Peroxiredoxins in malaria parasites: Parasitologic aspects

Available online at www.sciencedirect.com

Parasitology International 57 (2008) 1 – 7 www.elsevier.com/locate/parint

Review

Peroxiredoxins in malaria parasites: Parasitologic aspects Shin-ichiro Kawazu a,b,c,⁎, Kanako Komaki-Yasuda b , Hiroyuki Oku d , Shigeyuki Kano b a

c

National Research Center for Protozoan Diseases, Obihiro University of Agriculture and Veterinary Medicine, 2-13 Inada-cho, Obihiro, Hokkaido 080-8555, Japan b Research Institute, International Medical Center of Japan, 1-21-1 Toyama, Shinjuku-ku, Tokyo 162-8655, Japan Precursory Research for Embryonic Science and Technology, Japan Science and Technology Agency, Chiyoda-ku, Tokyo 102-0075, Japan d Department of Chemistry and Chemical Biology, Gunma University, Kiryu, Gunma 376-8515, Japan Received 2 July 2007; recieved in revised form 2 August 2007; accepted 4 August 2007 Available online 24 August 2007

Abstract Malaria is one of the most debilitating and life threatening diseases in tropical regions of the world. Over 500 million clinical cases occur, and 2–3 million people die of the disease each year. Because Plasmodium lacks genuine glutathione peroxidase and catalase, the two major antioxidant enzymes in the eukaryotic cell, malaria parasites are likely to utilize members of the peroxiredoxin (Prx) family as the principal enzymes to reduce peroxides, which increase in the parasite cell due to metabolism and parasitism during parasite development. In addition to its function of protecting macromolecules from H2O2, Prx has also been reported to regulate H2O2 as second messenger in transmission of redox signals, which mediate cell proliferation, differentiation, and apoptosis. In the malaria parasite, several lines of experimental data have suggested that the parasite uses Prxs as multifunctional molecules to adapt themselves to asexual and sexual development. In this review, we summarize the accumulated knowledge on the Prx family with respect to their functions in mammalian cells and their possible function(s) in malaria parasites. © 2007 Elsevier Ireland Ltd. All rights reserved. Keywords: Antioxidant; Malaria; Peroxiredoxin; Plasmodium; Thioredoxin peroxidase

Contents 1. 2.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . Peroxiredoxins . . . . . . . . . . . . . . . . . . . . . . . 2.1. Overview . . . . . . . . . . . . . . . . . . . . . . 2.2. Prx as regulator of intracellular signal transduction . 2.3. Targeted inactivation of Prx in mice . . . . . . . . 3. Prx and thioredoxin systems in malaria parasites . . . . . 3.1. Overview . . . . . . . . . . . . . . . . . . . . . . 3.2. 2-Cys Prx (TPx-1 and TPx-2) . . . . . . . . . . . . 3.3. 1-Cys Prx (1-Cys-Prx and AOP) . . . . . . . . . . 4. Concluding remarks . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

. . . . . . . . . . . .

2 2 2 2 3 3 3 3 5 6 6 6

Abbreviations: AOP, antioxidant protein; GSH, glutathione; GPx, GSH peroxidase; Grx, glutaredoxin; 8-OHdG, 8-hydroxy-2′-deoxyguanosine; Prx, peroxiredoxin; Plrx, plasmoredoxin; RNS, reactive nitrogen species; ROS, reactive oxygen species; SOD, superoxide dismutase; TSA, thiol-specific antioxidant; Trx, thioredoxin; TPx, Trx peroxidase; TrxR, Trx reductase. ⁎ Corresponding author. National Research Center for Protozoan Diseases, Obihiro University of Agriculture and Veterinary Medicine, 2-13 Inada-cho, Obihiro, Hokkaido 080-8555, Japan. Tel.: +81 155 495846; fax: +81 155 495643. E-mail address: [email protected] (S. Kawazu). 1383-5769/$ - see front matter © 2007 Elsevier Ireland Ltd. All rights reserved. doi:10.1016/j.parint.2007.08.001

2

S. Kawazu et al. / Parasitology International 57 (2008) 1–7

1. Introduction Malaria is a disease caused by infection with protozoan parasites of the genus Plasmodium and transmitted by Anopheles mosquitoes. Almost half of the world's population lives with a serious risk of contacting malaria, and 2–3 million people die of the disease each year [1]. Despite years of intensive research, an effective vaccine is still not available, and the parasite displays increasing resistance towards the commonly used anti-malarial drugs. To combat malaria, a better understanding of the basic biology of the parasite, especially the mechanism of adaptation to environmental conditions is needed. Reactive oxygen species (ROS) are produced in cells during normal aerobic metabolism in oxygen-containing environments [2,3]. To protect macromolecules from the effects of ROS, aerobes have evolved efficient defense systems composed of nonenzymatic and enzymatic antioxidants [3]. The four major cellular antioxidant enzymes are superoxide dismutase (SOD), catalase, glutathione (GSH) peroxidase (GPx), and peroxiredoxin (Prx) [4]. As Plasmodium spp. (malaria parasites) actively proliferate within erythrocytes of their vertebrate hosts, large quantities of ROS, which damage macromolecules, are generated [5,6]. A major source of ROS in parasite cells is heme, a byproduct of the digestion of hemoglobin for nutrition [7,8]. ROS are also generated when the parasite is exposed to various stress conditions, such as those induced by the host immune system [9]. The malaria parasites develop sexually in the digestive tract (midgut) of the vector mosquito, translocate to the salivary grand, and then mature intracellularly in the salivary gland to facilitate the transmission. In these environments, parasites are also likely to be under oxidative stress [10,11]. Because malaria parasites are highly susceptible to such oxidative burden, their antioxidant defenses are considered to play essential roles throughout the lifecycle. Such defenses are thus expected to be potential targets for malaria control strategies [12–14]. 2. Peroxiredoxins 2.1. Overview Peroxiredoxins (Prxs) are a ubiquitous family of antioxidant enzymes that catalyze the reduction of H2O2 to H2O with a thiol as the other substrate. The first member of the family, thiolspecific antioxidant (TSA), was discovered by Chae et al. [15,16] in Saccharomyces as a protein that protects glutamine synthetase from inactivation by a thiol/Fe(III)/oxygen mixedfunction oxidation system. Proteins structurally homologous to TSA have been identified in all living organisms, from bacteria to humans, and these proteins were named the peroxiredoxin (Prx) family [16,17]. Prx proteins have been identified in both helminth and protozoan parasites [18–20]. Prx of some parasites, such as Entamoeba histolytica [21,22], Fasciola hepatica [23], and Schistosoma japonicum [24], are characterized as antigens or secreted proteins, suggesting that these molecules may also participate in the defense against the host

attacks. Mammalian cells express six isoforms of Prx (Prx I to VI) [16,17]. Interestingly, several mammalian Prx proteins were discovered without reference to antioxidant function, suggesting the multi-functional nature of this family of proteins [4,16,17,25]. On the basis of the number and position of cysteine (Cys) residues that participate in catalysis, Prxs are divided into three subgroups, typical 2-Cys Prx, atypical 2-Cys Prx, and 1-Cys Prx [17,25]. Both 1-Cys and 2-Cys Prx contain a conserved cysteine residue at the N terminus that is oxidized to sulfenic acid (Cys-SOH) during the catalytic reaction. This Cys is referred to as the peroxidatic Cys (CysP-SH) and serves as the site of oxidation by H2O2. The amino acid sequences surrounding the CysP-SH in 2-Cys Prx and 1-Cys Prx are conserved, and these sequences are FVCP and PVCT, respectively [17,25]. Prx proteins do not contain redox cofactors, such as heme, flavin, or metal ions that might participate in the catalytic reaction [16,17,25]. Typical 2-Cys Prx forms a homodimer during the catalytic reaction; oxidation of CysP-SH to CysP-SOH is followed by formation of an intermolecular disulfide bond between Cysp and the second conserved Cys at the C terminus of the other subunit, which is referred to as the resolving Cys (CysR-SH). Finally, the 2-Cys disulfides are reduced by another biothiol, such as thioredoxin (Trx), and the dimer dissociates into two regenerated monomers [17,25]. Atypical 2-Cys Prx contains only Cysp but requires one additional, non-conserved Cys residue for the catalytic reaction. An intramolecular disulfide bond is formed that is also reduced by Trx [17,25]. In 1-Cys Prx, Cysp-SOH is reduced directly by a redox partner that has not yet been clearly identified [17,25]. In addition to the peroxidase activity, several 2-Cys Prx enzymes are able to reduce reactive nitrogen species (RNS), such as peroxynitrite (ONOO − ), that are formed by nitric oxide (NO) and superoxide (O2− ). [16]. The enzymatic activity of 2-Cys Prxs, including mammalian Prx II, are thought to be controlled by the structural change from the dimer to decamer, because the decameric enzyme exhibits higher peroxidase activity [16,25]. The redox state of the catalytic Cys of the enzyme is critical for determining dimerization (oxidized) and decamerization (reduced) of the enzyme [16,25]. 2.2. Prx as regulator of intracellular signal transduction Recently, intracellular H2O2 surge, which occurs in response to the activation of cell surface receptors, has attracted interest as a second messenger in receptor-mediated signaling [26,27]. This H2O2 surge has been proposed to modify protein function through oxidation of critical Cys residues of enzymes such as tyrosine phosphatases and kinases [26–28]. It has been suggested that, in addition to defending against oxidative stress, 2-Cys Prx (Prx I and II) may regulate of such an H2O2 surge [27–30]. With respect to the regulatory function of Prx in transduction of cell signals, two different mechanisms that allow temporary inactivation of the enzyme and accumulation of H2O2 at the site of the signaling action, have been proposed: (1) phosphorylation at Thr90 by cyclin-dependent kinase (Cdc2) during M phase [31] and (2)

S. Kawazu et al. / Parasitology International 57 (2008) 1–7

hyperoxidation of the active site CysP to Cys sulfinic acid (CysSO2H). Such hyperoxidation is not reversed by Trx but is reversed by the reaction catalyzed by sulfiredoxin [29,32–34]. These two control mechanisms probably favor different oligomeric states [16,25]. Another mechanism proposed to regulate peroxidase activity of Prx is specific proteolysis of the C terminus of the molecule, which makes the enzyme resistant to overoxidation but leaves it susceptible to inactivation by phosphorylation [16,25].

3

The Trx reductase (TrxR) of P. falciparum is a homodimeric, FAD-dependent oxidoreductase [5,6]. PfTrxR is not selenium dependent and has a C-terminal active site motif that differs from that of human TrxR [5,6]. Analysis of gene knockouts revealed that PfTrxR is essential for asexual development of the parasite [53]. Therefore, it is believed that GSH is the major redox buffer for transient H2O2 exposure and that the basal cellular peroxide flux in the parasite is handled by the Trx system, which includes four Prx subfamilies [5,6,14].

2.3. Targeted inactivation of Prx in mice 3.2. 2-Cys Prx (TPx-1 and TPx-2) The various functions and importance of Prx were indicated by the phenotypes of several knockout experiments in mice. The roles of Prx I in aging and carcinogenesis were suggested by the phenotypes of mice carrying a targeted deletion of the Prx I gene; the knockout mice had shortened lifespan owing to severe hemolytic anemia and increased frequency of several malignant cancers [35]. Prx II null (−/−) mice showed hemolytic anemia with higher intraerythrocytic ROS levels, and the phenotype suggested that Prx II plays a major role in protecting erythrocytes from oxidative stress [36]. Increased cellular senescence was observed in Prx II (−/−) mouse embryonic fibroblasts [37]. Targeted inactivation of Prx VI (1-Cys Prx) gene in mice resulted in increased susceptibility to paraquat-induced lung injury, suggesting importance of Prx in the lung [38]. 3. Prx and thioredoxin systems in malaria parasites 3.1. Overview Despite multiple entries of SOD [39,40], which dismute superoxide into hydrogen peroxide, in PlasmoDB (the Plasmodium genome resource), malaria parasites do not express two of the major peroxidases, catalase and genuine GPx [5,6]. To reduce peroxides, which are produced in each cellular compartment by metabolism and parasitism, the parasites are equipped with five peroxidases localized in the cytoplasm, mitochondrion, and presumably in apicoplast [20,41]. These peroxidases include 1-Cys Prx, two typical 2-Cys Prxs, a 1-Cys antioxidant protein (AOP), and a GSH peroxidase-like thioredoxin peroxidase (TPxGl) [20,41]. The 1-Cys Prx [42,43] and one of the 2-Cys Prxs (TPx-1) [43–45] are expressed in the cytosol, and the other 2-Cys Prx (TPx-2) is expressed in mitochondria [45,46]. The AOP has a signal that presumably target localization to apicoplasts [47], but the true localization of this Prx has not been identified. TPxGl has putative signal at the N terminus, but the localization of this enzyme is also unclear [6]. Genes encoding five thioredoxin (Trx) and Trx-like proteins have been identified in the parasite genome, and the proteins act as the redox-active molecules, which donate electrons through the thioredoxin system to peroxidases [41,48]. In addition to Trx, the parasite expresses a typical glutaredoxin (Grx) and Grxlike proteins [49,50] that are fueled by the GSH system and donate electrons to AOP [41]. In addition, there is a molecule unique to Plasmodium spp., plasmoredoxin (Plrx) that bridges the Trx and GSH systems [51] and can donate electrons to TPx-1 [52].

During the trophozoite stage, P. falciparum TPx-1 (PfTPx-1), which is one of the most abundantly expressed subfamilies in the parasite cytoplasm, accounts for 0.25% of the total cellular protein [44]. The kinetic parameters determined with the recombinant enzyme indicated high specificity for H2O2 (apparent Km = 0.25 μM) [52,54], suggesting that it contributes to peroxide reduction in the parasite cell. It has recently been reported that PfTPx-1 can also reduce peroxynitrite as its substrate [52]. Like 2-Cys Prx in other organisms, PfTPx-1 can form decamers consisting of five homodimers and having a doughnut-like shape [54]. As the structure-determining mechanism, redox state (reduction in Cysp: Cys50) and abundance (higher concentration) of the enzyme in the parasite cell may favor the formation of oligomers [54], which have higher activity than the dimers [25]. Molecular homology modeling of decameric PfTPx-1 protein from the crystal structure of mammalian Prx II revealed a conserved threonine residue (Thr88) that can be a phosphorylation target [31], likely triggering relaxation of the decameric structure and inactivation of the enzyme activity [25,52]. With regard to regulation of activity by overoxidation of Cysp [32,33], PfTPx-1 is inactivated at high peroxide concentrations and deviates from typical saturation kinetics in its catalytic reaction [14,45]. Homology modeling of PfTPx-1 with mammalian homologues suggested that PfTPx-1 possesses a redox-sensitive type of peroxidatic active-site structure, which enables the enzyme to act according to the floodgate model [29,32,33] (Fig. 1). These facts suggest that PfTPx-1 can act as an H2O2 regulator in the redox signaling model, similar to that proposed for other eukaryotic cells [25,27,30]. However, for recycling the overoxidized enzyme, there is no homologue of sulphiredoxin, which reduces sulfinylated enzyme [34], in the parasite genome. The parasite may possess a unique molecule or mechanism by which the overoxidized enzyme is reduced. If this is the case, overoxidation and recycling of Prx in malaria parasites needs further study. PfTPx-1 is expressed constitutively in the parasite cytoplasm throughout the erythrocytic stage, suggesting that it plays a housekeeping role including defense against oxidative stress, in the parasite cell [55]. PfTPx-1 reduces H2O2 via the PfTrx-1dependent catalytic cycle [56]. However, the expression profiles of these two proteins during erythrocytic development of the parasite do not overlap completely; PfTrx-1 shows lower expression during the ring stage. PfPlrx, which is an electron donor for the enzyme [52] and is expressed during ring stage, can be an alternative partner of PfTPx-1 in the early stage of

4

S. Kawazu et al. / Parasitology International 57 (2008) 1–7

parasite development. Expression of PfTPx-1 mRNA and protein can be induced in vitro with exogenously or endogenously applied oxidative stress [43]. An enhancer sequence that regulates transcription was mapped to the 5′-region of the PfTPx-1 gene; however, its impact in expression of the PfTPx-1 in response to oxidative stress is not clear (KomakiYasuda et al., unpublished data). Cultured P. falciparum carrying a disruption of the PfTPx-1 gene (knockout) showed lower growth than the wild-type (WT) P. falciparum when they are exposed to H2O2 and NO generated

by treatment of the cells with paraquat and sodium nitroprusside [57]. This finding suggests that PfTPx-1 protects the parasite cell from excessive oxidative and nitrosative stresses. However, in the absence of paraquat and sodium nitroprusside, the PfTPx1-knockout-P. falciparum could grow normally, suggesting that Prx is not essential for detoxification of H2O2 and NO, which are produced normally in the parasite cell under in vitro culture conditions. Analysis of the expression profile of TPx-1 in a rodent malaria parasite revealed that TPx-1 is expressed not only during the erythrocytic stage but also during the insect stage [58]. The constitutive expression of TPx-1 suggests that the molecule acts to defend against oxidative stress throughout the parasite lifecycle [43,52,58]. However, TPx-1-knockout-P. berghei (PbTPx-1 KO) develops normally in mouse erythrocytes and multiplies at a rate similar to that of WT parasite during experimental infection [59]. In contrast to mammalian Prx, PbTPx-1 does not prevent oxidation of parasite DNA [59]. Although these results suggest that Prx is not essential for P. berghei growth in mouse erythrocytes, the findings do not exclude the possibility that the gene is necessary for asexual growth of P. falciparum because these two parasites show differences in their life cycles. P. falciparum develops in erythrocytes sequestered in the microvasculature, where the parasite may be exposed to more severe stresses than in the circulating erythrocytes. PfTPx-1 may be required for preventing the parasite from oxidative and nitrosative stresses under the sequestration. The observation of protein nitration in brains from cerebral malaria patients suggests that NO is elevated at the site of sequestration [60]. This finding suggests that PfTPx-1 is as important as peroxynitrite reductase for protection of the parasites from nitrosative stresses. However, PbTPx-1 KO produced fewer (up to 60% of WT) gametocytes, which are sexual-stage parasites involved in the transmission between the mammalian host and the mosquito [59]. Gametocyte development can be induced by host factors or drug treatment, and there is consistent evidence for the involvement of signal transduction pathways in this process [61]. PbTPx-1 may participate in the signaling necessary to initiate gametocyte development in a manner similar to its mammalian homologue, which promotes cell division and differentiation. Homology modeling of PbTPx-1 with mammalian homologues suggests that the molecule possesses a Fig. 1. (A) Comparison of amino acid sequences of plasmodial and mammalian 2-Cys Prxs. P. falciparum PfTPx-1 [44], P. berghei PbTPx-1 [59], human Prx I (Protein data bank, PDB entry 1QMV) [71], and rat Prx II (PDB entry 1QQ2) [72] were compared. Sequences of three segments proposed to be specific for hydrogen peroxide-sensitive 2-Cys Prxs are shown [32]. The first (Loop–Helix) and third (C-Terminal Arm) segments are expected to undergo structural rearrangements during catalysis, while the second segment (310 Helix–Loop) is not. The sequence alignments suggest that both PfTPx-1 and PbTPx-1 are hydrogen peroxide-sensitive Prxs, and thus similar conformational changes are expected during catalysis. (B, C) The conformational change is computer modeled for P. falciparum PfTPx-1 based on the crystal structures of mammalian Prxs and the amino acid sequence alignments. Ribbon diagrams illustrate the peroxidatic active-site regions of (B) “fully folded” (modeled from 1QQ2) and (C) “locally unfolded” (modeled from 1QMV) conformers. In the first (Loop–Helix) and third (C-Terminal Arm) segments, the peroxidatic (C50 P : CysP) and resolving (C170 R : CysR) cysteine residues are depicted. Asterisk indicates the C-terminal unfolded disorder as observed in 1QMV.

S. Kawazu et al. / Parasitology International 57 (2008) 1–7

redox-sensitive type of peroxidatic active-site structure (Fig. 1). However PbTPx-1 does not appear to be involved in determination and development of sexually differentiated gametocytes because the male/female ratio of gametocyte and exflagellation activity of male gametocyte in PbTPx-1 KO are normal [59]. The PbTPx-1 affects oocyst maturation and sporozoite formation of the parasite in the vector mosquito Anopheles stephensi. The number of sporozoites that accumulated in the salivary gland of PbTPx-1 KO-infected mosquitoes is significantly decreased as compared to that in WT-infected mosquitoes (Yano et al., unpublished data). Immunoelectron microscopy revealed increased formation of 8-hydroxy-2′deoxyguanosine (8-OHdG), a marker of oxidative DNA damage, in nuclei of PbTPx-1 KO oocysts at an early stage of development (Yano et al., unpublished data). Although the specific mechanism by which the gene-knockout affects oocyst maturation is unknown, PbTPx-1 may be involved in formation of P. berghei sporozoites. The other typical 2-Cys Prx, PfTPx-2 was cloned from a cDNA library of P. falciparum as a Prx subfamily with mitochondria targeting signal at its N terminus [45]. The mitochondrial localization of PfTPx-2 and its expression during the trophozoite and schizont stages were confirmed microscopically with a specific antibody [55] and expression of a GFPfusion protein in the parasite cell [46]. Microarray data in PlasmoDB confirm that PfTPx-2 is expressed in gametocytes. PfTPx-2 prefers Trx-2, which localizes to mitochondria, to cytoplasmic Trx-1 as the electron donor [46]. Data from solution studies based on the crystal structure of the recombinant protein suggests that PfTPx-2 forms dimers with an intermolecular disulfide bridge and exists predominantly in a homodecameric form [46]. The physiologic role of PfTPx-2 with its electron donor in mitochondria remains unclear, and its role in the gametocyte, which possesses multiple mitochondria, will be interesting to study. 3.3. 1-Cys Prx (1-Cys-Prx and AOP) P. falciparum 1-Cys Prx (Pf1-Cys-Prx) is the other subfamily expressed abundantly in the parasite cytoplasm. During the trophozoite stage, Prx accounts for 0.5% of the total cellular protein [42]. The Pf1-Cys-Prx was identified independently by our group and an other group from the FCR-3 [42] and 3D7 P. falciparum strains [43], respectively; however, the amino acid sequences of the molecule from the two strains differ at several positions in the C terminus and the latter molecule is slightly larger. The discrepancy in the data was resolved by a third group who cloned the gene from the 3D7 strain [41]. Their sequence was identical to that of the FCR-3 strain (PF08-0131). We initially described a GSH-dependent peroxidase activity of Pf1Cys-Prx with H2O2 [42]; however, we could not confirm this result [62] and neither could an other group [41]. Trx-1- and Grx-dependent peroxidase activities were detected, but they were not sufficient such that the enzyme could be defined as Trx-1- or Grx-peroxidase [41,43]. It has been suggested that human 1-Cys Prx (Prx VI) is active with GSH only in the presence of a p-type glutathione S-transferase [63]; however,

5

such activity has not been confirmed for the parasite homologue [41,64]. Thus, the electron donor for Pf1-Cys-Prx remains controversial at this time. Ascorbate, which was recently proposed to be the substrate of Prx VI [65], should be tested for the parasite enzyme. Homology modeling of Pf1-Cys-Prx with the crystal structure of human Prx VI suggests that the parasite molecule forms a dimer [41] as has been reported for the human and Toxoplasma homologues (TgPrx2) [66]. However, Pf1-Cys-Prx lacks the oligomerization behavior observed for TgPrx2 [66]. Expression of Pf1-Cys-Prx is elevated in the parasite cytoplasm during the trophozoite and early schizont stages, and this expression profile suggests that this subfamily detoxifies metabolism-derived ROS, such as those released from heme iron. The trophozoite of the malaria parasite digests host hemoglobin to obtain amino acids for nutrition [7]. This process produces large quantities of heme in the parasite food vacuole, but the parasite does not possess a hemeoxygenase in the vacuole or cytoplasm [8]. For heme detoxification, we propose that Pf1-Cys-Prx competes with GSH for heme and decreases heme degradation by GSH and reduces the amount of heme-derived free iron that enters redox cycling and produces ROS in the parasite cytoplasm [8,62]. Additional functions of Pf1-Cys-Prx as a protector of parasite molecules from ironderived ROS and as an inhibitor of heme to association with the membrane have also been proposed [62]. Overexpression of Pf1-Cys-Prx yields an IC50 value for chloroquine higher than that in WT parasite (Table 1). Although the increase is not remarkable, it is statistically significant. This finding supports the idea that Pf1-Cys-Prx contributes at least partially to the parasite heme-detoxification mechanism. This hypothesis should be verified by observing how parasite with a disruption of the gene behaves in culture and in the host animals. PfAOP is another member of the 1-Cys Prx family identified from P. falciparum cDNA library presumably located in apicoplast because of the targeting signal found in the N terminus [41,47]. According to microarray data from PlasmoDB, the PfAOP gene is expressed during the erythrocytic stage and Table 1 Change in chloroquine (CQ) sensitivity of P. falciparum related to overexpression of 1-Cys Prx

Parent (FCR-3) 1-Cys Prx overexpressor Transfection control

IC50 (CQ μM) ± SD

P value

0.028 ± 0.0021 0.041 ± 0.0001 0.027 ± 0.0036

0.001 0.005

P. falciparum FCR-3 strain was transfected by electroporation [57] with the expression plasmid vector pHC1 [69], which carries the Pf1-Cys-Prx gene in the expression site. The parasite population that overexpresses Pf1-Cys-Prx (overexpressor) was established in vitro by selecting the transfectant with pyrimethamine (1 μM) in the culture medium. Expression of Pf1-Cys-Prx in the overexpressor at its trophozoite stage was estimated by Western blotting with a specific antibody [55], and the expression level was 4–8 times greater than that of the parent population (data not shown). The IC50 values for chloroquine (concentration of the drug that inhibits parasite growth in vitro by 50%) for the parasite populations were determined as described previously [70]. A parasite population transfected with pHC1 only and selected as the overexpressor was used as the transfection control. Data are mean ± standard deviation (SD) of four independent experiments, and differences were evaluated statistically with Student's t-test. P b 0.05 was considered statistically significant.

6

S. Kawazu et al. / Parasitology International 57 (2008) 1–7

expression is elevated during the trophozoite stage. This expression profile overlaps those of Grx1 and Plrx [67], which can donate electrons to this Prx [41]; however, whether Grx1 and Plrx can enter the apicoplast is unknown. Trx2 and Trx3, which also have the signal peptide, are candidate reductants for AOP in vivo [41,68]. Several lines of evidence based on the crystal structure and gel-filtration experiments suggest that PfAOP exists as a non-covalently linked dimer [47]. The physiologic role of PfAOP and its physiologic electron donor, which is localized at the apicoplast, are topics that remain to be clarified. 4. Concluding remarks The high abundance and distinct distribution of Prx subfamilies in different organelles suggests their importance in the antioxidant system of malaria parasites. Four Prx subfamilies, including two typical 2-Cys Prx and two 1-Cys Prx, have been identified in malaria parasites. TPx-1 is the major typical 2-Cys Prx in the parasite cell. This Prx is likely to be a key housekeeping protein for regulating redox homeostasis in the parasite cytoplasm. It is presumed from the phenotypes of the TPx-1-null parasite that Prx has dual roles in the parasite development as a regulator of the H2O2 signal during gametocyte development in the mammalian stage and as a protector of the parasite's macromolecules during the insect stage. If this is the case, how the parasite can make proper use of Prx according to the circumstances in the different hosts is a matter of great interest. In contrast, the cytoplasmic 1-Cys Prx, which constitutes approximately 0.5% of the total cellular protein during the trophozoite stage, appears to function specifically in moderating cellular redox imbalance, such as that created by heme metabolism. Whether these two Prx subfamilies can compensate for each other in the parasite cytoplasm is an interesting question still to be answered and may be answered by single- or doubleknockout experiments. Malaria parasites employ at least two additional Prx subfamilies, TPx-2 and AOP, in the mitochondrion and presumably in apicoplast, respectively. TPx-2 and AOP are characterized as typical 2-Cys Prx and 1-Cys Prx, respectively. Their physiologic roles are most likely related to the functions of the parasite organelles, and thus, these Prx subfamilies may be good targets for chemotherapy. Reverse genetics approaches will also help clarify the roles of such Prx subfamilies in the parasite life cycle. Further studies to elucidate the roles of these four Prx subfamilies in malaria parasites will further our understanding of the roles of these antioxidant proteins in asexual and sexual development of malaria parasites and may provide insights into novel ways to control this disease. Acknowledgements This work was supported by a Grant-in-Aid for Scientific Research (16590351 and 18590412 to S.I.K.) from the Japan Society for the Promotion of Science; Grants-in-Aid for Scientific Research (19550158 to H.O.) from the Ministry of Education, Culture, Sports, Science and Technology of Japan; and a grant for Precursory Research for Embryonic Science and Technology from the Japan Science and Technology Agency (to S.I.K.).

References [1] WHO. Global malaria situation. World Malaria Report 2005. Geneva: WHO and UNICEF; 2005. p. 5–17. [2] Finkel T. Oxidant signals and oxidative stress. Curr Opin Cell Biol 2003;15:247–54. [3] Sies H. Strategies of antioxidant defense. Eur J Biochem 1993;215:213–9. [4] Fujii J, Ikeda Y. Advances in our understanding of peroxiredoxin, a multifunctional, mammalian redox protein. Redox Rep 2002;7:123–30. [5] Becker K, Tilley L, Vennerstrom JL, Roberts D, Rogerson S, Ginsburg H. Oxidative stress in malaria parasite-infected erythrocytes: host–parasite interactions. Int J Parasitol 2004;34:163–89. [6] Müller S. Redox and antioxidant systems of the malaria parasite Plasmodium falciparum. Mol Microbiol 2004;53:1291–305. [7] Olliaro PL, Goldberg DE. The Plasmodium digestive vacuole: metabolic headquarters and choice drug target. Parasitol Today 1995;11:294–7. [8] Ginsburg H, Ward SA, Bray PG. An integrated model of chloroquine action. Parasitol Today 1999;15:357–60. [9] Postma NS, Mommers EC, Eling WM, Zuidema J. Oxidative stress in malaria; implications for prevention and therapy. Pharm World Sci 1996;18:121–9. [10] Han YS, Thompson J, Kafatos FC, Barillas-Mury C. Molecular interactions between Anopheles stephensi midgut cells and Plasmodium berghei: the time bomb theory of ookinete invasion of mosquitoes. EMBO J 2000;19:6030–40. [11] Radyuk SN, Klichko VI, Spinola B, Sohal RS, Orr WC. The peroxiredoxin gene family in Drosophila melanogaster. Free Radic Biol Med 2001;31:1090–100. [12] Flohé L, Jaeger T, Pilawa S, Sztajer H. Thiol-dependent peroxidases care little about homology-based assignments of function. Redox Rep 2003;8: 256–64. [13] Jaeger T, Flohé L. The thiol-based redox networks of pathogens: unexploited targets in the search for new drugs. Biofactors 2006;27:109–20. [14] Krauth-Siegel RL, Bauer H, Schirmer RH. Dithiol proteins as guardians of the intracellular redox milieu in parasites: old and new drug targets in trypanosomes and malaria-causing plasmodia. Angew Chem Int Ed Engl 2005;44:690–715. [15] Chae HZ, Chung SJ, Rhee SG. Thioredoxin-dependent peroxide reductase from yeast. J Biol Chem 1994;269:27670–8. [16] Rhee SG, Chae HZ, Kim K. Peroxiredoxins: a historical overview and speculative preview of novel mechanisms and emerging concepts in cell signaling. Free Radic Biol Med 2005;38:1543–52. [17] Chae HZ, Kang SW, Rhee SG. Isoforms of mammalian peroxiredoxin that reduce peroxides in presence of thioredoxin. Methods Enzymol 1999;300: 219–26. [18] McGonigle S, Dalton JP, James ER. Peroxidoxins: a new antioxidant family. Parasitol Today 1998;14:139–45. [19] Henkle-Dührsen K, Kampkötter A. Antioxidant enzyme families in parasitic nematodes. Mol Biochem Parasitol 2001;114:129–42. [20] Rahlfs S, Schirmer RH, Becker K. The thioredoxin system of Plasmodium falciparum and other parasites. Cell Mol Life Sci 2002;59:1024–41. [21] Torian BE, Flores BM, Stroeher VL, Hagen FS, Stamm WE. cDNA sequence analysis of a 29-kDa cysteine-rich surface antigen of pathogenic Entamoeba histolytica. Proc Natl Acad Sci U S A 1990;87:6358–62. [22] Tachibana H, Ihara S, Kobayashi S, Kaneda Y, Takeuchi T, Watanabe Y. Differences in genomic DNA sequences between pathogenic and nonpathogenic isolates of Entamoeba histolytica identified by polymerase chain reaction. J Clin Microbiol 1991;29:2234–9. [23] McGonigle S, Curley GP, Dalton JP. Cloning of peroxiredoxin, a novel antioxidant enzyme, from the helminth parasite Fasciola hepatica. Parasitology 1997;115:101–4. [24] Kumagai T, Osada Y, Kanazawa T. 2-Cys peroxiredoxins from Schistosoma japonicum: the expression profile and localization in the life cycle. Mol Biochem Parasitol 2006;149:135–43. [25] Wood ZA, Schröder E, Robin Harris J, Poole LB. Structure, mechanism and regulation of peroxiredoxins. Trends Biochem Sci 2003;28:32–40. [26] Finkel T. Oxygen radicals and signaling. Curr Opin Cell Biol 1998;10: 248–53.

S. Kawazu et al. / Parasitology International 57 (2008) 1–7 [27] Rhee SG, Kang SW, Jeong W, Chang TS, Yang KS, Woo HA. Intracellular messenger function of hydrogen peroxide and its regulation by peroxiredoxins. Curr Opin Cell Biol 2005;17:183–9. [28] Rhee SG. Cell signaling. H2O2, a necessary evil for cell signaling. Science 2006;312:1882–3. [29] Georgiou G, Masip L. Biochemistry. An overoxidation journey with a return ticket. Science 2003;300:592–4. [30] Kang SW, Rhee SG, Chang TS, Jeong W, Choi MH. 2-Cys peroxiredoxin function in intracellular signal transduction: therapeutic implications. Trends Mol Med 2005;11:571–8. [31] 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;277:25370–6. [32] Wood ZA, Poole LB, Karplus PA. Peroxiredoxin evolution and the regulation of hydrogen peroxide signaling. Science 2003;300:650–3. [33] Woo HA, Chae HZ, Hwang SC, Yang KS, Kang SW, Kim K, et al. Reversing the inactivation of peroxiredoxins caused by cysteine sulfinic acid formation. Science 2003;300:653–6. [34] Biteau B, Labarre J, Toledano MB. ATP-dependent reduction of cysteinesulphinic acid by S. cerevisiae sulphiredoxin. Nature 2003;425:980–4. [35] Neumann CA, Krause DS, Carman CV, Das S, Dubey DP, Abraham JL, et al. Essential role for the peroxiredoxin Prdx1 in erythrocyte antioxidant defence and tumour suppression. Nature 2003;424:561–5. [36] Lee TH, Kim SU, Yu SL, Kim SH, Park DS, Moon HB, et al. Peroxiredoxin II is essential for sustaining life span of erythrocytes in mice. Blood 2003;101:5033–8. [37] Han YH, Kim HS, Kim JM, Kim SK, Yu DY, Moon EY. Inhibitory role of peroxiredoxin II (Prx II) on cellular senescence. FEBS Lett 2005;579: 54897–902. [38] Wang Y, Feinstein SI, Manevich Y, Ho YS, Fisher AB. Peroxiredoxin 6 gene-targeted mice show increased lung injury with paraquat-induced oxidative stress. Antioxid Redox Signal 2006;8:229–37. [39] Bécuwe P, Gratepanche S, Fourmaux MN, Van Beeumen J, Samyn B, Mercereau-Puijalon O, et al. Characterization of iron-dependent endogenous superoxide dismutase of Plasmodium falciparum. Mol Biochem Parasitol 1996;76:125–34. [40] Sienkiewicz N, Daher W, Dive D, Wrenger C, Viscogliosi E, Wintjens R, et al. Identification of a mitochondrial superoxide dismutase with an unusual targeting sequence in Plasmodium falciparum. Mol Biochem Parasitol 2004;137:121–32. [41] Nickel C, Rahlfs S, Deponte M, Koncarevic S, Becker K. Thioredoxin networks in the malarial parasite Plasmodium falciparum. Antioxid Redox Signal 2006;8:1227–39. [42] Kawazu S, Tsuji N, Hatabu T, Kawai S, Matsumoto Y, Kano S. Molecular cloning and characterization of a peroxiredoxin from the human malaria parasite Plasmodium falciparum. Mol Biochem Parasitol 2000;109:165–9. [43] Krnajski Z, Walter RD, Müller S. Isolation and functional analysis of two thioredoxin peroxidases (peroxiredoxins) from Plasmodium falciparum. Mol Biochem Parasitol 2001;113:303–8. [44] Kawazu S, Komaki K, Tsuji N, Kawai S, Ikenoue N, Hatabu T, et al. Molecular characterization of a 2-Cys peroxiredoxin from the human malaria parasite Plasmodium falciparum. Mol Biochem Parasitol 2001;116:73–9. [45] Rahlfs S, Becker K. Thioredoxin peroxidases of the malarial parasite Plasmodium falciparum. Eur J Biochem 2001;268:1404–9. [46] Boucher IW, McMillan PJ, Gabrielsen M, Akerman SE, Brannigan JA, Schnick C, et al. Structural and biochemical characterization of a mitochondrial peroxiredoxin from Plasmodium falciparum. Mol Microbiol 2006;61:948–59. [47] 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;346:1021–34. [48] Kanzok SM, Schirmer RH, Türbachova I, Iozef R, Becker K. The thioredoxin system of the malaria parasite Plasmodium falciparum. Glutathione reduction revisited. J Biol Chem 2000;275:40180–6. [49] Rahlfs S, Fischer M, Becker K. Plasmodium falciparum possesses a classical glutaredoxin and a second, glutaredoxin-like protein with a PICOT homology domain. J Biol Chem 2001;276:37133–40.

7

[50] Deponte M, Becker K, Rahlfs S. Plasmodium falciparum glutaredoxinlike proteins. Biol Chem 2005;386:33–40. [51] Becker K, Kanzok SM, Iozof R, Fischer M, Schirmer RH, Rahlfs S. Plasmoredoxin, a novel redox-active protein unique for malaria parasites. Eur J Biochem 2003;270:1057–64. [52] Nickel C, Trujillo M, Rahlfs S, Deponte M, Radi R, Becker K. Plasmodium falciparum 2-Cys peroxiredoxin reacts with plasmoredoxin and peroxynitrite. Biol Chem 2005;386:1229–36. [53] Krnajski Z, Gilberger TW, Walter RD, Cowman AF, Müller S. Thioredoxin reductase is essential for the survival of Plasmodium falciparum erythrocytic stages. J Biol Chem 2002;277:25970–5. [54] Akerman SE, Müller S. 2-Cys peroxiredoxin PfTrx-Px1 is involved in the antioxidant defence of Plasmodium falciparum. Mol Biochem Parasitol 2003;130:75–81. [55] Yano K, Komaki-Yasuda K, Kobayashi T, Takemae H, Kita K, Kano S, et al. Expression of mRNAs and proteins for peroxiredoxins in Plasmodium falciparum erythrocytic stage. Parasitol Int 2005;54:35–41. [56] Kanzok SM, Rahlfs S, Becker K, Schirmer RH. Thioredoxin, thioredoxin reductase, and thioredoxin peroxidase of malaria parasite Plasmodium falciparum. Methods Enzymol 2002;347:370–81. [57] Komaki-Yasuda K, Kawazu S, Kano S. Disruption of the Plasmodium falciparum 2-Cys peroxiredoxin gene renders parasites hypersensitive to reactive oxygen and nitrogen species. FEBS Lett 2003;547:140–4. [58] Kawazu S, Nozaki T, Tsuboi T, Nakano Y, Komaki-Yasuda K, Ikenoue N, et al. Expression profiles of peroxiredoxin proteins of the rodent malaria parasite Plasmodium yoelii. Int J Parasitol 2003;33:1455–61. [59] Yano K, Komaki-Yasuda K, Tsuboi T, Torii M, Kano S, Kawazu S. 2-Cys peroxiredoxin TPx-1 is involved in gametocyte development in Plasmodium berghei. Mol Biochem Parasitol 2006;148:44–51. [60] Clark IA, Schofield L. Pathogenesis of malaria. Parasitol Today 2000;16: 451–4. [61] Talman AM, Domarle O, McKenzie FE, Ariey F, Robert V. Gametocytogenesis: the puberty of Plasmodium falciparum. Malar J 2004;3:24. [62] Kawazu S, Ikenoue N, Takemae H, Komaki-Yasuda K, Kano S. Roles of 1-Cys peroxiredoxin in haem detoxification in the human malaria parasite Plasmodium falciparum. FEBS J 2005;272:1784–91. [63] Manevich Y, Feinstein SI, Fisher AB. Activation of the antioxidant enzyme 1-CYS peroxiredoxin requires glutathionylation mediated by heterodimerization with πGST. Proc Natl Acad Sci U S A 2004;101:3780–5. [64] Fritz-Wolf K, Becker A, Rahlfs S, Harwaldt P, Schirmer RH, Kabsch W, et al. X-ray structure of glutathione S-transferase from the malarial parasite Plasmodium falciparum. Proc Natl Acad Sci U S A 2003;100:13821–6. [65] Monteiro G, Horta BB, Pimenta DC, Auqusto O, Netto LE. Reduction of 1-Cys peroxiredoxins by ascorbate changes the thiol-specific antioxidant paradigm, revealing another function of vitamin C. Proc Natl Acad Sci U S A 2007;104:4886–91. [66] Deponte M, Becker K. Biochemical characterization of Toxoplasma gondii 1-Cys peroxiredoxin 2 with mechanistic similarities to typical 2-Cys Prx. Mol Biochem Parasitol 2005;140:87–96. [67] Bozdech Z, Ginsburg H. Antioxidant defense in Plasmodium falciparum: data mining of the transcriptome. Malar J 2004;3:23. [68] Rahlfs S, Nickel C, Deponte M, Schirmer RH, Becker K. Plasmodium falciparum thioredoxins and glutaredoxins as central players in redox metabolism. Redox Rep 2003;5:246–50. [69] Crabb BS, Triglia T, Waterkeyn JG, Cowman AF. Stable transgene expression in Plasmodium falciparum. Mol Biochem Parasitol 1997;90: 131–44. [70] Hatabu T, Kawazu S, Suzuki J, Valenzuela RF, Villacorte EA, Suzuki M, et al. In vitro susceptibility of Plasmodium falciparum isolates to chloroquine and mefloquine in southeastern Mindanao Island, the Philippines. Southeast Asian J Trop Med Public Health 2003;34:546–51. [71] Schroder E, Littlechild JA, Lebedev AA, Errington N, Vagin AA, Isupov MN. Crystal structure of decameric 2-Cys peroxiredoxin from human erythrocytes at 1.7 A resolution. Structure 2000;8:605–15. [72] Hirotsu S, Abe Y, Okada K, Nagahara N, Hori H, Nishino T, et al. 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;96:12333–8.