization. J Mol Biol. 2010;402:194–209. [PMC free article] [PubMed]
  • Hall A, Nelson K, Poole L, Karplus PA. Structure-based insights into the catalytic power and conformational dexterity of peroxiredoxins. Antioxid Redox Signal. 2011 in press. [PMC free article] [PubMed]
  • Hirotsu S, Abe Y, Okada K, Nagahara N, Hori H, Nishino T, Hakoshima T. Crystal structure of a multifunctional 2-Cys peroxiredoxin heme-binding protein 23 kDa/proliferation-associated gene product. Proc Natl Acad Sci U S A. 1999;96:12333–12338. [PMC free article] [PubMed]
  • Horta BB, de Oliveira MA, Discola KF, Cussiol JR, Netto LE. Structural and biochemical characterization of peroxiredoxin Qbeta from Xylella fastidiosa: catalytic mechanism and high reactivity. J Biol Chem. 2010;285:16051–16065. [PMC free article] [PubMed]
  • Hugo M, Turell L, Manta B, Botti H, Monteiro G, Netto LE, Alvarez B, Radi R, Trujillo M. Thiol and sulfenic acid oxidation of AhpE, the one-cysteine peroxiredoxin from Mycobacterium tuberculosis: kinetics, acidity constants, and conformational dynamics. Biochemistry. 2009;48:9416–9426. [PubMed]
  • Imlay JA. Cellular defenses against superoxide and hydrogen peroxide. Annu Rev Biochem. 2008;77:755–776. [PMC free article] [PubMed]
  • Iwao-Koizumi K, Matoba R, Ueno N, Kim SJ, Ando A, Miyoshi Y, Maeda E, Noguchi S, Kato K. Prediction of docetaxel response in human breast cancer by gene expression profiling. J Clin Oncol. 2005;23:422–431. [PubMed]
  • Jang HH, Lee KO, Chi YH, Jung BG, Park SK, Park JH, Lee JR, Lee SS, Moon JC, Yun JW, Choi YO, Kim WY, Kang JS, Cheong GW, Yun DJ, Rhee SG, Cho MJ, Lee SY. Two enzymes in one; two yeast peroxiredoxins display oxidative stress-dependent switching from a peroxidase to a molecular chaperone function. Cell. 2004;117:625–635. [PubMed]
  • Jara M, Vivancos AP, Hidalgo E. C-terminal truncation of the peroxiredoxin Tpx1 decreases its sensitivity for hydrogen peroxide without compromising its role in signal transduction. Genes Cells. 2008;13:171–179. [PubMed]
  • Jeong W, Cha MK, Kim IH. Thioredoxin-dependent hydroperoxide peroxidase activity of bacterioferritin comigratory protein (BCP) as a new member of the thiol-specific antioxidant protein (TSA)/Alkyl hydroperoxide peroxidase C (AhpC) family. J Biol Chem. 2000;275:2924–2930. [PubMed]
  • Karplus PA, Hall A. Structural Survey of the Peroxiredoxins. In: Flohé L, Harris JR, editors. Peroxiredoxin Systems. Springer; New York: 2007. pp. 41–60.
  • Kim SK, Yang JW, Kim MR, Roh SH, Kim HG, Lee KY, Jeong HG, Kang KW. Increased expression of Nrf2/ARE-dependent anti-oxidant proteins in tamoxifen-resistant breast cancer cells. Free Radic Biol Med. 2008;45:537–546. [PubMed]
  • Klomsiri C, Karplus PA, Poole LB. Cysteine-Based Redox Switches in Enzymes. Antioxid Redox Signal. 2010 in press. [PMC free article] [PubMed]
  • Knoops B, Clippe A, Bogard C, Arsalane K, Wattiez R, Hermans C, Duconseille E, Falmagne P, Bernard A. Cloning and characterization of AOEB166, a novel mammalian antioxidant enzyme of the peroxiredoxin family. J Biol Chem. 1999;274:30451–30458. [PubMed]
  • Knoops B, Loumaye E, Van der Eecken V. Evolution of the peroxiredoxins: Taxonomy, homology and characterization. In: Flohé L, Harris JR, editors. Peroxiredoxin Systems. Springer; New York: 2007. pp. 27–40.
  • Koo KH, Lee S, Jeong SY, Kim ET, Kim HJ, Song K, Chae H-Z. Regulation of thioredoxin peroxidase activity by C-terminal truncation. Arch. Biochem. Biophys. 2002;397:312–318. [PubMed]
  • Koua D, Cerutti L, Falquet L, Sigrist CJ, Theiler G, Hulo N, Dunand C. PeroxiBase: a database with new tools for peroxidase family classification. Nucleic Acids Res. 2009;37:D261–266. [PMC free article] [PubMed]
  • Lee YM, Park SH, Shin DI, Hwang JY, Park B, Park YJ, Lee TH, Chae HZ, Jin BK, Oh TH, Oh YJ. Oxidative modification of peroxiredoxin is associated with drug-induced apoptotic signaling in experimental models of Parkinson disease. J Biol Chem. 2008;283:9986–9998. [PubMed]
  • Lee YS, Chang HW, Jeong JE, Lee SW, Kim SY. Proteomic analysis of two head and neck cancer cell lines presenting different radiation sensitivity. Acta otolaryngologica. 2008;128:86–92. [PubMed]
  • Lehtonen ST, Svensk AM, Soini Y, Paakko P, Hirvikoski P, Kang SW, Saily M, Kinnula VL. Peroxiredoxins, a novel protein family in lung cancer. Int J Cancer. 2004;111:514–521. [PubMed]
  • Li S, Peterson NA, Kim MY, Kim CY, Hung LW, Yu M, Lekin T, Segelke BW, Lott JS, Baker EN. Crystal Structure of AhpE from Mycobacterium tuberculosis, a 1-Cys Peroxiredoxin. J Mol Biol. 2005;346:1035–1046. [PubMed]
  • Lowther WT, Haynes AC. Reduction of Cysteine Sulfinic Acid in Eukaryotic, Typical 2-Cys Peroxiredoxins by Sulfiredoxin. Antioxid Redox Signal. 2011 in press. [PMC free article] [PubMed]
  • Manta B, Hugo M, Ortiz C, Ferrer-Sueta G, Trujillo M, Denicola A. The peroxidase and peroxynitrite reductase activity of human erythrocyte peroxiredoxin 2. Arch Biochem Biophys. 2009;484:146–154. [PubMed]
  • Moon JC, Hah YS, Kim WY, Jung BG, Jang HH, Lee JR, Kim SY, Lee YM, Jeon MG, Kim CW, Cho MJ, Lee SY. Oxidative stress-dependent structural and functional switching of a human 2-Cys peroxiredoxin isotype II that enhances HeLa cell resistance to H2O2-induced cell death. J Biol Chem. 2005;280:28775–28784. [PubMed]
  • Nelson KJ, Parsonage D, Hall A, Karplus PA, Poole LB. Cysteine pKa values for the bacterial peroxiredoxin AhpC. Biochemistry. 2008;47:12860–12868. [PMC free article] [PubMed]
  • Nelson KJ, Knutson ST, Soito L, Klomsiri C, Poole LB, Fetrow JS. Analysis of the peroxiredoxin family: Using active-site structure and sequence information for global classification and residue analysis. Proteins. 2011 in press. [PMC free article] [PubMed]
  • Neumann CA, Krause DS, Carman CV, Das S, Dubey DP, Abraham JL, Bronson RT, Fujiwara Y, Orkin SH, Van Etten RA. Essential role for the peroxiredoxin Prdx1 in erythrocyte antioxidant defence and tumour suppression. Nature. 2003;424:561–565. [PubMed]
  • Neumann CA, Fang Q. Are peroxiredoxins tumor suppressors? Current opinion in pharmacology. 2007;7:375–380. [PubMed]
  • Ogusucu R, Rettori D, Munhoz DC, Soares Netto LE, Augusto O. Reactions of yeast thioredoxin peroxidases I and II with hydrogen peroxide and peroxynitrite: Rate constants by competitive kinetics. Free Radic Biol Med. 2007;42:326–334. [PubMed]
  • Pak JH, Choi WH, Lee HM, Joo WD, Kim JH, Kim YT, Kim YM, Nam JH. Peroxiredoxin 6 overexpression attenuates cisplatin-induced apoptosis in human ovarian cancer cells. Cancer investigation. 2011;29:21–28. [PubMed]
  • Parmigiani RB, Xu WS, Venta-Perez G, Erdjument-Bromage H, Yaneva M, Tempst P, Marks PA. HDAC6 is a specific deacetylase of peroxiredoxins and is involved in redox regulation. Proc Natl Acad Sci U S A. 2008;105:9633–9638. [PMC free article] [PubMed]
  • Parsonage D, Youngblood DS, Sarma GN, Wood ZA, Karplus PA, Poole LB. Analysis of the link between enzymatic activity and oligomeric state in AhpC, a bacterial peroxiredoxin. Biochemistry. 2005;44:10583–10592. [PMC free article] [PubMed]
  • Parsonage D, Desrosiers DC, Hazlett KR, Sun Y, Nelson KJ, Cox DL, Radolf JD, Poole LB. Broad specificity AhpC-like peroxiredoxin and its thioredoxin reductant in the sparse antioxidant defense system of Treponema pallidum. Proc Natl Acad Sci U S A. 2010;107:6240–6245. [PMC free article] [PubMed]
  • Parsonage D, Reeves SA, Karplus PA, Poole LB. Engineering of fluorescent reporters into redox domains to monitor electron transfers. Methods Enzymol. 2010;474:1–21. [PubMed]
  • Pascual MB, Mata-Cabana A, Florencio FJ, Lindahl M, Cejudo FJ. Overoxidation of 2-Cys peroxiredoxin in prokaryotes: cyanobacterial 2-Cys peroxiredoxins sensitive to oxidative stress. J Biol Chem. 2010;285:34485–34492. [PMC free article] [PubMed]
  • Peskin AV, Low FM, Paton LN, Maghzal GJ, Hampton MB, Winterbourn CC. The high reactivity of peroxiredoxin 2 with H2O2 is not reflected in its reaction with other oxidants and thiol reagents. J Biol Chem. 2007;282:11885–11892. [PubMed]
  • Phalen TJ, Weirather K, Deming PB, Anathy V, Howe AK, van der Vliet A, Jönsson TJ, Poole LB, Heintz NH. Oxidation state governs structural transitions in peroxiredoxin II that correlate with cell cycle arrest and recovery. J Cell Biol. 2006;175:779–789. [PMC free article] [PubMed]
  • Poole LB, Claiborne A. The non-flavin redox center of the streptococcal NADH peroxidase. II. Evidence for a stabilized cysteine-sulfenic acid. J Biol Chem. 1989;264:12330–12338. [PubMed]
  • Poole LB, Reynolds CM, Wood ZA, Karplus PA, Ellis HR, Li Calzi M. AhpF and other NADH:peroxiredoxin oxidoreductases, homologues of low Mr thioredoxin reductase. Eur J Biochem. 2000;267:6126–6133. [PubMed]
  • Poole LB, Karplus PA, Claiborne A. Protein sulfenic acids in redox signaling. Annu. Rev. Pharmacol. Toxicol. 2004;44:325–347. [PubMed]
  • Poole LB. The Catalytic Mechanism of Peroxiredoxins. In: Flohé L, Harris JR, editors. Peroxiredoxin Systems. Springer; New York: 2007. pp. 61–81.
  • Poole LB, Nelson KJ. Discovering mechanisms of signaling-mediated cysteine oxidation. Curr Opin Chem Biol. 2008;12:18–24. [PMC free article] [PubMed]
  • Quan C, Cha EJ, Lee HL, Han KH, Lee KM, Kim WJ. Enhanced expression of peroxiredoxin I and VI correlates with development, recurrence and progression of human bladder cancer. The Journal of urology. 2006;175:1512–1516. [PubMed]
  • Rabilloud T, Heller M, Gasnier F, Luche S, Rey C, Aebersold R, Benahmed M, Louisot P, Lunardi J. Proteomics analysis of cellular response to oxidative stress. Evidence for in vivo overoxidation of peroxiredoxins at their active site. J Biol Chem. 2002;277:19396–19401. [PubMed]
  • Rhee SG, Yang KS, Kang SW, Woo HA, Chang TS. Controlled elimination of intracellular H(2)O(2): regulation of peroxiredoxin, catalase, and glutathione peroxidase via post-translational modification. Antioxid Redox Signal. 2005;7:619–626. [PubMed]
  • Roumes H, Pires-Alves A, Gonthier-Maurin L, Dargelos E, Cottin P. Investigation of peroxiredoxin IV as a calpain-regulated pathway in cancer. Anticancer Res. 2010;30:5085–5089. [PubMed]
  • Sarma GN, Nickel C, Rahlfs S, Fischer M, Becker K, Karplus PA. Crystal structure of a novel Plasmodium falciparum 1-Cys peroxiredoxin. J Mol Biol. 2005;346:1021–1034. [PubMed]
  • Sayed AA, Williams DL. Biochemical characterization of 2-Cys peroxiredoxins from Schistosoma mansoni. J Biol Chem. 2004;279:26159–26166. [PubMed]
  • Seo JH, Koo KH, Kim IG, Chae HZ. Ionizing radiation induced C-terminal truncation of PrxII: A noble peroxidase activity enhancing mechanism. Free Radic Biol Med. 2004;37(Supp. 1):S15.
  • Seo JH, Lim JC, Lee DY, Kim KS, Piszczek G, Nam HW, Kim YS, Ahn T, Yun CH, Kim K, Chock PB, Chae HZ. Novel protective mechanism against irreversible hyperoxidation of peroxiredoxin: Nalpha-terminal acetylation of human peroxiredoxin II. J Biol Chem. 2009;284:13455–13465. [PMC free article] [PubMed]
  • Seo MS, Kang SW, Kim K, Baines IC, Lee TH, Rhee SG. Identification of a new type of mammalian peroxiredoxin that forms an intramolecular disulfide as a reaction intermediate. J Biol Chem. 2000;275:20346–20354. [PubMed]
  • Smith-Pearson PS, Kooshki M, Spitz DR, Poole LB, Zhao W, Robbins ME. Decreasing peroxiredoxin II expression decreases glutathione, alters cell cycle distribution, and sensitizes glioma cells to ionizing radiation and H(2)O(2) Free Radic Biol Med. 2008;45:1178–1189. [PMC free article] [PubMed]
  • Soito L, Williamson C, Knutson ST, Fetrow JS, Poole LB, Nelson KJ. PREX: PeroxiRedoxin classification indEX, a database of subfamily assignments across the diverse peroxiredoxin family. Nucleic Acids Res. 2011;39:D332–337. [PMC free article] [PubMed]
  • Stacey MM, Peskin AV, Vissers MC, Winterbourn CC. Chloramines and hypochlorous acid oxidize erythrocyte peroxiredoxin 2. Free Radic Biol Med. 2009;47:1468–1476. [PubMed]
  • Stone JR, Yang S. Hydrogen peroxide: a signaling messenger. Antioxid Redox Signal. 2006;8:243–270. [PubMed]
  • Su D, Berndt C, Fomenko DE, Holmgren A, Gladyshev VN. A conserved cis-proline precludes metal binding by the active site thiolates in members of the thioredoxin family of proteins. Biochemistry. 2007;46:6903–6910. [PubMed]
  • Szatrowski TP, Nathan CF. Production of large amounts of hydrogen peroxide by human tumor cells. Cancer Res. 1991;51:794–798. [PubMed]
  • Trujillo M, Mauri P, Benazzi L, Comini M, De Palma A, Flohe L, Radi R, Stehr M, Singh M, Ursini F, Jaeger T. The mycobacterial thioredoxin peroxidase can act as a one-cysteine peroxiredoxin. J Biol Chem. 2006;281:20555–20566. [PubMed]
  • Trujillo M, Clippe A, Manta B, Ferrer-Sueta G, Smeets A, Declercq JP, Knoops B, Radi R. Pre-steady state kinetic characterization of human peroxiredoxin 5: taking advantage of Trp84 fluorescence increase upon oxidation. Arch Biochem Biophys. 2007;467:95–106. [PubMed]
  • Veal EA, Day AM, Morgan BA. Hydrogen peroxide sensing and signaling. Mol Cell. 2007;26:1–14. [PubMed]
  • Wang G, Olczak AA, Walton JP, Maier RJ. Contribution of the Helicobacter pylori thiol peroxidase bacterioferritin comigratory protein to oxidative stress resistance and host colonization. Infect Immun. 2005;73:378–384. [PMC free article] [PubMed]
  • Wang T, Tamae D, LeBon T, Shively JE, Yen Y, Li JJ. The role of peroxiredoxin II in radiation-resistant MCF-7 breast cancer cells. Cancer Res. 2005;65:10338–10346. [PubMed]
  • Winterbourn CC. Reconciling the chemistry and biology of reactive oxygen species. Nat Chem Biol. 2008;4:278–286. [PubMed]
  • Woo HA, Yim SH, Shin DH, Kang D, Yu DY, Rhee SG. Inactivation of Peroxiredoxin I by Phosphorylation Allows Localized H2O2 Accumulation for Cell Signaling. Cell. 2010;140:517–528. [PubMed]
  • Wood ZA, Poole LB, Hantgan RR, Karplus PA. Dimers to doughnuts: redox-sensitive oligomerization of 2-cysteine peroxiredoxins. Biochemistry. 2002;41:5493–5504. [PubMed]
  • Wood ZA, Poole LB, Karplus PA. Peroxiredoxin evolution and the regulation of hydrogen peroxide signaling. Science. 2003;300:650–653. [PubMed]
  • Wood ZA, Schröder E, Harris JR, Poole LB. Structure, mechanism and regulation of peroxiredoxins. Trends Biochem Sci. 2003;28:32–40. [PubMed]
  • Yamamoto Y, Ritz D, Planson AG, Jonsson TJ, Faulkner MJ, Boyd D, Beckwith J, Poole LB. Mutant AhpC peroxiredoxins suppress thiol-disulfide redox deficiencies and acquire deglutathionylating activity. Mol Cell. 2008;29:36–45. [PMC free article] [PubMed]
  • Yang KS, Kang SW, Woo HA, Hwang SC, Chae HZ, Kim K, Rhee SG. Inactivation of human peroxiredoxin I during catalysis as the result of the oxidation of the catalytic site cysteine to cysteine-sulfinic acid. J Biol Chem. 2002;277:38029–38036. [PubMed]
  • Yeh JI, Claiborne A, Hol WG. Structure of the native cysteine-sulfenic acid redox center of enterococcal NADH peroxidase refined at 2.8 Å resolution. Biochemistry. 1996;35:9951–9957. [PubMed]
  • Yuan Y, Knaggs MH, Poole LB, Fetrow JS, Salsbury FR. Conformational and oligomeric effects on the cysteine pK(a) of tryparedoxin peroxidase. Journal of biomolecular structure & dynamics. 2010;28:51–70. [PMC free article] [PubMed]
  • Zhang B, Wang Y, Su Y. Peroxiredoxins, a novel target in cancer radiotherapy. Cancer Lett. 2009;286:154–160. [PubMed]
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