The origin of catalase (EC 220.127.116.11), the enzyme that metabolizes H2O2 but also reacts with a multitude of other substrates, can be traced back to the 19th century (Figure 1) when Thénard discovered H2O2 and suspected that its tissue degradation in living organisms was the result of a ‘special’ substance activity (Thénard, 1811). Schönbein showed that a ‘ferment’ can detoxify H2O2 (Schönbein, 1863) and later on Loew gave the name ‘catalase’ to the enzyme converting H2O2 into water and oxygen (Loew, 1900). Loew found the presence of this enzyme in many organisms ranging from plants to mammals. In the 1920s, Warburg and coworkers demonstrated, by using cyanide as enzyme inhibitor, that the active site of catalase contains an iron atom (Warburg, 1923). Furthermore, Stern showed that the hemin group of the enzyme can react with compounds such as cyanides, sulfides, fluorides and he also showed that the enzyme active group had a ferric complex identical to the protoporphyrin found in the hemoglobin of red blood cells (Stern, 1937). Thereafter, Sumner and Dounce (1937) purified and crystallized bovine catalase.
The advances in biochemistry facilitated further elucidation about the mechanisms of the ‘catalase’ enzymatic reaction. Thus, the work carried out by Chance’s lab led to the discovery of the formation of the ‘Compound I’ that occurred during the reaction between catalase and the first molecule of H2O2. A few years later, he also discovered the ‘compounds II and III’ (Chance, 1948, 1949; Chance et al., 1952). In the 1960s, it was elucidated the role of key residues in the active site of the enzyme, such as the distal histidine, and their importance for the stabilization of the tertiary structure was discussed by Nakatani (1961). In 1970, catalase Compound I was identified in intact eukaryotic cells, proving the existence of H2O2 in normal aerobic metabolism (Sies and Chance, 1970). Kirkman and Gaetani reported that NADPH was the enzyme cofactor bound to catalase (1984), which was further confirmed following X-ray analysis of catalase structures (Fita and Rossmann, 1985a). Finally, recombinant phage clones containing the human catalase gene were isolated and characterized (Quan et al., 1986) and the use of molecular biology techniques have led to major advances in the understanding of catalase regulation mechanisms and the role that this enzyme could play in numerous biological processes.
In this context, Nenoi et al. (2001) reported that in the upstream catalase promoter, there is a region containing CCAAT and GGGCGG boxes where transcription factors like Nuclear factor Y (NF-Y) and Specificity protein 1 (Sp1) can bind to the catalase promoter regulating the transcriptional activation of the human catalase gene. Recently, Glorieux et al. (2016a) identified a new regulatory region in the human catalase promoter, in which chromatin remodeling is required to regulate catalase expression by retinoic acid receptor alpha (RARα) and JunB transcription factors. This novel regulatory mechanism is involved during cancer cells adaptation to chronic exposure to H2O2 and may have therapeutic consequences for various diseases and metabolic disorders.
Types of catalases
With the increasing number of complete sequences available, distinct homologies were detected and catalases came to be classified in three groups based on their structure and function. The first and the second group contain heme-containing enzymes, namely typical or true catalases and catalase-peroxidases, whereas the third group contains (non-heme) manganese catalases (Zamocky and Koller, 1999; Zamocky et al., 2008, 2010).
The members of this largest group are found in aerobically respiring organisms. In contrast, in anaerobic bacteria, catalases proteins are generally not expressed. Most of these catalases are homotetramers, between 200 and 340 kDa in size and contain four prosthetic groups. In the majority of true catalases, a ferric protoporphyrin IX was found in the active center (namely heme b, similar to the prosthetic group of human hemoglobin). Some variants do exist such as ‘heme d’ groups which can reside in some typical catalases.
Following phylogenetic analyses the typical catalases can also be divided into three main clades. Clade 1 contains bacterial, algal and plant catalases with small-subunit size (55–69 kDa) using heme b as the prosthetic group. Clade 2 regroups bacterial and fungal catalases and has a large subunit size (75–84 kDa), with heme d as the prosthetic group and an additional ‘flavodoxin-like’ domain. Clade 3 is the most abundant subfamily; catalases from this subgroup are found in archaebacterial, fungi, protists, plants and animals. The human catalase belongs to this clade and is characterized by a small subunit (62 kDa), with heme b as its prosthetic group and NADPH as cofactor.
These proteins have been found in fungi, archeobacteria and bacteria. Their molecular weight varies between 120 and 340 kDa and they are generally homodimers. The catalase activity (degrading hydrogen peroxide) is less efficient than in typical catalases but catalase-peroxidases have a better affinity for their substrate H2O2. Catalase-peroxidases are also significantly more sensitive than typical catalases to inactivation by pH and temperature. The well-known horseradish peroxidase, currently employed in immunoblotting experiments, is one example of catalase-peroxidase.
These enzymes have been found exclusively in bacteria. Manganese catalases utilize two manganese ions in the active site, they can form oligomeric structures measuring between 170 and 210 kDa and have no significant homology with either typical catalases or catalase-peroxidases. The catalytic reaction is completely different to other types of catalases. The dimanganese core is equally stable Mn2+–Mn2+ or Mn3+–Mn3+. Like typical catalases, the catalase reaction occurs in two-step.
Structure of human catalase
Human catalase contains four identical subunits of 62 kDa, each subunit containing four distinct domains and one prosthetic heme group (Nagem et al., 1999; Zamocky and Koller, 1999; Putnam et al., 2000). The four domains include: (1) a N-terminal arm which contains a distal histidine, an essential amino acid for the catalase reaction; (2) a β-barrel domain that contains eight β-barrels arranged in an antiparallel fashion with six α-helical insertions, conferring the hydrophobic core of the protein necessary for the tri-dimensional structure of the enzyme; (3) a connection domain which contains the tyrosine residue that binds the heme group; and finally (4) an α-helical domains, which is important for NADPH binding.
Although amino acid sequences do not have high identities between all typical catalases, the tridimensional structure is highly conserved. The tertiary structure of the β-barrel domain, the connection domain and the zone neighboring the distal histidine are highly conserved. The α-helical domain is moderately conserved between species and some typical catalases do not bind the cofactor NADPH.
The investigation of inhibitory mechanisms by which cyanide and 3-amino-1,2,4-triazole (ATA) inhibit human catalase allowed to understand the enzyme activity (Putnam et al., 2000). Cyanide nitrogen blocks heme access to other potential ligands. It interacts with the distal histidine and an asparagine residue suggesting that it competes with hydrogen peroxide for heme binding. Meanwhile, ATA interacts with the distal histidine leading to an adduct formation and thereby blocks the catalase reaction.
Mechanism of ‘catalase’ reaction
During the enzymatic reaction leading to H2O2 destruction, catalase is first oxidized to a hypervalent iron intermediate, known as compound I (Cpd I), which is then reduced back to the resting state by a second H2O2 molecule.
The first reaction is characterized by the oxidation of the heme protein by a single H2O2 molecule leading to the formation of Cpd I, an oxoferryl porphyrin cation radical (Jones and Dunford, 2005). Once Cpd I is formed, it reacts rapidly with a second molecule of H2O2 to generate H2O and O2 in a two-electron redox process. This second reaction is particularly efficient in some catalases compared to other heme proteins such as myoglobin (Matsui et al., 1999). Labeling studies have shown that both H2O and O2 molecules are formed from the same molecule of H2O2 (Vlasits et al., 2007). Fita and Rossmann (1985b) proposed that two molecules of H2O2 were sequentially transferred to the oxoferryl group of Cpd I, where the distal histidine residue plays a role of acid-base catalyst. Although mutation of distal histidine suppresses the ability to form Cpd I (Nakatani, 1961), some authors claimed that reaction occurs as a direct mechanism and histidine does not play a crucial role in catalysis (Kato et al., 2004).
In the presence of one-electron donors (such as phenols, ferrocyanide, salicylic acid, NO, superoxide anions) and low H2O2 concentrations, Cpd I may undergo a one-electron reduction towards the inactive compound II (Cpd II) intermediate, which transforms back to the resting state through another one-electron reduction step (reviewed in Bauer, 2015). Both Cpd I and Cpd II are generally described as an oxoferryl-heme species (Rovira, 2005). In the case of Cpd I, the porphyrin bears a cation radical (O=FeIV-heme˙+), while Cpd II lacks the porphyrin cation radical (O=FeIV-heme). Thus, Cpd II is best described as a hydroxoferryl bond (HO-FeIV-heme) instead of the traditional oxoferryl species, consistent with the fact that a proton is released upon conversion of Cpd I to Cpd II:
At higher H2O2 concentrations, NADPH prevents the generation of Cpd II by participating in a two-electron reduction process (Kirkman et al., 1999). In the presence of another one-electron donor, Cpd II will return to a resting state. But in presence of a H2O2 molecule, Cpd II will be transformed back to compound III (Cpd III), an inactive intermediate (Gabdoulline et al., 2003). In this intermediate state, iron is at an oxyferrous state (O2-FeII-heme). Then, Cpd III goes back to a resting state or leads to the inactivation of the catalase.
Cellular and tissue catalase distribution
Regarding catalase localization within the cell, it should be noted that catalase is mainly located in peroxisomes because it contains a sequence signal recognized by some peroxisome receptors. Contrary to mitochondria, proteins located within peroxisomes are all of nuclear origin and should be imported. Indeed, it is generally accepted that catalase monomers are imported into peroxisomes where tetramerization and heme addition occurs (Lazarow and De Duve, 1973). The apoprotein (monomer) enters into the peroxisome by a peroxisome-targeting signal sequence (PTS) present on the carboxy-terminal tail of catalase. The most common targeting signal is the SKL (serine-lysine-leucine) but catalase is characterized by a different signal, namely the KANL (lysine-alanine-asparagine-leucine) signal (Purdue and Lazarow, 1996). Proteins bearing these signals are recognized by the PTS1 receptor, called PEX5p for humans. Some diseases related to defects in peroxisome biogenesis, such as Zellweger syndrome, are characterized by mutations in the PEX5p receptor and cellular H2O2 overproduction due to a catalase default import in peroxisomes (Wanders et al., 1984). This syndrome is most commonly called ‘the syndrome of empty peroxisome’. It has been shown that overexpressing receptor mutants in PEX5-deficient CHO cells, drastically reduce the import of proteins such as catalase (Shimozawa et al., 1999). Some authors have observed that catalase with the SKL signal and not KANL had a better import capacity and could be transported into the peroxisome even in the case of mutations in the PTS1 receptor (Koepke et al., 2007). It has been shown that PEX5p recognizes catalase which is already folded and interacts with other receptors such as PEX13p for proper import into the peroxisome (Otera and Fujiki, 2012). Studies are underway to validate, through clinical trials, whether catalase SKL has a therapeutic potential for diseases with peroxisome biogenesis disorders. Recently, it has been reported that PEX19p, an essential protein for peroxisome biogenesis, interacts with Valosin-containing protein (VCP) and regulates the catalase cytoplasmic localization, a potential feedback mechanism modulating H2O2 levels (Murakami et al., 2013).
Interestingly, the existence of a cytosolic catalase, in its active tetrameric conformation, has been reported (Middelkoop et al., 1993) with varied percentage from one cell type to another, and different function to peroxisome catalase. Indeed, catalase may bind cytosolic proteins such as Grb2 and SHP2, to protect them from potential oxidative damage (Yano et al., 2004a,b). These proteins are linked to the membrane by a pleckstrin homology (PH) domain, so it is common to find catalase in fractions including membrane proteins and membrane-associated proteins. In this context, it has been shown that catalase can be localized at the cytoplasmic membrane, specifically at the surface of cancer cells (Bauer, 2012). This locally high expression of catalase on the membrane of tumor cells is in line with the findings by Deichman’s group that showed that tumor progression in vivo is dependent on increased resistance towards exogenous H2O2 (Deichman, 2000, 2002). Furthermore, localized expression of catalase on the membrane of tumor cells is not in disagreement with the finding of lower total catalase concentration in malignant cells, as the membrane comprises a minority of the total cellular material.
These observations open the way for new anti-cancer therapies targeting catalase with specific antibodies (Bauer, 2012; Bauer and Motz, 2016) or exogenous singlet oxygen (Riethmüller et al., 2015; Bauer and Graves, 2016) to induce apoptosis by reactivation of intercellular HOCl and/or NO/peroxynitrite signaling after catalase inhibition or inactivation. Furthermore, the modulation of the intracellular NO concentration has been shown to lead to the generation of cell-derived singlet oxygen that inactivates tumor cell protective catalase and reactivates intercellular ROS/RNS-mediated apoptosis-inducing signaling (Bauer, 2015; Scheit and Bauer, 2015).
In addition to classical intracellular catalase and membrane-associated catalase of tumor cells (Heinzelmann and Bauer, 2010; Böhm et al., 2015), the release of soluble catalase from tumor cells has also been reported (Sandstrom and Buttke, 1993; Moran et al., 2002; Böhm et al., 2015). This soluble extracellular catalase was protective for the tumor cells.
Finally, catalase has been also localized in the mitochondria of rat cardiomyocytes (Radi et al., 1991).
The human catalase is expressed in every organ and the highest levels of activity are measured in the liver, kidney and red blood cells (Winternitz and Meloy, 1908). In erythrocytes, a high production of H2O2 is generated due to oxygen transport and catalase is responsible for more than 50% of the H2O2 turnover (Mueller et al., 1997).
Biological functions of catalase and related diseases
The first function assigned to catalase is the dismutation of H2O2 into oxygen and water without consummation of endogenous reducing equivalents, an important role in cell defense against oxidative damage by H2O2. To note that H2O2 is not only toxic by its ability to form other ROS, like hydroxyl radical through the Fenton reaction (Fenton, 1894) but as was nicely recently reviewed by Sies, H2O2 acting as a second messenger is involved in many biological processes including changes of morphology, proliferation, signaling (i.e. NF-κB), apoptosis, etc (Sies, 2017). In addition to its dominant ‘catalatic’ activity (decomposition of H2O2), catalase can also act in its peroxidatic mode, i.e. decomposition of small substrates such as methanol, formate, azide, hydroperoxides (Sies, 1974; Chance et al., 1979; Johansson and Borg, 1988), and in case of ethanol, it is also capable of oxidize it to acetaldehyde contributing to its liver metabolism (Keilin and Hartree, 1945; Thurman et al., 1972; Oshino et al., 1973). It has also been reported that catalase may decompose peroxynitrite (Gebicka and Didik, 2009; Heinzelmann and Bauer, 2010), oxidize nitric oxide to nitrite (Wink and Mitchell, 1998; Brunelli et al., 2001). A discrete balance between oxidation of NO by compound I of catalase and inhibition of catalase by NO through formation of a CAT-FeIIINO complex has been reported (Brown, 1995). Catalase also exhibits low oxidase activity (O2-dependent oxidation of organic substrates) (Vetrano et al., 2005). Thus, catalase may also have additional roles such as the detoxification or activation of toxic and anti-tumor compounds. For instance, catalase has been detected in mouse oocytes most likely to protect the genome from oxidative damage during meiotic maturation (Park et al., 2016).
Within this framework, several studies have shown a change in catalase expression in cancer cells became resistant to chemotherapies (Akman et al., 1990; Kim et al., 2001; Kalinina et al., 2006). Thus, a potential role of catalase during the acquisition of cancer cell resistance to chemotherapeutic agents was explored by overexpressing the human enzyme in MCF-7 cells, a human derived breast cancer cell line (Glorieux et al., 2011). No particular resistance against conventional chemotherapies like doxorubicin, cisplatin and paclitaxel was observed in cells overexpressing catalase but they were more resistant to the pro-oxidant effect induced by an H2O2-generating system (Glorieux et al., 2011).
In addition, catalase mitochondria overexpression in mice enables an increase of lifespan by 20% (Schriner et al., 2005). In such animals, the development of mitochondrial deletions was reduced and heart disease and the onset of cataracts were delayed.
Regarding catalase down-regulation, no particular sensitivity was observed as catalase-deficient mice are viable and fertile (Ho et al., 2004). They develop normally with a normal hematological profile, but after trauma the mitochondria shows defects in the oxidative phosphorylation. Note that humans may also be deficient in catalase, a condition known as acatalasemia that is characterized by a low catalase rate, but it is still rare and usually benign (Goth et al., 2004).
In this context, there are benign polymorphisms of the catalase gene for which no change of catalase expression or activity was detected (Goth et al., 2004). They include single nucleotide substitutions in the promoter, 5′ untranslated region, intron 1, exon 1, exon 9 and 10 (Goth et al., 2004). To note that the catalase gene encodes one single protein of 526 amino acids and the single locus has been mapped to chromosome 11p13 (Wieacker et al., 1980). The length of the catalase gene is 34 kb, it contains 12 introns and 13 exons generating a mRNA of 2286 bp (Quan et al., 1986).
Conversely, some catalase mutations provoke changes in either catalase expression or activity and may be associated with some diseases. For instance, a higher transcriptional activity in two human cancer cell lines, namely HepG2 and K562 cells, was observed in the case of common functional C-T substitution polymorphism in the promoter region (−262) of the human catalase gene (Forsberg et al., 2001) but no mutations have been detected in the coding sequence, to our knowledge, in patients suffering from cancer. Recent studies have focused on the associations of catalase polymorphisms with various types of cancer but many inconsistent results about the relationship between the catalase gene polymorphism and cancer risk were reported. Recently, two meta-analyses pointed out a correlation exists between this polymorphism C-262T and prostate cancer (Liu et al., 2016; Wang et al., 2016).
Different catalase mutations in patients can cause decreased catalase activity leading to increased H2O2 concentrations in the blood and tissues. Table 1 shows the polymorphism of the catalase gene in patients suffering from hypocatalasemia (about 50% of catalase activities) or acatalasemia (less than 10% of catalase activities). Depending on the mutations, such patients may be subject to an increased risk of type 2 diabetes, vitiligo and increased blood pressure (Goth et al., 2004). When the mutations are located in exons as in Japanese and Hungarian acatalasemia, a truncated or mutated catalase is synthetized and functionally less active. Takahara was the first to describe acatalasemia and this pathology is often benign but Japanese patients can suffer from oral gangrenes and esophageal ulcerations (Takahara’s disease), probably promoted by H2O2 generated by phagocytic cells and bacterial actions (Takahara, 1952).
Decreased activity of catalase has also been observed in various genetic alterations, for example, loss of alleles (i.e. loss of heterozygosity) of the catalase gene in non-small-cell lung cancer cells (Ludwig et al., 1991; Fong et al., 1994; Shipman et al., 1998) or deletion of chromosome 11p, as has been observed in children affected by Wilms’ tumor, aniridia, gonadoblastoma and retardation (WAGR) syndrome (Dufier et al., 1981; Gregoire et al., 1983; Barletta et al., 1985; Michalopoulos et al., 1985; van Heyningen et al., 1985).
Regulation of catalase expression in cancer cells
It is generally accepted that the cellular maintenance of redox homeostasis is controlled by a complex network of antioxidant enzymes (i.e. superoxide dismutases and glutathione peroxidases) whose expression is under the fine-tuning control of the Keap1-Nrf2 signaling pathway (Menegon et al., 2016). Nevertheless, the molecular mechanisms regulating the expression of catalase – the oldest known and first discovered antioxidant enzyme – are independent of this pathway and not totally elucidated. Therefore, the fine-tuning regulation of this enzyme should be prior elucidated in order to find a new approach to modulate the antioxidant status in cancer cells.
Interestingly, despite the existence of diverse protection mechanisms against oxidant injuries, a consensus emerged in the scientific literature about an alteration of redox homeostasis within tumor cells. Indeed, they produce large amounts of ROS that are involved in the maintenance of genetic instability favoring cancer cell proliferation. Meanwhile, altered expression levels of catalase have been reported in cancer tissues as compared to their normal counterparts. Thus, as compared to normal tissues of the same origin, some authors reported an increased catalase expression in tumors (Sander et al., 2003; Hwang et al., 2007; Rainis et al., 2007), whereas other studies showed a catalase down-regulation (Marklund et al., 1982; Baker et al., 1997; Lauer et al., 1999; Chung-man et al., 2001; Cullen et al., 2003; Kwei et al., 2004), indicating that cancer cells are frequently more sensitive to an oxidative stress. For instance, we have reported an important decrease of catalase activity in different cancer cell lines, as shown in Table 2 (Verrax et al., 2009; Beck et al., 2011a; Glorieux et al., 2011). Briefly, catalase levels can vary after short treatments to H2O2 (Rohrdanz and Kahl, 1998; Rohrdanz et al., 2001; Sen et al., 2003) and catalase expression is modified in cancer cell lines rendered resistant to chronic exposures to H2O2 (Kasugai and Yamada, 1992; Nenoi et al., 2001) or certain chemotherapeutic compounds such as doxorubicin (Ramu et al., 1984; Akman et al., 1990; Kim et al., 2001; Kalinina et al., 2006). Although mechanisms controlling catalase expression have been partially elucidated, the decreased catalase expression in cancer cells still remains an unanswered question.
The regulation of catalase expression in cancer cells is a complex process because different levels of regulation are thought to be involved. In a recent review, we discussed the different mechanisms playing a potential role in the regulation of its expression in both healthy and tumor cells (Glorieux et al., 2015). They include transcriptional regulation, represented by the activity of transcription factors that induce or repress the transcriptional activity of catalase promoters, post-transcriptional regulation (mRNA stability) and post-translational modification (phosphorylation and ubiquitination of the protein). In addition, epigenetic (DNA methylation, modifications of histones) changes or genetic alterations can also be involved playing a role in governing proper levels of catalase activity in these cells.
Regarding transcription it should be noted that the catalase gene has all the characteristics of a housekeeping gene (no TATA box, no INR sequence, high GC content in promoter) and a core promoter which is highly conserved among species (Quan et al., 1986; Nakashima et al., 1990; Reimer et al., 1994). In this core promoter, the presence of DNA binding sites for transcription factors like NF-Y and Sp1 has an essential role in the positive regulation of catalase expression (Nenoi et al., 2001). Additional transcription factors have also been involved in this regulatory process. In fact, there is strong evidence that the protein Akt/PKB in the PI3K signaling pathway plays a major role in the expression of catalase by modulating the activity of FoxO3a (Turdi et al., 2007; Venkatesan et al., 2007; Akca et al., 2013). Therefore, targeting PI3K/Akt/mTOR (LoPiccolo et al., 2008; Rodon et al., 2013) may be an efficient way to increase the expression of catalase in tumors and inhibit tumor cell growth.
In this last decade, other transcription factors (PPARγ, Oct-1, etc.) as well as genetic, epigenetic and post-transcriptional processes are emerging as crucial contributors to the negative regulation of catalase expression (Glorieux et al., 2015). Specifically, we investigated the transcriptional regulatory mechanism controlling catalase expression in human mammary cell lines. To this end, we have made a human breast MCF-7 cancer cell line resistant to oxidative stress, the so-called Resox cells. These cells show decreased ROS basal levels and an increased activity of some antioxidant enzymes, notably catalase (Dejeans et al., 2012; Glorieux et al., 2015, 2016b). A novel promoter region, responsible for the regulation of catalase expression, was identified at −1518/−1226 locus in Resox cells (Glorieux et al., 2016a). The AP-1 family member JunB and retinoic acid receptor alpha (RARα) mediate catalase transcriptional activation and repression, respectively, by controlling chromatin remodeling through a histone deacetylases-dependent mechanism (Glorieux et al., 2016a). Indeed, RARα and JunB act in collaboration with transcription factors or other proteins on either a closed- or an open-promoter chromatin status regulating the expression of catalase in breast cancer cells (Figure 2). Thus, cancer adaptation to oxidative stress, regulated by transcriptional factors through chromatin remodeling appears as a new mechanism to target cancer cells.
Oxidative stress-based therapies against cancer
As previously mentioned, cancer cells are generally deficient in antioxidant enzymes, thus, any increase in ROS levels would be a menace to the precarious redox balance of cancer cells making them vulnerable to an additional oxidative stress. Given that weakness, the loss of redox homeostasis represents an interesting target for research and development of new molecules with antitumor activity and numerous drugs are currently being clinically evaluated. Therefore, several strategies have been developed looking for the disruption of tumor cell redox homeostasis and a subsequent cancer cell death (Demizu et al., 2008; Trachootham et al., 2009; Verrax et al., 2009).
One of these strategies is the use of ROS-generating compounds such as arsenic trioxide (ATO), currently employed against promyelocytic leukemia (Valenzuela et al., 2014; Lo-Coco et al., 2016), or doxorubicin, an anthracycline used for the treatment of several types of cancer (i.e. breast cancer). Indeed, the impairment of mitochondrial function due to increased levels of superoxide anion is supposed to be the main mechanism of both chemotherapeutic drugs (Thayer, 1977; Pelicano et al., 2003). A decrease in antioxidant levels has also been proposed. For instance, glutathione (GSH) levels may be decreased either by its direct binding with phenylethylisothiocyanate (PEITC) and sulphoraphane (Trachootham et al., 2009) or by inhibition of γ-glutamylcysteine synthetase activity (a key enzyme involved in GSH synthesis) as done by Buthionine sulfoximine (Griffith and Meister, 1979). The development of specific inhibitors of thioredoxin and thioredoxin reductases was also carried out. Inhibitors of thioredoxin, such as PX-12, were shown to have potent antitumor activities (Welsh et al., 2003; Baker et al., 2006). Conversely, the overexpression of Trx1 is correlated with resistance to anti-cancer drugs (Baker et al., 2006; Kaimul et al. 2007).
As quinones display redox cycling abilities thus generating ROS (Kappus and Sies, 1981), the association of menadione (a naphthoquinone derivative) and ascorbate (Figure 3), was employed to trigger tumor cell death (Verrax et al., 2011b). This association (Asc/Men) has been shown to have potent in vitro and in vivo antitumor activities enhancing as well the therapeutic effect of currently used anticancer drugs (Taper et al., 1987; Buc Calderon et al., 2002; Verrax et al., 2003). We hypothesized that H2O2, issued from the redox cycling, is the oxidant species responsible for antitumor effects observed both in vitro and in vivo (Verrax et al., 2006, 2007; Verrax and Buc Calderon, 2009). The induced oxidative stress provokes cell necrosis by a wide variety of processes including ATP depletion (Verrax et al., 2004, 2005, 2011a), disruption of Ca2+ homeostasis (Dejeans et al., 2010) and loss of HSP90 chaperone activity (Beck et al., 2009, 2011b, 2012). Based on the vulnerability of tumor cells to an oxidative stress, we have induced the alteration of their intracellular redox homeostasis as a new strategy in the research and development of new antitumor drugs (Benites et al., 2010; Arenas et al., 2013; Valderrama et al., 2015). A similar approach has been recently developed by using the SnFe2O4 nanocrystals, a heterogeneous Fenton catalyst, which once internalized into the cancer cells, convert H2O2 into hydroxyl radicals inducing apoptotic cell death. In normal cells, the oxidative injury induced by SnFe2O4 is prevented by catalase (Lee et al., 2017).
As ATO increases the levels of ROS (Valenzuela et al., 2014) and Asc/Men increases the cytotoxicity of chemotherapeutic drugs (Taper et al., 1987), we formulated the following hypothesis: Asc/Men potentiates ATO-mediated cytotoxicity (Figure 4). ATO decreases catalase expression most likely by activating Akt pathway and/or inducing RARα, meanwhile Asc/Men generates H2O2. Despite that Resox cells display high antioxidant defenses (Dejeans et al., 2012; Glorieux et al., 2015, 2016b), an enhanced cell death was observed when they were exposed to a mixture containing sublethal doses of Asc/Men and ATO. This is likely due to a decreased transcriptional activity of the human catalase −1518/+16 promoter caused by ATO resulting in less catalase protein (Glorieux et al., 2017, unpublished results).
Under stress conditions, the antioxidant enzyme catalase plays a major role by detoxifying H2O2. As consequences, a change of its activity or expression will lead to pathological processes as Zellweger syndrome, acatalasemia or WAGR syndrome. The subcellular localization of catalase is mainly peroxisomal but a shuttle between this organelle and cytoplasm exists and may be involved in the protection of key cellular elements (i.e. proteins, chromosomes) against an oxidative damage.
Our group and others demonstrated that catalase expression is also altered in cancer cells, most likely to favor cell proliferation by inducing genetic instability and activation of oncogenes. The regulation of catalase expression appears to be mainly controlled at transcriptional levels although other mechanisms may also be involved. In addition of transcription factors like Sp1 and NF-Y, JunB and RARα transcription factors are crucial regulators in breast cancer cells by recruiting proteins involved in transcriptional complexes and chromatin remodeling.
Therefore, catalase can be a future therapeutic target in the context of cancer by using pro-oxidant approaches.
The authors thank Professor Helmut Sies for the splendid discussion and his precious input. This project was funded by FNRS-Televie Grant (grant n° 7.4575.12F).
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Published Online: 2017-04-06
Published in Print: 2017-09-26