Encyclopedia of Signaling Molecules

2018 Edition
| Editors: Sangdun Choi


Reference work entry
DOI: https://doi.org/10.1007/978-3-319-67199-4_101839


 AhpC;  Peroxiredoxin;  Prx;  Thioredoxin peroxidase (antioxidant enzyme);  Thioredoxin peroxidase;  Thioredoxin-dependent peroxide reductase;  Tpx;  Tsa

From NCBI, Homo sapiens:

PRDX1 = Prx1, PrxI, HBP23, MSP23, OSF-3, TPx-A, NKEF-A, PAG, TDPX2 (location: Chromosome 1, NC_000001.11: 45511035..45522890, complement)

PRDX3 = Prx3, PrxIII, AOP-1, HBC189, MER5, PRO1748, SP-22 (Chromosome 10, NC_000010.11: 119167699..119178865, complement)

PRDX4 = Prx4, PrxIV, AOE372, TRANK, HEL-S-97n (Chromosome X, NC_000023.11: 23664260..23686399)

PRDX5 = Prx5, PrxV, ACR1, AOEB166, B166, HEL-S-55, PLP, PMP20, AOPP, SBBI10 (Chromosome 11, NC_000011.10: 64318088..64321823)

PRDX6 = Prx6, PrxVI, hORF6, 1-Cys, AOP2, HEL-S-128m, NSGPx, aiPLA2, LTW4, p29 (Chromosome 1, NC_000001.11: 173477347..173488807)

Historical Background

The first reports of the family of thiol peroxidases eventually known as peroxiredoxins (PRDXs or Prxs) came in the late 1960s, when a protein named “torin,” isolated from human erythrocytes, was shown by transmission electron microscopy to form discrete complexes with tenfold symmetry (Harris 1969). This protein is now recognized as PrxII, the third most abundant protein in erythrocytes. In the late 1980s, a “protector protein” from Saccharomyces cerevisiae was shown to preserve glutamine synthetase activity in a metal and thiol-containing oxidizing system by Sue Goo Rhee and colleagues working with Earl Stadtman at the National Institutes of Health (Kim et al. 1988). This “thiol-specific antioxidant” protein was subsequently discovered to be evolutionarily related to a bacterial peroxidase discovered around the same time (Chae et al. 1994), the alkyl hydroperoxide reductase “C-22 component” (now known as AhpC) identified in Bruce Ames’ group as part of a two-component, OxyR-inducible peroxidase system capable of scavenging organic hydroperoxides (Jacobson et al. 1989). By the mid-1990s, these enzymes were discovered to be widely distributed and very abundant (Chae et al. 1994), although it took most of another decade for them to be recognized as important oxidant defense enzymes alongside the better-known catalase and glutathione peroxidase enzymes (Seaver and Imlay 2001). With the increasing recognition that these proteins serve important roles in both oxidant defense and regulation of signal transduction, peroxiredoxin research has continued to intensify.

Peroxiredoxin Catalysis and Structure

Peroxiredoxins (EC rely on a single, conserved cysteine at the active site, often referred to as the peroxidatic cysteine (CP) which in its deprotonated (thiolate) form performs a nucleophilic attack on the O–O bond of a hydroperoxide substrate, generating a hydroxyl-containing product as well as an oxidized form of the Prx (the sulfenic acid form of CP, shown as SpOH in Fig. 1). This step of peroxidation (Fig. 1, step 1) is common to all Prxs and proceeds, in many Prxs studied, with a second order rate constant as high as 106 to 108 M-1 s-1 for the preferred substrate of each. Degree of specificity can vary, but Prx substrates can include hydrogen peroxide, lipid hydroperoxides, protein and amino acid hydroperoxides, peroxynitrite, and model peroxides like cumene hydroperoxide and t-butyl hydroperoxide. For the catalytic cycle to proceed, the sulfenic acid must leave the active-site environment through an unfolding event to yield a locally unfolded (LU) conformation that allows disulfide bond formation between CP and another thiol group, termed the resolving thiol (Fig. 1, step 2, shown as SrH). In many Prxs, the resolving thiol comes from another cysteine in the protein, termed the resolving cysteine (CR), forming either an inter- or intrasubunit disulfide bond. The disulfide-bonded Prx must then be reduced, typically by thioredoxin or by another thioredoxin-like reductase protein or domain (Fig. 1, step 3), and the enzyme returned to its fully folded (FF) conformation, in order to restore its activated state. Prxs in their sulfenic acid form (just after step 1) can also be sensitive toward being inactivated by their own peroxide substrates (Fig. 1, step 4), generating a regulatory modification, the sulfinic acid (SpO2H in Fig. 1), that inactivates them. A repair protein capable of the ATP-dependent reversal of this oxidation (Fig. 1, step 5) was first identified in yeast and named sulfiredoxin by the group of Michel Toledano (Biteau et al. 2003). The existence of this repair protein in many eukaryotes highlights the biological significance of this oxidative modification, often referred to as hyperoxidation.
Peroxiredoxins, Fig. 1

Prx catalytic cycle. In the first step (1) of the Prx catalytic cycle, conserved across all Prxs, the thiolate form of the peroxidatic cysteine (-Sp- in green) reacts with peroxide to form cysteine sulfenic acid (-SpOH in purple). In step 2, the sulfenic acid forms a disulfide (-Sp-Sr- in blue) with a resolving thiol (-SrH) from the same subunit, from another Prx subunit, or from an exogenous molecule. A structural change involving local unfolding of the fully folded (FF) active site loop to a locally unfolded (LU) conformation must occur to expose the sulfenic acid so that the disulfide bond can form in step 2. In step 3, the Prx is recycled back to the reduced form through a disulfide exchange reaction with thioredoxin or a similar protein or domain, and local refolding reestablishes the stubstrate-ready, FF form of the active site. Hyperoxidation (step 4) can in some cases occur to form the catalytically inactive sulfinic acid (-SPO2H in red) but can be reversed by sulfiredoxin (Srx; step 5) (Adapted from Poole et al. (2013), copyright 2013, with permission from Springer)

Structurally, Prxs contain a core thioredoxin fold, with a central β sheet and flanking α helices (Fig. 2a), and a highly conserved active site architecture around the TXXC motif that is conserved and substitutes for the CXXC motif of thioredoxins and glutaredoxins (Fig. 2b). Additional conserved residues include an upstream Pro (in the characteristic PXXXTXXC Prx motif) and an Arg that activates both the CP (lowering its pKa) and the incoming peroxide substrate (Perkins et al. 2015). Stabilization of the transition state as one of the two oxygens from the peroxide is transferred to the sulfur occurs through optimized hydrogen-bonding interactions and stabilization of the developing negative charge (Hall et al. 2010; Ferrer-Sueta et al. 2011). Six classes of Prxs can be distinguished structurally and bioinformatically (Hall et al. 2011; Perkins et al. 2015): E. coli has members from three of the groups, Prx1 (named AhpC in E. coli), PrxQ (named BCP in E. coli), and Tpx (a “thiol peroxidase”). Human Prxs, of which there are six, include four Prx1 members (PrxI through PrxIV), the canonical Prx5 member (human PrxV), and the canonical Prx6 or “1-Cys Prx” member (human PrxVI). The final group, AhpE, is present only in a subset of bacteria, the Actinomycetes (which includes Mycobacterium tuberculosis), and less information is available about this group. Note: we are here using Roman numerals for the specific human Prx designation, and Arabic for the overall group name, but there is variability in the names used in the literature.
Peroxiredoxins, Fig. 2

Core secondary structure and conserved active site of Prx proteins. (a) Shown is the back view of a representative monomer of a fully folded (FF) Prx (Salmonella typhimurium AhpC; PDB entry 1N8J) with the conserved structural elements labeled and β-sheets shown in black. The CP is located in the first turn of helix α2 and is shown as spheres (readily seen in panel b). An asterisk represents the position of attachment of the rest of the C-terminal tail that has been hidden in this view (normally absent in other groups of Prxs except Prx6); the C-terminus of Prx1 and Prx6 subfamily members extends from here to make extensive interactions with an adjacent monomer. (b) The same monomer viewed from the front side of the cradle (related to A by a ~180° rotation around a vertical axis). (c) This overlay of Prx active sites includes high-resolution structures of (i) DTT-bound human PrxV (light blue; PDB 3MNG, 1.45 Å resolution), (ii) peroxide-bound Aeropyrum pernixTpx (actually a Prx6 group member) (green; PDB 3A2V, 1.65 Å resolution), and (iii) water-bound wild-type Xanthomonas campestris PrxQ (white; PDB 5IIZ, 1.05 Å resolution). Heteroatoms are colored red (oxygens), blue (nitrogens), and yellow (sulfurs), and ligands are colored cyan (DTT), lime (H2O2), and dark gray (waters) (Panels a and b are reprinted with permission from Springer (Poole et al. (2013)). Panel c is reprinted with permission from Elsevier from Perkins et al. (2016)

With the identification of this as a widespread family in the mid-1990s (Chae et al. 1994), the designation of “1-Cys” and “2-Cys” Prxs was introduced as it was noticed that, among the 22 Prx sequences known at the time, only one Cys residue, fairly near the N-terminus (around residue 46–52 out of 156–199 residues total) was absolutely conserved; this is the residue now recognized as CP (Fig. 1). The second cysteine recognized at that time as semiconserved (hence distinguishing these as the “2-Cys Prxs”) is the resolving cysteine, CR, which forms an intersubunit disulfide bond with CP in the widespread and abundant Prx1 group, sometimes referred to as the “typical 2-Cys Prxs.” With accumulating data, it became clear that the CR resides in multiple positions in the Prx family members other than just in the C-terminal region of a partner subunit (Fig. 3) and the designation of “atypical 2-Cys Prx” arose (Prxs for which a CR is present but not in the originally identified position characteristic of the Prx1 group). As our knowledge of this protein family has grown, these other designations are not particularly meaningful and are better replaced with the group name to which a given Prx belongs. [To assist with identification of new members and the subgroup to which they belong, a searchable database with GenBank sequences through October 2011 is publicly available at “http://csb.wfu.edu/PREX” (Soito et al. 2011)]. Also, the “1-Cys Prx” designation was originally associated with the human PrxVI, but it is now recognized that the existence and location of CR across all six groups is heterogeneous, with “1-Cys” members present in all of the subgroups and five known locations for CR (distributed unevenly among the Prx groups; Fig. 3).
Peroxiredoxins, Fig. 3

Variable locations of the resolving cysteine (CR). (a) Parts of three subunits of a decameric Prx are shown as ribbons (gray, white, and gold). Also shown are the various positions of the peroxiredoxin CR (colored sidechains) in relation to the active site peroxidatic cysteine (CP, circled and in red). Intrasubunit CP-CR disulfides are formed when CR is located in α2 (yellow), α3 (green), and α5 (blue), and intersubunit disulfides are formed when CR is located near the N-terminus (Nt orange, CR in the gold chain) or C-terminus (Ct magenta, CR in the black chain). (CR residues are mapped onto a composite structure based on S. typhimurium AhpC, Protein Databank Identifier 4MA9). (b) Pie charts illustrating the prevalence of each resolving Cys (CR) location among all six subfamilies of peroxiredoxins (Prxs). Wedges are colored by CR position consistent with the structure shown in panel a. No CR (red) indicates that there is no other Cys in the protein and uncertain (gray) indicates that there are other Cys residues but not in recognized CR positions. Hybrid proteins in the Prx5 subfamily with a C-terminally appended Grx domain, as exemplified by Haemophilus influenza Prx5–Grx, are represented in cyan (no clear CR) (Reproduced with permission from Elsevier from Perkins et al. (2015))

Oligomeric States of Peroxiredoxins

Almost all members of the six groups of Prxs are dimeric or larger assemblies of dimers; only the PrxQ group is known to include monomeric Prxs. Two types of dimers are formed, however, depending on the Prx group. B-type dimers are formed for members of the Prx1, Prx6, and AhpE groups, bringing β-strands together side-by-side to form an extended β-sheet (Fig. 4, B-type dimers). A different type of interface, called the A-type interface, is used to form dimeric Prx5, Tpx, and PrxQ proteins. Interestingly, B-type dimers can also come together through the A interface to form decamers (or sometimes dodecamers), and in Prx1 group members, this dimer-decamer equilibrium can be influenced by redox state, concentration, and pH (Hall et al. 2011; Perkins et al. 2015).
Peroxiredoxins, Fig. 4

Quaternary structures of Prxs. Dimeric α2 complexes are formed using either an A-type interface, where the monomers interact near helix α3, or the B-type interface, where the interaction is through the β-strands, generating an extended 10–14 strand β-sheet. Further interactions at the A-interfaces of some Prx1 and Prx6 members generate (α2)5 decamers [or in rare cases (α2)6 dodecamers]. The blue subunit is displayed in approximately the same orientation in each of the structures to illustrate these interaction interfaces that together build the decamer. For a number of Prx1 members, the structural change upon disulfide bond formation destabilizes the A-type interface of the decamers, promoting dissociation to B-type dimers. The structures depicted are: Aeropyrum pernix PrxQ (A-type dimer, Protein Data Bank Identifier 4GQF) and wild-type S. typhimurium AhpC (B-type dimer and decamer, Protein Data Bank Identifier 4MA9) (Reproduced with permission from Elsevier from (Perkins et al. 2015))

Functions of Prxs in Oxidant Defense and Cell Signaling

The first functions recognized for Prxs were in catalyzing peroxide reduction and protecting DNA and proteins from oxidative damage (Kim et al. 1988; Jacobson et al. 1989), and subsequent studies of diverse proteins in many organisms confirm the importance of Prxs in oxidant defense. However, considering that H2O2 is an important signaling mediator and regulator in multicellular organisms, these efficient peroxide scavengers must somehow be regulated or harnessed in order to allow or perhaps mediate cell signaling through H2O2, lipid hydroperoxides, and potentially other substrates. The multiple ways that Prxs can be regulated (e.g., cysteine hyperoxidation; threonine, serine, and tyrosine phosphorylation; acetylation on lysine and the N-terminal amine; cysteine S-nitrosation; tyrosine nitration; cysteine glutathiolation; and C-terminal truncation) emphasize their roles beyond defense (Chae et al. 2012; Randall et al. 2014; Peskin et al. 2016) (Fig. 5). Higher order oligomers (beyond decamers) can even exhibit chaperone activity, as demonstrated for hyperoxidized and aggregated forms of Tsa1 and Tsa2 from Saccharomyces cerevisiae (Chae et al. 2012). Loss of Prx activity through hyperoxidation or phosphorylation in a highly localized manner that can then allow for H2O2-mediated oxidation of other targets was proposed as the “floodgate hypothesis” in 2003 (Wood et al. 2003) and subsequently demonstrated in several scenarios, including hyperoxidation of PrxI during cell cycle progression (Lim et al. 2015) and phosphorylation of PrxI during proliferative signaling (Woo et al. 2010). Evidence for PrxII as a sensor and “carrier” of the oxidation signal from H2O2 has also been presented in the case of STAT3 signaling in inflammatory responses, and a number of signaling proteins (e.g., c-Abl, c-Myc, Mst1, Ask1, and PTEN) are known to bind to Prxs, in some cases in a redox-dependent manner (Chae et al. 2012; Sobotta et al. 2015). Elucidation of pathways and mechanisms of cell signaling involving Prxs continues to be a very active area of research.
Peroxiredoxins, Fig. 5

Posttranslational modifications (PTMs) of human PRDX1 and PRDX2 (also known as PrxI or Prx1 and as PrxII or Prx2, respectively). Shown between the bars representing full-length proteins are PTMs common between the two proteins, including lysine acetylation (green ellipse), phosphorylation (orange triangles), S-glutathiolation (purple “SSG”), and cysteine oxidation at either the peroxidatic cysteine (CPox) or the resolving cysteine (CRox); see Fig. 1 for cysteine roles in catalysis. A previously recognized “YF” motif in both, near the C-terminus, is often found in peroxiredoxins sensitive to peroxide-mediated hyperoxidation of CP to sulfinic (CP-SO2H) [or sulfonic (CP-SO3H)] acids (Wood et al. 2003). CP-SO2H can be reversed by the enzyme sulfiredoxin (Srx) for some (Prx1 group) peroxiredoxins. Note that CP sulfenic acid (CP-SOH) and disulfide-bonded CP-CR are formed normally during catalysis (Fig. 1); CR may also be otherwise oxidized or modified under certain conditions. Oxidative modifications can be caused by excess reactive oxygen species (ROS) or reactive nitrogen species (RNS). Modifications shown above PRDX1 or below PRDX2 have only been demonstrated for these specific proteins but this does not rule out their occurrence on the other homolog (except C83 glutathiolation, as this cysteine is not present in PRDX2). Acetylation at the N-terminal amine group occurs after processing to remove the initiator methionine in PRDX2 but not PRDX1 (green ellipse). Sites shown are phosphorylated by the following kinases: TOPK (S32), CDK1 (T89 and T90), Mst1 (T183), and Src family kinases (Y194). Blue zig-zags illustrate the locations of α helices α2, α3, α4, and α5; helix α6, which is not observed in crystal structures of disulfide-bonded PRDX1 and PRDX2, is similarly shown in pink


Peroxiredoxins (PRDXs) are cysteine-based hydroperoxide reductases that provide cellular defenses against these oxidants (including hydrogen peroxide, lipid hydroperoxides, and peroxynitrite) but also play a role in regulating or mediating cell signaling processes. These enzymes fall into six structural subfamilies, and many organisms possess more than one PRDX, usually from more than one subfamily. For example, humans express six PRDXs (from six distinct genes) which represent three of the subfamilies. A variety of posttranslational modifications have been observed to regulate PRDX activities and/or interactions, and a repair enzyme, sulfiredoxin, can resurrect a subset of these proteins following inactivation by hyperoxidation.


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Copyright information

© Springer International Publishing AG 2018

Authors and Affiliations

  1. 1.Department of Biochemistry and Center for Redox Biology and MedicineWake Forest School of MedicineWinston-SalemUSA