The Uniqueness of the Trypanosoma cruzi ...

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Current Pharmaceutical Design, 2011, 17, 2074-2099

The Uniqueness of the Trypanosoma cruzi Mitochondrion: Opportunities to Target New Drugs Against Chagas´ Disease Lisvane Silva Paes§, Brian Suárez Mantilla §, María Julia Barisón§, Carsten Wrenger and Ariel Mariano Silber Departamento de Parasitologia, Instituto de Ciências Biomédicas, Universidade de São Paulo, São Paulo, Brasil Abstract: Trypanosoma cruzi is the causative agent of Chagas´ disease, which affects some 8 – 10 million people in the Americas. The only two drugs approved for the etiological treatment of the disease in humans were launched more than 40 years ago and have serious drawbacks. In the present work, we revisit the unique characteristics of T. cruzi mitochondria and mitochondrial metabolism. The possibility of taking advantage of these peculiarities to target new drugs against this parasite is also discussed.

Keywords: Trypanosoma cruzi, mitochondria, metabolism, bioenergetics, oxidative stress, chemotherapy. INTRODUCTION: CHAGAS´ DISEASE AND ITS ETIOLOGICAL AGENT Chagas´ disease (or American trypanosomiasis) belongs to a group of neglected diseases according to the WHO. It is an infectious disease caused by the protozoan parasite, Trypanosoma cruzi. This infection affects 8 to 12 million people on the American continent, exposing approximately 40 million people to the risk of infection. The infection is mainly transmitted by a triatomine insect vector present in the endemic areas (from southern Argentina and Chile to the southern USA). However, other modes of transmission are also well established. These modes of transmission include the following in decreasing order of epidemiological relevance; the transmission via blood transfusion and solid organ transplantations from infected donors to healthy receptors; vertical transmission during pregnancy or birth; and contamination through laboratory accidents. Oral transmission (through the ingestion of contaminated foods or liquids) is also well demonstrated and is considered epidemiologically relevant [1]. Chagas´ disease presents two clinical forms: acute and chronic. The acute phase starts immediately after contamination and is characterized by a conspicuous parasitemia and the absence of an immune response. This phase lasts for several weeks to several months and is generally asymptomatic. In some cases, the acute phase may present with non-specific symptoms, such as fever and headaches; eventually, a more complex symptomatology can be observed, including lymphadenopathy and splenomegaly, myalgia, malaise, muscle pains, sweating and hepatosplenomegaly. In the more severe presentations, heart failure from myocarditis or pericardial effusion, or meningoencephalitis can lead to death. The chronic phase follows the acute phase and is characterized by the absence of noticeable parasitemia and the presence of a strong cellular and humoral immune response, which is unable to eliminate the parasite. The chronic phase of the disease lasts for the rest of the patient´s life and is asymptomatic in most cases (indeterminate form). In 10 – 30% of cases, patients can develop one of the symptomatic forms of the chronic disease, which mainly include the cardiac form (manifested as arrhythmias, thromboembolism and cardiomegalia caused by inflammatory infiltrations) or the digestive form (which can consist of megaesophagus, which causes dysphagia and regurgitation and megacolon, which causes severe constipation). In immunocompromised patients, severe compromise of the central nervous system can also occur [1, 2]. *Address correspondence to this author at the Prédio Biomédicas II, Av. Lineu Prestes 1374, Sala 24, Cidade Universitária (05508-900) São Paulo (SP) Brazil; Tel: +55-11-3091-7335: Fax: +55-11-3091-7417: E-mail: [email protected] § Indicated authors contributed equally. 1381-6128/11 $58.00+.00

T. cruzi is a hemoflagellated protozoan parasite with a life cycle between invertebrate and vertebrate hosts, including several species of triatomine insects and more than 150 species of mammalian hosts, including humans (Fig. 1). The invertebrate insects become infected after a blood meal on mammals that host trypomastigotes (non-dividing, infective forms of the parasite) circulating in the blood. Most of these trypomastigotes die in the anterior region of the triatomine digestive tube, but a small proportion of this population reaches the insect midgut where they differentiate to epimastigotes (not infective, dividing form). Epimastigotes actively replicate and colonize the digestive tube from the medium to the distal region. In the distal region, epimastigotes differentiate into metacyclic trypomastigotes (infective non-dividing forms similar to

Fig. (1). Schematic representation of the T. cruzi life cycle. The cycle is complex and occurs between an invertebrate (insect vector) and a vertebrate (mammalian) host. The parasite differentiates among several stages inside each host, as shown. A: amastigote (intracellular dividing stage); Ei: intracellular epimastigote (intracellular dividing stage); T: trypomastigote (nondividing, infective extracellular forms in the mammalian host); E: epimastigote (dividing non-infective form in the midgut of the insect vector); M: metacyclic trypomastigote (non-dividing, infective form derived from epimastigotes in the insect vector). © 2011 Bentham Science Publishers

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those that occur inside the mammalian hosts) through a differentiation process known as metacyclogenesis. After feeding on a new mammal, the insect vector defecates and releases the metacyclic trypomastigotes in the feces, which can infect the new host via the mucosa or even through microscopic skin lesions. Once inside the mammalian host, these forms must invade the host cells by forming a transient parasitophorous vacuole, from, which it escapes to the cytoplasm. Once in the cytoplasm, trypomastigotes are able to differentiate into amastigotes (replicative and infective forms), which divide by binary fission [3, 4]. After replication, the amastigotes differentiate into trypomastigotes, passing transiently through an epimastigote-like (intracellular epimastigote) stage [5]. The trypomastigotes are released from the infected cells via cell lysis. Once outside the host cells, these forms can invade other cells in the infected tissue or enter the blood stream, thereby enabling them to infect other organs. While present in the blood stream, the trypomastigotes can be transmitted to a non-infected vector after the vector consumes a new blood meal. In summary, this parasite goes through several types of environments during its life cycle, and each environment has its own specific physical, physicochemical and chemical characteristics. WHY IS IT NECESSARY TO IDENTIFY NEW DRUG TARGETS TO COMBAT CHAGAS´ DISEASE? Some 40 years ago, two compounds were launched as therapeutic agents to treat T. cruzi infection in patients: benznidazole (Bz) and nifurtimox (NF) (Fig. 2). These compounds are the only currently available therapeutic weapons against Chagas’ disease, and since their approval for the use in humans, no other drugs have been developed that can treat T. cruzi infection. Bz is a nitroimidazole that seems to act mainly by interfering with the synthesis of macromolecules via covalent bonding between nitroreduction intermediates and cellular components, such as DNA, lipids and proteins. In addition, Bz acts by improving phagocytosis, stimulating the host immune response through the production of gamma-interferon (IFN), inhibiting the fumarate reductase, and interfering with the oxidative stress resistance mechanisms of the parasite. NF acts in the presence of oxygen through the generation of nitroanion radicals through nitroanion reductase to produce reactive oxygen species (ROS). However, these therapeutic drugs have serious drawbacks. Both Bz and NF are highly toxic and can cause secondary effects that in some cases force the interruption of treatment. In addition, both drugs are effective in the acute phase of the infection (when it is rarely diagnosed), and in congenital and reactivated infections, but their efficacy in the chronic phase is controversial, meaning that most of patients are not treated. These drawbacks highlight the need for new alternative chemotherapeutic treatments for Chagas´ disease. PROMISING TARGETS FOR THE DEVELOPMENT OF NOVEL DRUGS Despite the fact that Bz and NF are currently used to treat Chagas disease, they are not the ideal chemotherapeutics because of their side effects and their controversial performance in treating the chronic phase of infection. A detailed review of the new drugs with chemotherapeutic potential is beyond the scope of this review. However, some T. cruzi specific pathways involve proteins/enzymes that are currently being evaluated as targets, and several drugs that are known to interfere with these pathways are promising potential therapies [6]. A selected number of these targets are mentioned below and are shown in Table 1. THE CYSTEINE PROTEINASES The cysteine proteinases of T. cruzi participate in several cellular processes, such as energy metabolism, differentiation, host cell invasion and evasion of the immune system [7, 8]. The most abundant member of the cysteine proteinase family in T. cruzi is

Silva Paes et al. O N

O

N N

N H

O

N-benzyl-2-(2-nitro-1H-imidazol-1-yl)acetamide O S O

O N

N

O

N

O N-(3-methyl-1,1-dioxo-1,4-thiazinan-4-yl)-1-(5-nitro-2-furyl)methanimine

Fig. (2). Chemical structures of therapeutic drugs used against Chagas disease. Two drugs are currently used for the human treatment of Chagas disease. A. Benznidazole (N-benzyl-2-(2-nitro-1H-imidazol-1-yl)acetamide), B. Nifurtimox (N-(3-methyl-1,1-dioxo-1,4-thiazinan-4-yl)-1-(5-nitro2-furyl)methanimine).

cruzipain [9-11], which is found in a variety of isoforms with subcellular localizations [12]. Several studies have validated cysteine proteinases, more specifically cruzipain, as drug targets [7]. In particular, the use of synthetic inhibitors, such as vinyl sulfonederivatized dipeptides, showed promising results in treating the in vivo infection [13-16]. STEROL BIOSYNTHESIS PATHWAY The sterol biosynthesis pathway provides promising targets because ergosterol rather than cholesterol is the main sterol in T. cruzi membranes [17]. Therefore, specific inhibitors against enzymes specific for the ergosterol pathway will likely specifically target T. cruzi without harming mammalian cells. Several specific inhibitors, such as azole derivatives, directed against enzymes in this pathway (sterol C14 -demethylases [18-22], oxidosqualene cyclase (or lanosterol synthase) [23, 24] and 2D-squalene synthase [25, 26] were tested in epimastigote cultures, in in vitro or in vivo infections [27]. BIOSYNTHESIS OF POLYISOPRENOIDS Enzymes of the polyisoprenoid biosynthesis pathway, such as farnesylpyrophosphate synthase [28-30] and protein farnesyltransferase [31], were also shown to be targets of various inhibitors. Among them, promising results were obtained with biphosphonates and their derivatives, which proved to be potent inhibitors of farnesylpyrophosphate synthase [28]. They were also effective in cell culture, and decreased the parasitemia in infected mice [32-34]. Biphosphonates also accumulate in acidocalcisomes, which are important for Ca2+ metabolism in the parasite, suggesting that these organelles are also a relevant target for these compounds [35]. Interestingly, they are largely used for the treatment of calcium resorption disorders in human bones. These facts make these drugs good candidates for the treatment of the clinical phases of T. cruzi infections. Among the inhibitors of protein farnesyltransferase, Rphenylalanine and N-propylpyperazinil derivatives were selective in in vitro assays and extended the survival of infected animals [31]. PURINE SALVAGE PATHWAYS AND NUCLEOTIDE SYNTHESIS Enzymes in these metabolic pathways are essential for trypanosomatids because these parasites are auxotrophic for purine and are strongly dependant on purine uptake and salvage [36, 37]. The proteins involved in this metabolic pathway that have been evaluated as drug targets include purine (hypoxanthine/guanine)-

The Uniqueness of the Trypanosoma cruzi Mitochondrion

Table 1.

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Summary of the Druggability of Enzymatic Pathways in T. cruzi Metabolism

Enzymes

Energy metabolism, differentiation, host cell invasion and evasion of the immune system

Cystein-proteinases (cruzipain)

Sterol biosynthesis pathway

Sterol C14-demetylases Oxidosqualene cyclase Lanosterol synthase

Localisation

Drug

Ref.

Vinyl sulphone derivatized dipeptides

[13-16]

Azole derivatives

[18-26]

Acidocalcisomes

Derivatives of biphosphonates R-phenylalanine and Npropylpyperazinil

[28-31]

Cytosol

Allopurinal

[38-43]

Nucleus, Mitochondrion

Coumermycin A1 Clorobiocin

[46, 124, 125, 119]

Kinetoplast

Berenil Hydroxystilbamidine

Several subcellular localisations

2D-squalene synthase Biosynthesis of polyisoprenoids

Farnesylpyrophosphate synthase Protein-farnesyl transferase

Purine salvage pathways and nucleotide synthesis

Purine phosphoribosyl-transferase

Topoisomerases

Topoisomerase I Topoisomerase II

Dihydrofolate reductase Pteridine reductase Dihydroorotate dehydrogenase

Camptothecin Rebeccamycin RNA editing proteins

Endonuclease

Mitochondrion

RNA ligase KREL1

GW5074, mitoxantrone

[89, 90]

NF 023, protoporphyrin IX D-sphingosine Naphthalene-based scaffolds

Oxidative stress

Trypanothione reductase

Mitochondrion

Organo-metallic complexes

[354-362]

Tricyclic compounds polyamine

phosphoribosyltransferase [38-40], dihydrofolate reductase [41, 42], pteridine reductase, and dihydroorotate dehydrogenase [43]. In addition, allopurinol, which is an inhibitor of purine (hypoxanthine/guanine)-phosphoribosyltransferase, was proposed for the treatment of the reactivation of T. cruzi infection in patients after heart transplantation [44, 45]. TOPOISOMERASES AND DNA Enzymes and proteins related to DNA replication have also been targeted for selective inhibition. In addition to enzymes involved in kDNA replication, which will be discussed below, enzymes such as topoisomerase I [46], which is involved in nuclear DNA replication, have proven to be potential targets. Inhibitors of these enzymes were found to efficiently block T. cruzi growth. Parasite DNA was also proposed as a target for intercalators and binders that showed trypanocide activity [47-49]. THE UNIQUENESS OF SOME KINETOPLASTID ORGANELLES The Glycosome and the Reservosome T. cruzi, similar to other pathogenic trypanosomatids, possesses several peculiar characteristics that, most of which are beyond the scope of this review. A brief overview will provide a better understanding of the uniqueness of the organism’s “cell design”; in particular, there are two specific organelles: the glycosome and the reservosome. Glycosomes are modified peroxisomes, which contain some enzymes of the glycolytic pathway (in T. cruzi, the first seven glycolytic enzymes participate in the conversion of glucose into glyceraldehyde 1,3-bisphosphate) and enzymes of the pentose

phosphate pathway [50, 51]. Glycolysis in trypanosomatids also has some special characteristics in addition to its compartmentalization between the glycosome and the cytosol, including the lack of negative regulation in the presence of oxygen (Pasteur effect) and the lack of the allosteric regulation of hexokinase and phosphofructokinase [52]. Some authors have proposed that compartmentalization and the lack of regulation of these enzymes are related phenomena because the glycolytic flux in these organisms could be controlled at the level of glucose transport into the glycosome and the intraglycosomal balance of ADP/ATP [53]. Reservosomes are lysosome-related organelles, which are the main compartments for the storage of ingested proteins and lipids, as well as proteolytic enzymes originating from the secretory pathway of the protozoan that are used as energy sources (these parasites do not have polysaccharides as energy reserve) [54-56]. The reservosomes are conspicuously present in epimastigotes, but they are not found in other stages, suggesting that their content (proteins) is degraded and consumed during metacyclogenesis. This fact, together with early works showing that these parasites actively oxidize several amino acids, led several researchers to focus on the degradation of amino acids as an energy source. As occurs in other protists, T. cruzi has acidocalcisomes, which are acidic organelles that resemble lysosome-related organelles (LROs) from mammalian cells and store polyphosphates and pyrophosphates complexed with Ca2+. It has been proposed that acidocalcisomes are involved in the storage of cations and phosphorus, phosphorus metabolism, the maintenance of Ca2+ and intracellular pH homeostasis, and osmoregulation [57, 58]. As is the case for the previously described organelles, T. cruzi (as well as other kinetoplastid) single mitochondrion presents its


unique characteristics. The aim of the present review is to describe these peculiarities, and discuss their use as chemotherapeutic targets. THE UNIQUE MITOCHONDRIA OF KINETOPLASTIDS Some General Characteristics Flagellates of the order Kinetoplastidia bear a single and usually well developed mitochondrion that spans the entire cell body and is distributed in branches under the subpellicular microtubules. This mitochondrion is comprised of the classical compartments: the outer membrane, the intermembrane space, the inner mitochondrial membrane that forms infoldings known as mitochondrial cristae, and the mitochondrial matrix. Depending on the available environmental and nutritional resources, the mitochondrion can occupy a variable amount of the total cellular volume [59]. The mitochondrial DNA of trypanosomatids is localized in a certain portion of the organelle, near the basal body, enclosed in an unusual structure known as the kinetoplast (kDNA). More precisely, the kDNA is located within the mitochondrial matrix, perpendicular to the axis of the flagellum (reviews in [60] and [61]). Regarding the electron transport machinery, the presence of complex II to V has been demonstrated. However, as will be addressed later in more detail, there is controversy with respect to the presence and contribution of complex I (NADH:ubiquinone oxidoreductase) to energy metabolism [62]. Structure and Organization of the Mitochondrial Genome (kDNA) kDNA is the most structurally complex mitochondrial DNA in nature. The morphology of the kinetoplast varies depending on the developmental stage of the parasite. In T. cruzi epimastigotes and amastigotes, the kinetoplast forms a bar-like structure. In contrast, trypomastigotes have a roundish kinetoplast [63, 64]. Unlike any other DNA in nature, the kDNA of trypanosomatids is composed of circular molecules that are topologically relaxed and interlocked to form a single network. Two types of DNA rings, the minicircles (0.65-2.5 kb) and the maxicircles (20-40 kb in size depending on the species), are present in the kinetoplast. Approximately 50 units of maxicircle DNA are found in each organelle. Minicircles are found in the range of 5,000-10,000 units per organelle, representing more than 90% of the network in terms of mass. Maxicircles are the functional homologs of mitochondrial DNA in other eukaryotes and encode ribosomal RNAs and several proteins, most of which are involved in mitochondrial energy transduction [65]. Some genes in maxicircles have a very unusual structure: these genes have incomplete or immature open reading frames and have to be remodeled by a complex process called RNA editing in order to convert them into translatable transcripts [66]. All of the tRNAs necessary for translation are imported from the cytosol [67]. Mitochondrial Gene Expression RNA Editing It was apparent from the sequence data of the maxicircles of several kinetoplastids that the maxicircles contain genes without proper open reading frames (ORFs) and genes that lack start or stop codons [68]. The analysis of this unconventional gene organization led to the discovery of a process known as RNA editing, which generates correct ORFs through the addition or removal of uridine residues in specific sites of the transcripts [66]. This complex process is conducted by trans-acting RNA molecules called guide RNAs (gRNAs) (Fig. 3). The gRNAs are encoded by the minicircles and have a characteristic primary structure. They have a 5’ region, or anchor, and a complementary region (10 bases) that hybridizes to the RNA to be edited at its 3’ position, counting from the first unpaired nucleotide. This latter region provides the landmark to label the editing site. The central portion of the gRNA contains the sequence information

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necessary for editing and is complementary to the sequence that will become the mature mRNA. The transfer of information between the gRNA and mRNA is based on both Watson-Crick interactions and an unconventional G:U base pairing [69]. Finally, the 3’ region consists of a poly-uracil (poly(U)) tail, which is posttranscriptionally added to the molecule. This sequence stabilizes the interaction between the gRNA and the mRNA by annealing with purine-rich regions present in the pre-edited transcript [70]. The mechanism of the addition/deletion of uridines involves several steps of an “enzymatic cascade” [71, 72] (Fig. 3). First, the gRNA hybridizes downstream of the first editing site, via the anchor region, forming the gRNA-mRNA duplex. Subsequently, an endonuclease cleaves the site where the mismatch occurred. In the case of the addition of uridine, a terminal uridyltransferase (TUTase) transfers U residues to the 3’ end of the 5’ fragment; these U residues are complementary to the nucleotides in the gRNA. Free UTP molecules are used as the uridine source for this process. Subsequently, a 3’ to 5’ exonuclease removes the extra unpaired U residues and finally, a ligase links the processed fragments. When the editing involves the removal of Us, a U-specific exonuclease (Exo-Uase) removes the residues. This cycle of cleavage, addition/deletion and ligation can continue in an upstream editing site along the primary transcript. A helicase also seems to be involved in this complex process [for further review on this topic see [73]]. Editing Machinery The protein machinery responsible for the editing process involves different enzymatic activities, which have not been fully identified and characterized. Several research groups have described a protein complex that sediments at 20 Svedberg (20S) in mitochondrial lysates and was named the “20S editosome”. This complex is involved in the key activities needed for the editing process, including an endonuclease activity that cleaves the mRNA at the editing site, 3’ to 5’ exonuclease (exo-Uase) activity for terminal deletions, uridylyl transferase (TUTase) activity for the insertion of Us and ligase activity that joins the resulting products (reviewed in [74, 75]). The 20S editosome was mainly described for T. brucei and L. tarentolae by a variety of different groups, which led to confusing nomenclature for each of the protein factors. In this review will use the nomenclature introduced by Stuart et al. (2005) [74]. With respect to the other trypanosomatids, genes encoding proteins that are orthologous to those already described in the editing process in T. brucei and L. tarentolae were identified in genomic databases of T. cruzi and Leishmania spp [76-78]. A description of the enzymatic activities involved in the RNA editing process in trypanosomatids is included below. - Endonucleases: In spite of the fact that proteins with endonuclease motifs have been found among the components of the 20S editosome, endonuclease activities were minimally described. The KREBP1-5 proteins were identified as type III RNAses, which contain a motif that can confer endonuclease activity. Moreover, these factors have an N-terminal U1-like zinc finger domain, which could facilitate their interaction with RNA or other proteins. In addition, other RNA binding motifs were also identified; for KREBP1-3, they have a dsRNA binding domain, whereas in the case of KREBP 4 and 5, they have a Pumilio-like domain, which recognizes specific sequences in the mRNA [76, 79, 80]. These latter proteins are recognized as probable structural factors involved in the stabilization of the complex rather than in any exonuclease activity. Three others proteins with U1-like zinc finger domains (KREPB6-8) with no RNAse III-like domains were identified, so they are not considered potential exonucleases [74, 76]. - Exonucleases: The 3’ to 5’ exonuclease (exo-Uase) activity is responsible for the elimination of U residues and is required



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Fig. (3). Schematic representation of the RNA editing process. The editing process is complex, consisting of the addition or deletion of uridines in the primary transcript. ES: editing site, TUTase: terminal uridyltransferase, Exo-Uase: U-specific exonuclease.

-

-

for the editing process. The proteins identified with this activity are KREPC1 and KREPC2. The latter was purified with other factors involved in deletion editing [81] and has a Cterminal EEP motif with endo- and exonuclease activities [76]. KREPC1 has a similar structure as KREPC2, especially in relation to its EEP motif, and likely also acts as a 3’ to 5’ exonuclease [79]. The presence of two domains in this proteins suggests that it could be constituted by multifunctional factors, that could participate in the catalysis of different substrates or could be differentially expressed, being each one characteristic of different life cycle stages [74]. Terminal uridylyl transferase (TUTase): This activity is essential for the addition of Us during RNA editing. The activity was described in two protein factors: KRET1 [82] and KRET2 [83]. Although these proteins have nucleotidyl transferase and poly(A) polymerase activity, it was shown that they have different roles. KRET2 is part of the insertion subcomplex and is the TUTase responsible for gRNA-dependent U addition in the transcript that is being edited [84]. In contrast, KRET1 has the ability to add Us in a gRNA independent manner [83, 84] and the absence of KRET1 leads to gRNAs with shorter poly(U) regions [82, 84]. These results indicate that the KRET1 TUTase is responsible for adding Us to the poly(U) tail in the gRNA. In addition, KRET1 has a zinc-finger domain that is absent in KRET2, which could be important for its function [83]. RNA Ligases: Two proteins were identified as RNA ligases in the 20S editosome complex: KREL1 and KREL2 [85]. Using techniques, such as tandem-affinity purification (TAP), yeast two-hybrid analyses and co-immunoprecipitation, two subcomplexes have been identified. One of these subcomplexes contains KREL1, which can perform the exo-Uase and ligation steps of RNA editing by deletion in vitro. The other subcomplex contains KREL2, which can insert U residues and ligate during RNA editing [74, 81].

RNA helicase: Due to the need to recycle the gRNAs for different rounds of editing, the presence of a helicase activity in the editosome complex was suggested, which could facilitate the opening of the gRNA:mRNA duplex. Using biochemical techniques and immunoaffinity purification, it was possible to isolate an editosomal KREH1 helicase activity, which contains a DEAD box helicase domain that was shown to be nonessential for parasite growth [86]. However, it was not possible to isolate the activity associated with the 20S editosome by using the TAP-tagging technique, suggesting that this factor is not a stable component of the complex [79]. Additional studies are necessary to establish the precise functions of this activity. - Additional factors: Other 20S editosomal proteins were identified, but their functions are still unclear. The factors KREPA16 have RNA and protein interaction domains [76, 81]. It was demonstrated that the absence of KREPA1 and KREPA2 alters editosome formation and activity, suggesting a role for these proteins in maintaining the integrity of the 20S editosome. In addition to the aforementioned factors, accessory proteins involved in RNA editing, such as MRP1 and MRP2 (RNA binding proteins that form a heterodimeric complex), were also identified. Three other mitochondrial proteins, RBP16, REAP1 (RNA editing-associated protein 1) and TbRGG1, also possess the ability to bind RNA. Together with those factors mentioned above, they may play roles in both RNA editing and regulation and in other processes involving mitochondrial RNAs (e.g., the maturation of polycistronic pre-mRNAs). For details, see [74, 87]. From all of the information obtained regarding the components and structure of the 20S editosome complex, it was proposed that this complex can be divided into two discrete subcomplexes, in which the functions of U insertion and deletion are physically and functionally separated. Inside each of these subcomplexes, pairs of related proteins perform similar functions, which consist mainly of insertional or deletional editing [88]. In addition to the 20S edito-


some, a larger complex that sediments at approximately 40S was described for T. brucei [89, 90]. This finding implies the presence of other molecules and protein factors that are required for RNA editing in addition to the essential activities mentioned above, which are part of the enzymatic “core” [75]. RNA Editing is Differentially Regulated In T. brucei, an RNA editing process has been described that is differentially regulated throughout the life cycle. This phenomenon suggests that RNA editing may contribute to the regulation of parasite gene expression, and more specifically, with respect to the changes in energy metabolism that are experienced by the parasite. For example, in T. brucei, mRNAs encoding the cytochrome system are mainly edited in the procyclic forms, whereas those for the NADH dehydrogenase complex are edited in the bloodstream form of the parasite. This differential regulation corresponds to the energy-generating processes for each of the stages: the bloodstream forms are mainly dependent on glycolysis as an energy source, whereas the procyclic forms obtain their energy mainly from cytochrome-mediated oxidative phosphorylation [88, 91]. The mechanism by which this differential editing is regulated is still unknown. Riley and colleagues showed that this regulation is not dependent on the number of gRNAs, and their abundance remains constant throughout the life cycle of T. brucei [92, 93]. It has been proposed that RNA editing may be controlled at the level of gRNA utilization, most likely involving the protein factors mentioned above [94]. This differential RNA editing observed in T. brucei is less studied in other trypanosomatids, such as T. cruzi, and Leishmania spp. Kim et al. compared the differential expression of the gene encoding subunit II of cytochrome oxidase (COII). In contrast to T. brucei, no differences were observed in the COII mRNA levels in both insect and mammalian stages of T. cruzi [95]. Because T. cruzi (as well as T. brucei) goes through a complex life cycle and alternates between different energy sources, the contribution of the RNA editing process to metabolism cannot be ruled out. Presently, this possibility should be considered as a hypothesis, and additional studies are needed for confirmation. RNA Editing Proteins as Therapeutic Targets Mitochondrial RNA editing is a vital and unique process, which occurs in the mitochondria of microorganisms, in general, and predominantly trypanosomatids. This specificity makes RNA editing a potential target for new antiparasitic drugs. In this context, Liang and Connell used high-throughput screening to identify specific inhibitors of RNA editing. Five compounds were identified in this screen: GW5074, mitoxantrone, NF 023, protoporphyrin IX, and D-sphingosine, which proved to be inhibitors of insertional editing. These compounds had IC50 values between 1 and 3 M (Table 1). More specifically, GW5074 and protoporphyrin IX inhibited the editing process at the level of endonuclease cleavage, which begins the editing process [96]. Another potential target in the RNA editing process is the RNA ligase KREL1, as proposed by Durrant et al. In a recent paper, this group showed four inhibitors with unique naphthalene-based scaffolds, which could act as inhibitors of the KREL1activity in T. brucei [97] (Table 1). The Replication of kDNA Virtually all the information on kDNA replication were derived from studies conducted primarily in C. fasciculata and T. brucei. kDNA replication involves not only the duplication of the two populations of DNA molecules (mini- and maxicircles) but also the proper segregation of all of the kDNA pieces of the replicated genomes between both daughter mitochondria. The improper segregation of a mini- or a maxicircle could result in the loss of essential information for one of the daughter cells, so that kinetoplastids have developed a sophisticated system to ensure the proper segregation of the replicated kDNA.

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Duplication of kDNA occurs in a specific period during the cell cycle, close to S phase [98]. This differs from the asynchronicity of mDNA duplication in mammals, which occurs randomly throughout the life cycle [99]. As mentioned above, the kDNA network is condensed and highly ordered in a disk-shaped structure. This disk is always positioned close to the flagellum basal body and perpendicular to it (reviews in [60] and [61]). There is evidence of direct physical contact between these structures, despite being separated by the double mitochondrial membrane [100]. The area of the mitochondrial matrix between the kDNA and the basal body is called the kinetoflagellar zone (KFZ) [101]. Replication Mechanism The proteins necessary for initiating the replication of the kDNA are located in the kinetoflagellar zone. The replication mechanisms of the mini- and maxicircles are similar to one another; both are replicated unidirectionally through type intermediates. The main difference between these two mechanisms is that maxicircles are duplicated while they are integrated into the network, wheras minicircles need to be liberated from the network in order to be replicated as free molecules [102, 103]. At the beginning of the replication process, individual minicircles are released vectorially from the network to the KFZ by a topoisomerase II activity [101]. Replication begins at conserved sites that are recognized by the universal minicircle sequence binding protein (UMSBP). This protein was identified in T. cruzi with two highly polymorphic and differentially expressed alleles at the TcUMSBP locus [104, 105]. Several proteins appear to bind to the replication origin, including a primase, which is responsible for the synthesis of RNA primers, and replicative DNA polymerases (probably DNA polymerases IB and IC, two of the six mitochondrial DNA polymerases present in trypanosomes) [106]. The replication initiation complex is formed with these and probably other proteins, thus enabling the initiation of DNA synthesis [107]. Flanking the kinetoplast, and separated by 180°, are recognized structures named as antipodal sites. These structures are constituted by protein complexes that contain the activities that are necessary for the next steps of replication, such as the removal of RNA primers by an RNAse H called SSE1 [108], the sealing of gaps between Okazaki fragments by a type DNA polymerase [109] and the closing of the resulting nicks by a specific ligase [110, 111]. Once the minicircles have been duplicated, the replication mechanism in T. cruzi differs from that of T. brucei because the distribution of duplicated molecules changes from one organism to another. In the case of T. cruzi, C. fasciculata and Leishmania tarentolae, replicated minicircles are uniformly re-linked around the network periphery, ~180° apart, forming a ring that grows in thickness as the replication advances. This distribution results from the relative motion of the kDNA disk and the two antipodal sites [112, 113]. In contrast, the minicircles replicated in T. brucei are re-linked to the network with a polar distribution, together with the antipodal sites [114, 115]. It was recently demonstrated using fluorescence methods that the kinetoplasts in C. fasciculata and T. brucei move relative to the fixed antipodal sites, presenting a rotating and oscillating motion, respectively, which contributes to the distribution previously described [116]. The duplicated minicircles still contain at least one gap, which serves as a “landmark” to tag the replicated minicircles and ensure that each one has been duplicated once and only once [107]. Minicircles are subsequently re-attached to the network by a topoisomerase II activity [117] and must then be split in two for distribution to daughter cells. That division also involves a topoisomerase II activity and probably other still unknown factors that have their activities restricted to diameter of the network. This process would generate progeny kDNA of identical size and content [118]. Finally, the two newly formed networks must be segregated to daughter


cells during cytokinesis. This process is achieved through molecular connections between the basal bodies of the flagella and the kDNA networks that are being segregated [100]. The identities of the molecules that bind both structures are unknown, but a filament system called the tripartite attachment complex (TAC) has been described. This structure goes through the mitochondrial double membrane and it may be responsible for kinetoplast positioning and segregation [119]. DNA Topoisomerases as Drug Targets in T. cruzi The DNA topoisomerases are a potential drug target in trypanosomatids because of their role in kDNA replication and due to their uniqueness compared to the corresponding enzymes present in the host. Several topoisomerases have been characterized in several trypanosomatids [46, 120-128]. With regard to T. cruzi, Riou and colleagues described a topoisomerase I with ATP independent activity [129]. Another work showed that T. brucei topoisomerase I is comprised of two subunits encoded by different genes, forming a heterodimeric enzyme [128]. This unusual structure is likely shared by other trypanosomatids, as suggested by information in genome databases and other recent studies [130]. Regarding topoisomerase II activity, Fragoso et al. have isolated and sequenced the gene encoding this enzyme [131]. In a later study, the enzyme was characterized and was shown to be expressed to a greater extent in the replicative forms of the parasite [127]. There are two classes of drugs that target the topoisomerase enzymes: the “topoisomerase inhibitors”, which compete with ATP for binding to the catalytic site by interfering with their function; and the “topoisomerase poisons”, which stabilize the DNA-enzyme complex, resulting in the breakdown of DNA (reviewed by [132] and [59]). Many of these drugs have been used in the treatment of cancer and bacterial infections, and to a lesser extent, their effects have been evaluated in trypanosomatids. Douc-Rasy and colleagues studied the effect of these drugs on the proliferation of T. cruzi and described a Topo II with catenation and decatenation activity that could be inhibited by coumermycin A1 and clorobiocin and by the trypanocides, berenil and hydroxystilbamidine [126] (Table 1). In addition, intercalating drugs, such as ellipticine derivates, were also potent inhibitors of not only Topo II activity but also the reactions catalyzed by Topo I [133]. This same activity was found to be sensitive to the drug camptothecin, which is an antitumor drug and a eukaryotic topoisomerase II inhibitor. This drug was shown to be cytotoxic to T. cruzi, T. brucei and L. donovani [46]. In another work, Gonzales-Perdomo et al. showed that bacterial topoisomerase II inhibitors, such as ofloxacin, nalidixic acid and novobiocin inhibit the transformation of epimastigotes into metacyclic trypomastigotes or amastigotes into trypomastigotes in a dose-dependent manner. Because these processes involve replication or changes in the kDNA, the results suggest that topoisomerase II could be a target of these drugs and may play a key role in the proliferation and differentiation of T. cruzi [134]. In a recent study performed by Zuma et al., the effects of different topoisomerase inhibitors and DNA binding drugs on the proliferation and ultrastructure of T. cruzi epimastigotes were evaluated. The results showed that eukaryotic type I topoisomerase inhibitors, such as the abovementioned camptothecin and rebeccamycin, were the most effective in arresting the proliferation of T. cruzi. With regard to eukaryotic type II topoisomerase inhibitors, mitoxantrone was the most effective (Table 1). As previously shown, these compounds can block cellular proliferation and promote ultrastructural alterations of the kinetoplast [135]. With respect to the DNA binding drugs, it was observed that only berenil significantly disrupted kDNA [136]. All results presented demonstrate the importance of DNA topoisomerases, which are essential enzymes that demonstrate a high potential for the development of new drugs against trypanosomatids.


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Transcription and Protein Synthesis As occurs in the trypanosomatid nucleus, transcription in the mitochondria is polycistronic for both mini- and maxicircles. The polycistronic RNA is processed and matured to form monocistronic RNAs through precise cutting events, polyadenylation, and in some cases, editing [87]. The polyadenylation mechanism in the mitochondria of kinetoplastids is not well understood. In this complex process, poly(A) length is partially dependent on RNA editing status. In addition, differences in poly(A) lengths may be related to the life cycle stage [137-139]. In most organisms, only some tRNAs must be imported from the cytosol to conduct protein synthesis. Interestingly, trypanosomatids must import the complete set of tRNAs for the translation of mitochondrial proteins because none of them are encoded by the mitochondrial genome [67, 140]. This requirement results in a paradox: mitochondrial translation in trypanosomatids is, from the evolutionary point of view, similar to the prokaryotic method of translation; however, only eukaryotic tRNAs are imported from the cytosol [141]. T. cruzi Mitochondrial Metabolism: a general View The mitochondrial metabolism of T. cruzi involves a variety of enzymes and proteins, which are connected to many metabolic processes. While the bioenergetic role of mitochondria is linked to the electron transport chain and enzymes for oxidative phosphorylation, there are other important biosynthetic activities that take place in mitochondria [142]. To date, energy metabolism has been more deeply studied in T. brucei than other trypanosomatids [143]. However, after of the completion of the genome sequence of T. cruzi [77], as well as that of the related kinetoplastids T. brucei and L. major [78, 144] it has been accepted that energy metabolism is similar in most trypanosomatids. However, few but significant differences may exist between these different organisms. In addition, the integration of proteomic and genomic data with the analysis of the primary biochemical literature has extended our understanding of T. cruzi metabolism (for a summary of this topic, see Fig. 4) [145-149]. Acetyl-CoA - an Important Mitochondrial Intermediate The successful survival of the parasite in different environments throughout its life cycle (i.e, the vector intestinal tract, the mammalian host blood stream and the host cell cytoplasm) depends on its ability to maintain the intracellular homeostasis of ions and nutrients and its ability to catabolize different substrates for energy [150]. It is well established that trypanosomatids are able to uptake and consume both glucose and amino acids as their main carbon and energy sources [52, 150, 151]. As previously mentioned, the major portion of the glycolytic pathway is compartmentalized in glycosomes in all trypanosomatids (Trypanosoma and Leishmania species) studied so far [152, 153]. Glucose is partially oxidized to form end products such as succinate, L-alanine, acetate, lactate, pyruvate, glycerol and CO 2 [143, 151, 154], although potential pathways for oxidative phosphorylation have not been found [155]. Whereas the first seven steps of glycolysis (from glucose to glycerate 1,3-bisphosphate) occur inside the glycosomes, the conversion of glycerate-2phosphate to phosphoenolpyruvate (PEP) occurs in the cytosol. Cytosolic pyruvate is a branch point for two possible pathways: 1. The pyruvate produced in the cytosol from PEP enters into the mitochondrion and is converted to acetate via acetyl-CoA [156158]. 2. Alternatively, the PEP can re-enter the glycosomes to be carboxylated to oxaloacetate by phosphoenolpyruvate carboxykinase (PEPCK). The resulting oxaloacetate is converted to malate by


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Fig. (4). Schematic representation of the amino acid metabolic pathways in the mitochondria of T. cruzi. Solid lines represent pathways or reactions that were biochemically demonstrated. Blue boxes represent excreted end products (acetate, L-alanine, succinate and CO2 ). Pink boxes represent the amino acids involved in mitochondrial metabolism in T. cruzi. Dotted lines represent putative pathways, reactions inferred from the T. cruzi genome project or those that are still being studied. AOB: amino oxobutyrate; P5C: pyrroline-5-carboxylate. The enzymes are numbered as follows: 1- aspartate aminotransferase, 2mitochondrial malic enzyme, 3- pyruvate dehydrogenase complex, 4- acetate: succinate CoA-transferase, 5- succinyl-CoA synthetase, 6-unknown enzyme, 7acetyl-L-carnitine transferase, 8- acetyl-CoA: glycine C-acetyltransferase, 9- L-threonine dehydrogenase, 10- acetyl-CoA-synthetase, 11- aconitase, 12cytosolic NADP-linked isocitrate dehydrogenase,13- mitochondrial NADP-linked isocitrate dehydrogenase, 14- 2-ketoglutarate dehydrogenase, 15-succinylCoA synthase, 16-mitochondrial NADH-dependent fumarate reductase, 17-succinate dehydrogenase, 18- mitochondrial fumarate hydratase, 19- mitochondrial malate dehydrogenase, 20-citrate synthase, 21-alanine racemase, 22-alanine aminotransferase, 23- glutamate dehydrogenase, 24- glutamine synthetase, 25aspartate aminotransferase, 26-formiminoglutamase, 27-asparagine synthetase A, 28- L-asparaginase, 29- pyrroline-5-carboxylate dehydrogenase, 30- Lproline dehydrogenase, 31- proline racemase, 32- arginine kinase, 33- nitric oxide synthase, 34-methionine adenosyltransferase, 35-spermidine synthase, 36cystathionine -synthase, 37-serine hydroxymethyltransferase.

glycosomal malate dehydrogenase. As a branching point of this pathway, malate can either leave the glycosome and be converted into pyruvate again by a cytoplasmic malic enzyme or it can enter the mitochondrion where it can be converted again into pyruvate by the mitochondrial malic enzyme. Inside the mitochondrion, pyruvate can be coupled to coenzyme A (CoA) to form acetyl-CoA (catalyzed by the pyruvate dehydrogenase complex (PDH)) and then liberated as acetate. The malate can also be directly oxidized in the tricarboxylic acid cycle (TCA), forming succinate [150, 156, 159-161]. Recent evidence showed that in T. cruzi, as in T. brucei, the Krebs cycle has two main branches, one involved in the generation of citrate via the condensation of oxaloacetate and acetyl-CoA by the pyruvate dehydrogenase complex and a second, which is associated with the conversion of

the -ketoglutarate derived from amino acid oxidation (L-aspartate and L-glutamate) into succinate [162, 163]. Acetyl-CoA is a key intermediary metabolite at a crossroad between the catabolic and anabolic pathways of carbohydrates, amino acids and fatty acids. For several years, it was believed that in most trypanosomatids acetyl-CoA produced during metabolism is converted into CO2 and acetate through the TCA [154]. However, some studies concluded that most of the acetyl-CoA is converted into the excreted acetate [154, 156, 164, 165]. Acetate is produced by a two-step cycle. First, the acetate:succinate CoA transferase (ASCT) transfers the CoA moiety of acetyl-CoA to succinate, yielding acetate and succinyl-CoA. Then, CoA is recycled by the subsequent liberation of succinate by succinyl-CoA synthetase (SCS), an enzyme that generates extra ATP [154, 158, 166].


In trypanosomatids, the conversion of the amino acid Lthreonine into equimolar amounts of glycine and acetate is considered to be the main source of acetyl-CoA for lipid biosynthesis [156, 167, 168]. In most organisms, the degradation of ketogenic amino acids, such as leucine, isoleucine and lysine, directly results in the formation of acetyl-CoA (not from the pyruvate generated in glycolysis) in some organisms. Results obtained by Ginger et al. provided some evidence that T. cruzi epimastigotes are able to incorporate leucine directly into the sterol biosynthetic pathway when there is a need for a carbon source alternative to acetyl-CoA [169]. Because the production of acetyl-CoA occurs inside the mitochondrion and the biosynthesis of fatty acids takes place in cytosol, acetyl-CoA must be transferred to the cytoplasm [165]. It is widely accepted that all eukaryotes analyzed so far use the citrate/malate shuttle to transfer acetyl group equivalents between the two compartments [170]. In trypanosomes, it was assumed that the existence of a citrate/malate shuttle is necessary to supply the raw materials for cytosolic fatty acid biosynthesis [165]. In this shuttle, intramitochondrial acetyl-CoA first reacts with oxaloacetate to form citrate in the TCA cycle reaction catalyzed by citrate synthase (CS). Citrate passes through the mitochondrial inner membrane on a citrate transporter or a citrate/malate exchanger and is converted back to acetyl-CoA and oxaloacetate by the cytosolic citrate lyase [156, 165, 170]. Although the genome of T. cruzi possesses genes encoding a putative citrate synthase, the cytosolic citrate lyase gene is missing and the cell-free extracts of T. cruzi epimastigotes do not exhibit ATP-dependent citrate lyase activity [162] as reported in other trypanosomatids [165, 170]. Experimental data suggest the existence of other exchange systems that feed anabolic pathways and lipid biosynthesis in T. brucei (T. cruzi may also use these alternative pathways). For instance, the L-carnitine acetyl-transferase (CAT) mitochondrial system, which is able to transfer the acetyl group of the acetate to acyl groups, and the cytosolic L-carnitine acetyl-transferase, which converts acetate into acetyl-CoA [154, 171-173] or converts acetyl-CoA to acetate in the mitochondrion. The acetate can then cross the mitochondrial inner membrane (by passive diffusion or through an acetate carrier) to be converted into acetyl-CoA by the cytosolic acetyl-CoA synthetase (AceCS) [170]. L-amino Acid Metabolism in T. cruzi Mitochondria Early studies showed that epimastigotes of T. cruzi are able to take up and consume glucose first and continue with amino acids degradation [151]. T. cruzi obtains glucose from the extracellular medium through a single transport system [174, 175] which is regulated throughout the parasite’s life cycle [176]. On the other hand, the considerable pool of free intracellular amino acids can be obtained through their transport from the extracellular medium, from their biosynthesis from metabolic precursors and/or the degradation of proteins [150, 151]. It is worth noting the multiple roles that amino acids and their metabolites have in T. cruzi beyond their participation in protein synthesis and energy metabolism. In T. cruzi, amino acids and some derived metabolites are also involved in osmoregulation [177], polyamine biosynthesis [178, 179], stress resistance [180-183], energetic state management [184, 185], differentiation [186-188] and intercellular communication through the generation of NO [189]. In view of all these functions, efforts to unveil the relevance of amino acids and their metabolism during the T. cruzi life cycle are currently underway. In terms of metabolism, the relevance of several amino acids was established early on. It has long been known that, in part, the Krebs cycle operates based on the conversion of histidine into glutamate and, subsequently, into -ketoglutarate in the insect stage of T.cruzi [145]. It has also been shown that glutamine can be synthesized from glutamate and NH3 in the presence of glutamine


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synthetase [181, 190] (T. cruzi genome initiative of the Sanger Center systematic names Tc00.1047053508175.370 and Tc00.1047053503405.10). In turn, some studies support the idea that glutamine might be oxidized through the glutamate pathway. However, no genes for glutamate synthase or glutaminase have yet been found [150]. Other classic works showed that several different amino acids, such as asparagine, aspartate, glutamate, glycine, leucine, isoleucine and proline, can be metabolized by the parasite [191-193]. All of these amino acids also seem to feed the TCA cycle through their conversion into glutamate or aspartate [150]. In T. cruzi, amino acids are actively metabolized in the mitochondrion, generating the precursors needed for energy production as well as the intermediates required for metabolic processes that occur in the cytosol [194]. Amino acid catabolism in T. cruzi epimastigotes requires the elimination of the -NH2 group to generate the corresponding metabolizable ketoacid. As these parasites lack a functional urea cycle, the excess nitrogen is released as NH3 [195]. However, these parasites are equipped with an enzymatic transamination network for the exchange and disposal of the -NH2, allowing the transfer of these groups from different donor species (amino acids) to acceptor species (keto acids, mainly -ketoglutarate, oxaloacetate and pyruvate), and dehydrogenases, which can transfer the -NH2 to H2O, generating NH3 [196]. The dehydrogenases have many functions, such as the detoxification of NH3 through its transfer to acceptor ketoacids, such as ketoglutarate, again forming glutamate [197]. In turn, the aminotransferases, in particular, tyrosine aminotransferase (TAT), alanine aminotransferase (ALAT), aspartate aminotransferase (ASAT) and L--hydroxyacid dehydrogenase (AHADH), are responsible for the transfer of the -NH2 groups of almost all amino acids to pyruvate, yielding alanine and releasing the corresponding ketoacids (e.g., regenerating -ketoglutarate) [179]. The resulting ketoacids can be oxidized or recycled as intermediaries of the TCA cycle. In T. cruzi epimastigotes, a number of amino acids are able to be transaminated with -ketoglutarate or pyruvate to yield Lglutamate or L-alanine, respectively [151, 193]. This robust transamination system helps prevent the accumulation of NH3, and instead results in the accumulation of alanine, which is less toxic [150, 151]. Thus, both cytoplasmic and mitochondrial pools of pyruvate can be transaminated yielding pools of L-alanine. The occurrence of D-alanine in L. amazonensis [198] and the fact that a gene for alanine racemase (AR) was found in the T. cruzi genome led us to hypothesize a role for this isomer. When glycolysis is active, pyruvate accumulates. Upon glucose exhaustion, metabolism shifts from a metabolism based on glucose consumption to a metabolism based on amino acid consumption. L-alanine is actively generated as a product of transamination [52, 150, 151]. It is possible that the racemization of this amino acid could act as a regulator of the intracellular balance between L-alanine and pyruvate by creating a chemical compartment (D-alanine does not interfere with metabolism) as a way to circumvent the lack of the urea cycle and to control the excess ammonia. Experiments are being conducted in our laboratory to confirm this proposed role of D-alanine. Due to their recognized relevance, T. cruzi transaminases have been studied in some detail. All of them are pyridoxal phosphate (PLP)-dependent enzymes, are present in both subcellular compartments, the cytosol and the mitochondrial matrix, and are developmentally regulated at the protein level during the life cycle of T. cruzi [179, 196, 199-202]. ASATs from T. cruzi transaminate aromatic and dicarboxylic amino acids using -ketoglutarate as a cosubstrate, in addition to other –NH2 donors such as glutamine, alanine and leucine (among others). Furthermore, the mitochondrial isozyme is highly specific towards aspartate/-ketoglutarate, yielding the glutamate and oxaloacetate required for the Krebs cycle [150, 179, 202]. It has also been postulated that ASATs in parasitic protozoa might also be involved in methionine recycling [203]. T.


cruzi TAT is also able to transaminate the three aromatic amino acids (tyrosine, phenylalanine and tryptophan) as well as leucine, methionine and alanine [204, 205]. Among the acceptor groups are pyruvate (which is converted into alanine), -ketoglutarate (which is converted into glutamate) and oxaloacetate (which is converted into aspartate). Although pyruvate has been described as the best – NH2 acceptor for transamination, -ketoglutarate and oxaloacetate are also efficient co-substrates [179, 200, 205]. As TAT catalyses a reversible transamination of aspartate or alanine and the aromatic amino acids, it also seems to be a key enzyme in the biosynthesis of these amino acids [150]. Unlike the others transaminases, ALATs are highly specific and almost as efficient as TAT to reversibly transaminate the substrate pair alanine/-ketoglutarate and pyruvate/glutamate [201]. The fact of being pyruvate a main acceptor for the excess of –NH2 produced from amino acids metabolism is consistent with the fact that, in T. cruzi epimastigotes, alanine is one of the most abundant free amino acids [177]. Glutamate can also be interconverted into -ketoglutarate via glutamate dehydrogenases. Both isoforms are involved in the reversible metabolism of amino acids through their ability to transfer -NH2 to ketoglutarate. Two isoforms of this enzyme have been described in T. cruzi, one being NADP+-dependent (cytosolic) [197] and the other being NAD+-dependent (mitochondrial) [206]. The combined effects of aminotransferase and glutamate dehydrogenase activity replace the need for individual dehydrogenases for each amino acid [179, 207]. T. cruzi is also capable of metabolizing histidine into glutamate, which can again be converted into -ketoglutarate [145]. In the T. cruzi genome database, several putative ORFs for key enzymes involved in this pathway were identified: histidine ammonia-lyase (systematic name Tc00.1047053506247.220), urocanate hydratase (Tc00.1047053504045.110), imidazolone propionase (Tc00.1047053509137.30 and Tc00.1047053508741.140) and formimino-glutamase (Tc00.1047053507963.20 and Tc00.1047053507031.90). The corresponding products are currently being expressed and studied by our group. The enzymes required for the metabolism of histidine are upregulated in parasite forms found in the insect vector [145, 208]. Importantly, this result correlates with the occurrence of histidine as one of the predominant free amino acids in the excreta and hemolymph of the T. cruzi vector, Rhodnius prolixus [209-211]. Leucine, isoleucine and valine are all usually oxidized by the same enzymes. In the T. cruzi genome database, no genes coding for specific branched chain amino acid dehydrogenases or transaminases were found. However, as already mentioned, TAT could fulfill this activity by using -ketoglutarate as the amine acceptor [205]. Next, the degradation of these amino acids is catalyzed by the subunit of 2-oxoisovalerate dehydrogenase (T. cruzi genome initiative at the Sanger Center [www.genedb.org]; systematic names Tc00.1047053506629.220, Tc00.1047053509829.20 and Tc00.1047053506367.10), yielding 3-methylbutanoyl-CoA from leucine, (S)-2-methylbutanoyl-CoA from isoleucine and isobutyrylCoA from valine. Leucine might ultimately be converted to acetylCoA and acetoacetate, whereas isoleucine and valine are converted to acetyl-CoA [77]. The Specific Case of L-Proline Whereas the trypomastigote forms of T. cruzi use mainly glucose, the epimastigotes preferentially obtain their energy from amino acids, particularly proline, aspartate and glutamate, which are constituents of the hemolymph and tissue fluids of the invertebrate hosts [154, 212]. Interestingly, intracellular forms (amastigotes and intracellular epimastigotes) fulfill their energy requirements mainly from metabolites other than glucose, especially proline [176]. As previously mentioned, this amino acid also participates in the differentiation process from T. cruzi epimastigote to trypomastigote in the insect vector [186, 187, 213] and in the dif-

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ferentiation of the intracellular epimastigote to the trypomastigote forms in the mammalian host, a process that occurs in the cytoplasm of infected mammalian host cells [188]. In addition, it is well established that adhesion to the host cell surface prior to invasion requires the parasite to expend energy [214]. Martins et al. demonstrated that this energy requirement, in the case of metacyclic trypomastigotes, is supplied mainly by proline, which is able to rapidly restore ATP levels after starvation. This finding demonstrates that proline is able to provide the energy needed to support parasite infectivity, growth and differentiation [215]. In T. cruzi epimastigotes, L-proline is converted into five intermediates of the Krebs cycle (citrate, isocitrate, malate, succinate and oxaloacetate) via glutamate [192]. In most eukaryotic organisms, proline is converted into glutamate through two enzymatic steps coupled to a non-enzymatic reaction, all occurring in the mitochondria. The first step of this conversion is the oxidation of Lproline to 1-pyrroline-5-carboxylate (P5C) by the FAD-dependent enzyme, proline dehydrogenase (PRODH). The P5C (in the mitochondrial environment) is spontaneously interconverted into glutamate semi-aldehyde (GSA). Both species exist in equilibrium through this non-enzymatic reaction. In the second step of the pathway, the 1-pyrroline-5-carboxylate dehydrogenase (P5CDH), which is an NAD(P)+-dependent enzyme, completes the conversion of proline into glutamate by catalyzing the oxidation of GSA [216, 217]. In most organisms, the biosynthesis of proline has two formation routes. The enzyme 1-pyrroline-5-carboxylate synthetase (P5CS) catalyzes the transfer of the phosphoryl group from ATP to glutamate, which in turn is reduced to glutamate semi-aldehyde (GSA). This reduction depends on NADPH. As mentioned earlier, GSA spontaneously interconverts into P5C, which is used as a substrate that can be reduced to proline by 1-pyrroline-5carboxylate reductase (P5CR), an enzyme that uses NADPH as an electron donor. Another route for the formation of proline is through GSA catalyzed by the enzyme ornithine aminotransferase [217, 218]. In the case of T. cruzi, there is no biochemical evidence for the existence of the latter enzyme and no putative genes coding for it have been detected [150]. However, genes encoding a P5CS (P5CS, systematic name Tc00.1047053509067.70) and P5CR (P5CR, systematic name Tc00.1047053506857.20) were found in the parasite genome [77]. These facts are consistent with previous observations of the lack of a functional urea cycle in T. cruzi [195, 219]. It was also established that in other organisms (fungi, plants, bacteria and mammalian cells), the proline degradation pathway also plays several important roles in the control of the cellular redox state [220], apoptosis [221-223], and the resistance to multiple types of stress, including osmotic [224], thermal [225, 226] and oxidative [227-229] stress. Furthermore, the role of proline seems to be paradoxical because it was established in other organisms that proline oxidation generates reactive oxygen species (ROS), consequently being involved in the induction of apoptosis [222], but in some organisms under stressful conditions (oxidative, nutritional, osmotic) the accumulation of free proline appears to exert a protective effect against oxidative stress. In this sense, it was proposed that proline could play either role, depending on the metabolic state of the cell [230-232]. The participation of the accumulation of proline, as well as that of a main metabolite of its oxidation (glutamate) in the resistance to oxidative stress in T. cruzi has already been demonstrated [180, 181]. Amino Acid-related Pathways and Enzymes as Drug Targets As most amino acid metabolic routes involve the mitochondria, here we have grouped targets which are not necessarily mitochondrial, but which affect mitochondrial function. Amino acids have not yet been exploited for therapy. However, as several peculiarities have been identified in T. cruzi amino acid metabolism they are


now being explored for the development of new drugs [150]. The T. cruzi arginine kinase [233], as well as the proline racemase [186], both of which are absent in the mammalian hosts, were shown to be interesting targets for therapy. Due to their structural and functional peculiarities, ALAT and TAT were proposed as interesting drug targets [204, 234]. In addition, it was recently shown that proline transporters could be relevant targets [180]. Glucose- and Proline-derived Metabolites in the Mitochondria For many years it was believed that most trypanosomatids, when grown in glucose-rich medium, obtain their ATP primarily through the mitochondrial F0/F1 ATP synthase (oxidative phosphorylation), by exploiting the proton gradient between both sides of the inner mitochondrial membrane generated by respiratory chain. However, the role of the F0/F1 ATP synthase in energy production in the presence of glucose has been questioned [143, 235]. Indeed, it was shown that procyclic forms of T. brucei did not experience a change in intracellular ATP levels and cell viability in the presence of glucose and an excess of oligomycin (a specific inhibitor of F0/F1 ATP synthase). However, when the parasites were cultured in a medium poor in glucose, the cells were more sensitive to oligomycin. This finding supports the idea that, in the presence of glucose, procyclic forms are not dependent on oxidative phosphorylation for ATP production and that the majority of the ATP is produced by substrate level phosphorylation via phosphoglycerate kinase (PGK), pyruvate kinase (PK) and mitochondrial succinyl CoA synthase (SCS). In the absence of glucose, cells go through a metabolic switch to an amino acid-based catabolism, in which oxidative phosphorylation is essential [154, 236]. Experiments with intact T. cruzi and T. brucei cells showed that glucose, succinate and proline are suitable substrates for stimulating oxygen consumption [192, 237, 238]. Both glucose and proline may generate succinate as a result of their degradation [151]. Proline is converted into glutamate and further converted to -ketoglutarate, which in turn can produce succinate [192]. Proline metabolism has the advantage of avoiding the glycolytic process in T. cruzi, generating malate, and further succinate from fumarate [238]. Moreover iRNA studies in T. brucei revealed that the loss of proline dehydrogenase, succinate dehydrogenase and F0/F1-ATP synthase activities in glucosedepleted medium are conditionally lethal for cells. These results indicate that L-proline (among other oxidizable sources) is extremely relevant for trypanosomatid survival, probably by feeding into oxidative phosphorylation which is critical to manage energy metabolism in glucose-poor media [236, 239]. Succinate is a Key Intermediate Linking the Krebs Cycle and the Respiratory Chain There is evidence that when O2 is largely available and CO2 is kept low, a considerable portion of carbohydrate metabolism functions via the Krebs cycle, which feeds NADH to a respiratory chain with functional oxidative phosphorylation. However, in low O 2 tensions and high CO2, a likely situation inside the host cell, the regulation of succinate dehydrogenase by oxaloacetate (reported as an inhibitor of this enzyme) may be crucial to modulating the metabolic pathways [240]. The glucose degradation produces malate which can be a substrate of three enzymes: 1- malate dehydrogenase producing oxaloacetate either in the glycosome or mitochondria [241, 242]; 2malic enzyme producing cytoplasmic pyruvate [243]; and 3- enter to the mitochondrial matrix to be converted into fumarate or NADH. Thus, when fumarate is generated, both products can form succinate through the reaction catalyzed by the NADH-fumarate reductase [238, 244]. The NADH-fumarate reductase is absent in mammalian cells but it has been identified in trypanosomes and related species [245]. Its function has been a subject of controversy due to its particular activity [194, 246]. Some authors postulate that this enzyme is of bifunctional nature and is involved in both succinate and fumarate


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regeneration [194]. However, Turrens et al. demonstrated that both enzymes (NADH-fumarate reductase and succinate dehydrogenase) act separately. In addition, an important function attributed to this enzyme is NAD+ generation in the mitochondria, especially considering the proposed absence of a functional complex I [247]. Malonate is a competitive inhibitor of succinate dehydrogenase. Moreover, it may decrease the rate of oxygen consumption by 50%, but its effect is reversible upon succinate addition. It also inhibits proline- and succinate- supported respiration by 50% and can compete for the mitochondrial dicarboxylic acid carrier, hampering malate import into the mitochondrial matrix [246]. NADH-fumarate reductase is located mainly in the mitochondria [245], but it was also found in the glycosomal fraction [159, 160]. The identification of this enzyme outside the mitochondria suggested that the enzyme may provide more than one route to the regeneration of extramitochondrial NAD+ during anaerobic conditions in trypanosomatids [245]. For this reason, this enzyme may be a unique target for the treatment of Chagas disease [245]. Succinate, -glycerophosphate and NADH were the first substrates shown to be oxidizable by mitochondrial extracts from T. cruzi [248]. These oxidation processes, with the exception of NADH oxidation, are the only ones that are partially sensitive to relatively high concentrations of cyanide and antimycin, but are not sensitive to salicylhydroxamic acid (SHAM) and considered insensitive to rotenone [248]. Interestingly, NADH oxidation is completely insensitive to cyanide and 2-heptyl-4-hydroxyquinoline-Noxide (HOQNO). These data constituted the first evidence suggesting an NADH oxidation mechanism that involves either a cyanidesensitive pathway, a cytochrome-independent oxidase or peroxidases [248]. The succinate produced either by proline or glucose metabolism is considered the main electron donor to succinate dehydrogenase (the substrate of respiratory complex II) [238, 244]. However, it was observed that, in addition to being oxidized, it can also be excreted into the medium [238, 244]. Additionally, the presence of a gene encoding a succinate semialdehyde dehydrogenase gives support to the existence of an additional source of this metabolite from succinate semialdehyde, a byproduct of -aminobutyric acid (GABA) transamination. It is worth to stress that GABA specific uptake and occurrence in T. cruzi epimastigotes was recently shown (Mantilla et al., unpublished data). Thus, succinate produced on either side of the inner mitochondrial membrane may be transported across the inner membrane for oxidation by the respiratory chain under aerobic conditions [245]. Succinate excretion in trypanosomes, particularly under anaerobic conditions, suggests that this metabolite may become a sink of reducing equivalents [238, 245]. Considering this dual involvement, it has been proposed that interfering with succinate production in T. cruzi should be lethal for the parasite [245, 246]. Peculiarities of T. cruzi Bioenergetics Due to its large size, shape and structure, it is not possible to obtain whole mitochondrial preparations from trypanosomatids in a classical manner. Moreover, this isolation is complicated by the presence of microtubules in the cell membrane, which require stronger forces to homogenize the cells than those necessary for disrupting the mitochondrial membrane [238]. Since Vercesi et al. [249, 250] demonstrated that the treatment of T. cruzi epimastigote cells with digitonin (60 μM) enables the in situ analysis of intact mitochondria, several open questions about this organelle have been elucidated. The data obtained using this approach indicate that T. cruzi mitochondria behave similarly to vertebrate mitochondria regarding the properties of their electrochemical proton gradient. Thus, by permeabilizing the parasite plasma membrane with the appropriate concentration of digitonin, it was possible to determine several biochemical parameters such as the respiration rates at different energy states, the mitochondrial membrane potential ( m


= 70 mV), O2 consumption, as well as ions fluxes (Ca2+, K+) and the redox state. To this extent, some relevant aspects of the respiratory chain components (participating an alternative terminal oxidase) [155] and functions related to them have been described. For instance, the electrical membrane potential, generated by the pH gradient, is the major component of the electrochemical proton gradient across the inner mitochondrial membrane in this species [250]. Studies performed using T. brucei mitochondria indicate that the Krebs cycle is inefficient [158]. However, in the case of T. cruzi, this assumption should be considered carefully, particularly when taking into account the differences in the physiological conditions and nutrients available to supply its energetic requirements throughout this parasitic life cycle. In the invertebrate hematophagous (triatomine vector) host, the assumed catabolism of glucose by epimastigotes should trigger oxidative phosphorylationbased ATP synthesis [52, 151, 154]. Glucose is aerobically oxidized to produce pyruvate [251, 252]. Therefore, the parasites migrate to the terminal portion of insect gut (poor nutrient medium). Then, a metabolic switch occurs so that the parasites now rely on the energy obtained mainly via proline, glutamate and aspartate degradation, which are predominantly abundant in the hemolymph and tissue fluids [192, 253]. The presence of these amino acids promotes metacyclogenesis, probably by acting as the fuel necessary for the whole cell remodeling process [213, 254]. It is well known that NH3, succinate (the substrate of the complex II), ATP and reducing equivalents that can be reoxidized in the mitochondria are produced [52, 151, 154, 236]. Succinate and NH3 have been reported to be excreted metabolites [151]. As previously mentioned, succinate excretion depends on oxygen availability [240]. Trypanosomatids are described as urotelic and ammonotelic on the basis of the proven absence of urea excretion or urea-cycle enzymes; being the NH3 secreted [195]. After the invertebrate host takes a blood meal, the invasion process begins. Metacyclic forms infect the mammal host cells reaching the cell cytoplasm to differentiate and initiate their replication process. Therefore, the parasite faces a medium that is poor in free glucose, among other changes. For instance, neither mRNA expression nor glucose uptake activity was seen in the intracellular amastigotes, giving (this form) priority to the proline metabolism as the main ATP source [176, 188]. Furthermore, after bursting into the extracellular medium, trypomastigotes are able to degrade amino acids but their metabolism seems to rely on glucose metabolism consumption [176] (Fig. 3). Due to this alternation between replicative and infective forms, with different energetic supplies, this species represents a particularly interesting model for bioenergetics studies. Electron Transport Chain in T. cruzi It is well established in almost all eukaryotic cells that the electron transport chain is composed of four integral complexes in the inner membrane of the mitochondria. NADH ubiquinone oxidoreductase (complex I), succinate:ubiquinone reductase (complex II), ubiquinol:cytochrome c3+ oxidoreductase (complex III) and cytochrome c2+ oxidase (complex IV), together with ubiquinone (Coenzyme Q) and cyt c, function as mobile electron carriers between the complexes [142]. The electrochemical gradient generated by complexes III, IV, and I in some organisms, is used by the FoF 1 ATPase (complex V) to generate ATP (Fig. 5). In trypanosomatids (for example T. brucei), the mitochondrion switches between an active and fully repressed state along the different life cycle stages [154]. In this organism, the presence of a fully functional electron transport from complex II to complex IV has been demonstrated, but the participation of complex I is debated because four membrane subunits are missing [62]. The T. brucei complex I consists of 19 subunits with a molecular mass of 660kDa, as predicted by genomic data. It contains all of the subunits with redox centers, so it should be capable of contributing to electron transport across the mitochondrial inner membrane. However,

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the fact that it lacks four integral membrane subunits that are possibly involved in the vectorial translocation of H+ across the membrane supports the proposal that T. brucei complex I is not involved in energy generation, but rather its major function is related to mitochondrial NADH reoxidation [62]. Furthermore, an alternative rotenone-sensitive NADH-dehydrogenase has been found in the T. brucei procyclic form. It is located in the inner mitochondrial membrane where it oxidizes NADH and transfers the electrons to ubiquinone [255]. Two sequences in for the T. cruzi CL-Brener strain in the genome database (accession numbers: XM807104 and XM805934) showed 67% identity with the previously reported rotenone-insensitive NADH-dehydrogenase from T. brucei [256]. Because these enzymes are absent in the mammalian host, their study would be of immense interest [257]. A recent study using natural T. cruzi mutants with deletions in the ND4, ND5 and ND7 genes, which encode complex I subunits, showed that no significant differences were detected in oxygen consumption or respiratory control ratios in the presence of NADH-linked substrates or FADH2-generating succinate when compared to the wild type. Indeed, no correlation could be established between the m and the deletions analyzed. These data, in addition to previous reports that showed a decreased sensitivity to rotenone [194, 248], support the idea that complex I has limited function in T. cruzi [257] (Fig. 5). Mitochondrial complex II, involving succinate dehydrogenase (a membrane-bound Krebs cycle enzyme), often plays a pivotal role in the adaptation of parasites to environments in their host, as reported for helminth parasites [258, 259]. This complex consists of four subunits. The flavoprotein subunit, called SDH1-Fp, is covalently bound to FAD and catalyzes the oxidation of fumarate to succinate. The iron-sulfur subunit, SDH2-Ip forms a soluble heterodimer which binds to a membrane anchor heterodimer referred to as SDH3-CybL and SDH4-CybS. SDH2-Ip transfers electrons to ubiquinone via two functional highly conserved domains that are also present in plants (ferredoxin domain IpN) and bacteria (ferredoxin domain Ipc), in which they located in the amino- and carboxy-terminus, respectively [260]. Ubiquinone is bound and reduced in a pocket formed by SDH2-Ip, SDH3-CybL and SDH4CybS. The latter two contain three transmembrane helices and coordinate protoheme IX in a histidine-depenedent manner (reviewed in [260]). The T. cruzi complex II is an unusual supramolecular complex with a heterodimeric iron-sulfur subunit and seven novel non-catalytic subunits. It is also organized into six hydrophilic (SDH1, SDH2N,SDH2C, and SDH5–SDH7) and six hydrophobic (SDH3, SDH4, and SDH8–SDH11) nuclear encoded subunits each (Fig. 5). This supramolecular structure and the heterodimeric SDH2 subunit (SDH2N and SDH2C) are conserved in Trypanosomatida [260]. The T. cruzi complex II molecular mass was determined experimentally as ~550 kDa, which is four-fold larger than the bovine and yeast (130 kDa) or potato complex II (150 kDa). A two-dimensional analysis of the purified complex II showed that it was composed of 12 subunits that form a homodimer [260]. T. cruzi complex II binds a stoichiometric amount of protoheme, indicating that the monomeric enzyme complex contains one heme. Although heme has an important role in the assembly of complex II, it is not essential for the reduction of ubiquinones [260]. These structural and catalytic features are unique, prompting the further study of this enzyme as another potential therapeutic target. Proteomic analysis during the metacyclogenesis process in T. cruzi revealed the presence of subunits IV and V for complex IV [146, 261]. Subunit IV (ID number: EAO00075) showed a 1.56fold of expression in the epimastigote when compared to the metacyclic stage [261]. The presence of the FoF1 ATPase (complex V) was also evident [146, 261]. Additionally, the protein expression levels of an NADH dehydrogenase, a Rieske iron–sulfur protein, cytochrome c1, and cytochrome c oxidase subunits VI and VIII, as



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Fig. (5). Schematic representation of the electron transport chain in T. cruzi. The major complexes of the electron transport chain are present in the inner membrane mitochondrial (IMM), and some enzymes/intermediates are generated in the inter membrane space (IMS) or mitochondrial matrix. The composition of each subunit is based on the available data from the literature. Complex I (NADH-dehydrogenase) is formed from different subunits, as predicted in other related species (more information at http: //www.icp.be/_opperd/nadh_dh.html). Four subunits are absent (underlined) from the natural strains of T. cruzi (CLBrener, Esmeraldo). The T. cruzi complex I is not sensitive to rotenone (dot line), suggesting that NAD+ regeneration occurs through FMR. Complex II (succinate dehydrogenase) was recently characterized. SDH contains a heme center, six hydrophilic subunits (sdh1, sdh2n, sdh2c, sdh5-sdh7,) and six hydrophobic subunits (sdh3, sdh4, sdh8-sdh11). FMR: fumarate reductase; UQ: ubiquinone; GPD: FAD-dependent glycerol-3-phosphate dehydrogenase; Gly-3P: glycerol3-phosphate; DHAP: dihydroxyacetone phosphate; SHAM: salicylhydroxamic acid; SOD: superoxide Ddsmutase; Cyt c, Cyt b, Cyt a611 , Cyt c558, aa3: cytochromes c, b (antimycin A-sensitive), a611 (referred to as the terminal oxidase, which absorbs at 611 nm), c558 (cyanide-sensitive), CCCP (carbonyl cyanide 3chlorophenylhydrazone). The F0/F1 ATP synthase is responsible for ADP phosphorylation resulting from the proton-motive force generated through an electrochemical gradient. This enzyme also showed Mg-dependent phosphatase activity.

well as the -subunit of the ATP-synthase were analyzed in epimastigotes. These results are in agreement with the model purposed for the respiratory chain and oxidative phosphorylation in these stages [146, 155] (Fig. 5). Studies with epimastigote cells from T. cruzi showed cell growth inhibition when using low cyanide concentrations (ranged from: 0-300 μM) [155]. The effect of cyanide on cytochrome aa3-deficient, dyskinetoplastic epimastigotes supported cytochrome aa3 as T. cruzi’s main terminal oxidase. This work was complemented with cytochrome studies with intact epimastigotes and mitochondrial membranes, revealing the presence of cytochromes aa3, b, c558, o and possibly d, as components of the T. cruzi electron transport system [262]. It was then possible to compare the functionality of cytochrome aa3 with the canonical cytochrome c oxidase for other eukaryotes. Cytochrome o was also described in T. cruzi, but its function as a terminal oxidase is not clear because antimycin considerably inhibits the respiration rates [263]. The T. cruzi respiratory mechanism does not appear to be regulated during its life cycle because all stages are sensitive to cyanide [264]. The sensitivity to both cyanide and antimycin points to the presence of an alternative oxidase (cytochromes of the b, c, or a type) [155, 263]. The fact that the terminal oxidase, (Abs611

nm) does not oxidize horse heart ferrocytochrome c shows that the a-type oxidases from trypanosomes are likely specific for ferrocytochrome c558 (Fig. 5) [240, 265]. The cytochrome c oxidase requires heme A as the prosthetic group [266]. In S. cerevisiae, heme A biosynthesis occurs at the mitochondrial level. This reaction is catalyzed by two mitochondrial inner membrane enzymes, heme O synthase (HOS or COX10) and heme A synthase (HAS or COX15) [266]. The products of the T. cruzi genes COX10 and COX15 were cloned and their functionality was confirmed by functional complementation of yeast mutants lacking the orthologous genes [267]. The mRNA expression level of the genes coding for these proteins changes during the parasite life cycle, suggesting that this variation could reflect the varying respiratory requirements in the different parasite life stages [267]. Taken together, all of these data show that T. cruzi possesses a functional transport electron chain, with evidence for complexes IIV (Fig. 5). The data related to complex I indicate that either complex I has low activity or, less likely, that complex I mutations do not affect mitochondrial respiration and the coupling of respiration with oxidative phosphorylation [257]. This conclusion is only valid for axenic cultures of epimastigotes grown in a glucose-rich me-


dium. Under this condition, ATP production would be obtained mainly by substrate level phosphorylation and, probably, in the mammalian bloodstream trypomastigotes [151]. However, for the insect stage forms and the intracellular amastigotes, which rely on amino acid catabolism, oxidative phosphorylation becomes essential [151, 154]. The Mitochondrion as a Key Organelle in Reactive Oxygen Species (ROS) Production in Aerobic Organisms and T. cruzi Aerobic metabolism utilizes molecular oxygen (O2) as the final electron acceptor in the respiration chain. The O2 is, in turn, reduced by four electrons to two molecules of water. Nevertheless, the O2 may also be reduced to relatively stable species by accepting one, two or three electrons, with the formation of superoxide anion (O2-), hydrogen peroxide (H2O2) and hydroxyl radical (OH-), respectively [268, 269]. The term reactive oxygen species (ROS) is frequently used to describe a group of molecules and free radicals with one unpaired electron derived from redox processes involving O2 [270]. It was first proposed that mitochondria can produce ROS in 1966 [271]. It was found that a significant amount of H2O2 was generated as a byproduct of respiration when the respiratory chain was overloaded. H2O2 is not reactive by itself, but it can oxidize amino acid residues, such as cysteine or methionine, in proteins. Furthermore, H2O2 can be transported outside of the mitochondrion in the presence of metals (such as Fe2+ and Cu2+) and can also form hydroxyl radicals (OH-), which are very reactive for a great number of molecules [reviewed in [272]]. H2O2 has also been described as a cellular signaling molecule that can modify the activity of redoxsensitive proteins in the mitochondria and cytosol [272]. Other studies have demonstrated that H2O2 arises from the dismutation of the proximal ROS, anion superoxide (O2-), which is produced directly by the interaction of molecular O2 with electrons leaking from the respiratory chain [273, 274]. The O2- is dismutated to H2O2 by a mitochondrial manganese superoxide dismutase (MnSOD), which enables the H2O2 to then diffuse from the mitochondria to the cytoplasm. O2- can also be produced in the respiratory chain by complex III when treated with the inhibitor antimycin [274]. An additional source of induced O2- production in the electron chain transport is complex I. The mechanism described involves the presence of the complex I substrate, NADH, leading to superoxide formation [272, 275, 276]. In 1980, Turrens and Boveris [277] showed that complex I is able to produce O2- under two conditions: 1) when there was a high concentration of NADH in the presence of rotenone (complex I inhibitor) and 2) during reverse electron transfer when a high proton motive force and a reduced coenzyme Q-pool (Qred) occur at the same time. Thus, reverse electron transport through complex I is favored. The O2- anion is not dangerous by itself, in spite of being able to disrupt certain ironsulfur-center proteins [269, 270]. O2- formation occurs on the outer mitochondrial membrane, in the matrix and on both sides of the inner mitochondrial membrane. When the O2- generated in the mitochondrial matrix disappears, another portion of the O2- produced in the intermembrane space may be carried to the cytosol through voltage-dependent anion channels [278]. The respiratory chain includes a variety of redox centers with standard reduction potentials between -0.32 V (NAD(P)H) and +0.39 V (cytochrome a3 in complex IV). The standard reduction potential for the conversion of molecular oxygen is -0.160 V [279]. Given the highly reducing conditions that are found inside the mitochondria, various respiratory components, including flavoproteins, iron–sulfur clusters and ubisemiquinone, are thermodynamically capable of transferring one electron to oxygen. Moreover, most steps in the respiratory chain involve single-electron reactions, further favoring the monovalent reduction of oxygen [280] (reviewed in [270]). The state when mitochondria are actively respiring is known as “state 3”, and when the respiratory chain is highly reduced it is called “state 4” [281]. This last condition is reached in the absence of ADP (required by the ATP synthase complex) when

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the H+ flux through the ATP synthase diminishes and the H+ gradient builds up, causing electron flow to slow down and the electron transfer along respiratory chain to become more reduced. Thus, the physiological steady state concentration of O2- increases [282], which is the most common condition in most of studies [281]. The rates of O2- production by the electron transport chain is controlled primarily by mass action, increasing both when electron flow slows down (favoring the concentration of electron donors, R•) and when the concentration of oxygen increases [270, 283]. The available information about ROS release in vivo is controversial with respect to the information from isolated mitochondria, and these extrapolations are misleading. In this sense, ROS can be generated by mitochondrial matrix oxidases. For instance, the cytochrome oxidase retains all partially reduced intermediates until full reduction is achieved. It can impair several processes inside the mitochondria such as the Krebs cycle, ATP synthesis, fatty acid oxidation, the urea cycle, amino acid catabolism, heme synthesis and sulfateferrous center assembly [284]. O2- may also reduce cytochrome c (in the intermembrane space) or may be converted to H2O2 and oxygen (in both the matrix and the intermembrane space). Increased steady state concentrations of O2- may reduce transition metals which in turn react with H2O2, producing OH- radicals, or they may react with nitric oxide (NO) to form peroxynitrite (ONOO-). Both, OH- and ONOO- are strong oxidants and nitrating agents, which react indiscriminately with nucleic acids, lipids and proteins [270]. The abovementioned nitrogen derivatives are termed reactive nitrogen species (RNS) [reviewed in [272]]. NO is a vasodilator resulting from the conversion of arginine to citrulline. NO is produced mainly by the macrophages as a first line of defense during phagocytosis [285, 286]. NO production is catalyzed by a an NADPH-dependent enzyme called nitric oxide synthases (iNOS), which is the focus of a large number of studies [287]. NO synthesized at the mitochondrial level can bind to heme groups from cytochrome oxidases and inhibit respiration rates or stimulate the rates of O2- production from complex I [285, 287]. Under conditions in which the cells can maintain homeostasis between ROS levels and anti-oxidant defenses, all these species (ROS and RNS) are maintained at relatively constant intracellular steady state concentrations by different enzymes and antioxidant molecules (as discussed subsequently). An imbalance in this steady state leads to oxidative stress which may be deleterious and even lethal to all forms of life. H2O2 Generation in T. cruzi The formation of H2O2 is well documented in T. cruzi. It was observed that lipid peroxidation and, H2O2 and O2- production can be stimulated by the addition of -lapachone to epimastigote cultures from T. cruzi [288-290]. The H2O2 was mainly concentrated in the mitochondria (92%), with the remaining 8% produced in the endoplasmic reticulum [288-290]. A recent study of epimastigotes treated with naphthofuranquinones demonstrated mitochondrial H2O2 production and specifically decreased mitochondrial complex I-III activities in both epimastigotes and trypomastigotes. These events occurred in parallel to a reduction in succinate-induced oxygen consumption [291]. Assays performed with epimastigotes treated with digitonin and stimulated with succinate supported NADH-dependent O2 consumption and oxidative phosphorylation. This activity fell by 50% when catalase was added, suggesting that electrons from NADH oxidation reduce the O2 to H2O2. This result was confirmed by cytochrome c peroxidase activity and the effect could be reversed by fumarate addition. Taken together, these results suggest a competition between fumarate and O2 for the electrons from NADH, probably at the fumarate reductase level [244, 245]. Another H2O2 producing enzyme is dihydrolipoamide dehydrogenase (LADH) [1.8.1.4]. This enzyme is involved in branched amino acid catabolism, and its activity generates reduced equivalents in the form of NADH. The addition of catalase, which hydrolyzes this molecule to produce H2O and O2, prevented the


action of the NADH-dependent systems, thus supporting H2O 2 production by NADH-supplemented LADH [292]. It was also shown that natural mutants of ND genes (codifying for three complex I subunits) exhibit higher levels of H2O2 release in the mitochondrion [257]. This result cannot be ascribed to a reduced NADH pool, but to a higher availability of total NAD (NAD+ + NADH), as well as to lower respiratory rates and the high m observed in these strains. These data indicate, again, that alterations in complex I subunits do not correlate with ROS production or NADH oxidation in different T. cruzi strains [257]. All of these results support the idea that the T. cruzi mitochondrion naturally produces H2O2 in variable quantities, depending on genetic and environmental factors. In this context, it is relevant to remark that T. cruzi lacks catalase and peroxidase, enzymes that degrade the naturally formed H2O2 [293]. The absence of an efficient system to detoxify T. cruzi from H2O2 release was shown to be relevant in the control of the infection. The trypanocidal compound, nifurtimox (a nitrofuran prodrug), stimulates the respiration rate and H2O2 production in epimastigotes cells from T. cruzi [294] and in mammal tissues [295, 296]. It was recently shown that a Bothrops jararaca L-amino acid oxidase [1.4.3.2], a flavoenzyme that catalyzes the oxidative deamination of an L-amino acid substrate (preferentially hydrophobic amino acids) to an -ketoacid with the concomitant release of H2O2 and NH4, has a consistent trypanocidal activity when evaluated against T. cruzi epimastigotes [297]. Regarding the trypanocidal mechanism of this enzyme, its effect was abolished in the presence of catalase, suggesting that toxicity is derived from the H2O2 production [297]. In vivo experiments showed that rats infected with T. cruzi trypomastigotes showed increased H2O2 levels produced by peripheral blood monocytes and splenic macrophages. H2O2 production seems to be related to specific mononuclear cells that are able to produce H2O2 in the case of a high parasite load [298]. These data also highlight the relevance of innate immunity and H2O2 production in the host resistance to the acute chagasic infection. O2- Generation in T. cruzi and the Infected Host The anion O2- can be produced by cells both enzymatically and non-enzymatically. Enzymatic sources include the NADPH oxidases located on the cell membrane of polymorphonuclear cells, macrophages and endothelial cells [299, 300] and through cytochrome P450-dependent oxygenases [301]. Further studies, with human polymorphonuclear leukocytes (PMNs), which are able to generate potentially cytotoxic metabolites against pathogens, were described. These cells exhibited a burst in O2 consumption associated with the generation of O2-, H2O2 and possibly OH- and singlet oxygen (iO2) [reviewed in [302]]. It is well known that mitochondria-derived ROS act as mediators for different signal transduction pathways such as apoptotic processes [303]. In this sense, it was shown that enhanced ROS production, mainly in the form of O2-, is directly involved in T. cruzi PCD, demonstrating that the T. cruzi mitochondrion is also involved in the modulation of this process. Concomitant changes in m and respiration rate inhibition, probably due to the impairment of ADP/ATP exchange with the cytosol, were observed. Indeed, the oxidative stress state produced by mitochondrial O2- is extremely detrimental to parasite viability [304]. The induction of the production of ROS from both endogenous and exogenous origins is also heavily involved in parasite viability and the evolution of infection. Early reports about the production of O2- by T. cruzi components when inoculated into mice or during experimental infection were approached by using mice immunized with cruzipain, the most abundant T. cruzi cysteine protease. It was also observed that the subcellular localization of the NADPH oxidase subunits was altered in splenocytes from non-immune and cruzipain immune mice, as well as in a macrophage cell line related


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to cytokine production (IL-6 and IL-1). These data led to the proposed use of immunizing agents that favor oxidative conditions that would affect parasite viability [305]. In addition, other studies focused on the effects of T. cruzi infection in cardiac tissues have provided strong data to suggest that during this process, the myocardium is exposed to injuries induced by oxidative stress and that these may contribute to disease progression [306]. Thus, high levels of ROS (mainly O2- and OH-) can be produced as a consequence of the tissue destruction caused by toxic secretions from the parasite, immune-mediated cytotoxic reactions and secondary damage to the mitochondria [306]. H2O2 release has also been related to O2generation by whole parasite extracts or mitochondrial fractions, being and lapachone, nifurtimox and phenazine methosulfate (PMS) well documented cases [290, 294, 307, 308]. In 1983, Docampo et al. [309] provided biochemical and spectroscopic evidences that O2 consumption and O2- and H2O2 release are stimulated in PMNs that are in contact with antibody-coated T. cruzi epimastigotes and trypomastigotes [309]. In addition, free radical intermediates that are apparently unrelated to oxygen-reduction products have also been found in the metabolic pathways of other trypanocidal drugs (benznidazole and crystal violet) that are used to clinically treat Chagas disease or infected blood from donors (used in blood banks of endemic areas in attempts to eliminate parasite transmission by blood transfusion) [310]. The photodynamic action of crystal violet against T. cruzi trypomastigotes was unveiled with some detail. The trypanocidal activity is gated by the photoirradiation of whole blood containing crystal violet and ascorbate, resulting in an increase in the concentration of ascorbyl radicals and the generation of O2- anions [311]. It was also shown that O2- and H2O2 released by splenic cells contribute to the control of the infection in mice that were experimentally infected with T. cruzi [312], confirming the relevance of the host cell redox-state with regard to the treatment of the T. cruzi infection. Ca2+ Involvement at the Mitochondrial Level The first evidence regarding Ca2+ transport into the T. cruzi mitochondria and its effects at the mitochondrial level was reported by Docampo and Vercesi in 1989 [249, 313]. They reported that the addition of Ca2+ to permeabilized epimastigotes evoked a respiratory stimulation. The Ca2+ uptake was shown to happen in a similar manner as in vertebrate cells: through an electrochemically driven uniport mechanism (from a negative-inside membrane potential), with an efflux pathway that appeared to promote the electrochemical exchange of matrix Ca2+ by external Na+ or H+ pumping [314316]. This uptake causes mitochondrial membrane depolarization which is also compatible with the existence of an electrogenic gradient-mediated Ca2+ transport mechanism [249, 313]. The Ca2+ uptake is stimulated by phosphate and inhibited with ruthenium red and is dependent on the ATP content [249, 317]. The fact that Ca2+ depletion cells inhibited respiration and led to the collapse in the mitochondrial membrane potential highlights the connection between this cation and mitochondrial function [317]. In addition, it was shown an extremely high resistance to the deleterious effects of massive Ca2+ loads in comparison with most types of mammalian mitochondria [249]. High Ca2+ levels depend on the redox state of the mitochondria, which are modulated by thiols and NAD(P)H oxidants such as t-butyl hydroperoxide, diamide, and the 1,2naphthoquinone -lapachone [313]. In turn, high levels of Ca2+ produce the release of O2- triggering the death of the cell by apoptosis (with cyt c release) instead of necrosis [318]. This is the opposite result as that reported for mammals [319]. The mechanisms underlying the link between O2- production and Ca2+ accumulation have not been clearly unveiled. However, in summary, these data support the Docampo findings [58, 249, 313] that showed that T. cruzi mitochondria have a high capacity for Ca2+-storage with elevated resistance to mitochondrial permeability transition (MPT) [249]. This allows the organelle to buffer the cytosolic Ca2+ without collapsing completely, and to serve as a source of redox signals, as


described [318]. The Ca2+ content is also involved in other biochemical processes in T. cruzi, such as the regulation of some enzymes with Ca2+-binding domains. It was recently shown that T. cruzi proline dehydrogenase, a recently characterized mitochondrial enzyme, is Ca2+-regulated, which, in turn, also participates in the regulation of the redox state of the mitochondria (unpublished results). Some compounds that alter the Ca2+ homeostasis in T. cruzi have been proposed as chemotherapeutic targets. The first that was tested as a trypanocide was crystal violet, which induces an efflux of Ca2+ from the epimastigote´s mitochondria. Concomitantly, it induces a rapid, dose-dependent collapse of the inner mitochondrial membrane potential [320]. In addition, this compound inhibits the ATP-dependent, oligomycin-, and antimycin A-insensitive Ca2+ uptake in digitonin-permeabilized cells. It was the first assumption that crystal violet is believed to induce its physiological effects by a deregulation of Ca2+ homeostasis, leading to trypanosome cell injury [320, 321]. T. cruzi Defense Machinery Against Oxidative Damage The main mechanisms developed by aerobic organisms in order to attenuate the damage caused by ROS include: 1) the enzymatic defenses and 2) the low molecular weight antioxidants [268]. The enzymatic mechanisms classically rely on the activities exerted by enzymes, such as the following: (i) the cytosolic or mitochondrial copper, zinc-superoxide dismutase (Cu, Zn-SOD) which converts the O2- in H2O2; (ii) manganese-superoxide dismutase (Mn-SOD) which catalyzes the same reaction, but exclusively in the mitochondrial matrix; (iii) glutathione peroxidase, a peroxidase responsible for the reduction of H2O2 using glutathione as an electron donor; (iv) Catalase which catalyses the dismutation of H2O2 to O2 and H2O; and (v) another group of enzymes named peroxiredoxins that work via an enzymatic substitution (ping-pong mechanism) in which a redox-active cysteine center (the peroxidatic cysteine) is oxidized to a sulfenic acid by the peroxide substrate. Furthermore, the recycling of the sulfenic acid back to a thiol may be reduced by ascorbic acid or glutathione [322-324]. The small molecular weight antioxidants include a variety of molecules that directly react with radicals and prevent them from oxidizing important biomolecules such as tocopherols (Vit E), ascorbic acid (Vit C), uric acid, ubiquininone, and various thiolcontaining metabolites such as cysteine, glutathione and ovothiol [reviewed in [269]]. As already mentioned, trypanosomatids lack catalase [325]. Therefore, the production and diffusion of H2O2 through the cell membrane to the extracellular medium does not seem to be operative as a defense mechanism under physiological conditions [289]. Because H2O2 is toxic for T. cruzi and related species, substances that increase H2O2 generation, decrease its consumption by other enzymatic systems, or that catalyze the homolytic breakdown of H2O2 [326] have been considered as potential trypanocidal compounds. This is consistent with the fact that enzymes belonging to the parasite antioxidant network (mainly in the metacyclic stage) are involved in the initial survival of the parasites in the mammalian host. This fact led to the theory that these molecules are new virulence factors for Chagas disease [327, 328]. Superoxide Dismutase Considering these previous findings, the first studies that tried to elucidate the main defense mechanisms of T. cruzi epimastigotes against the accumulation of H2O2, revealed two enzymes, namely, superoxide dismutase (SOD) and peroxidase [293, 329]. SOD is cyanide-insensitive, but azide- and peroxide-sensitive, which suggests an iron-containing enzyme (Fe-SOD) closely related to the bacterial Fe-SOD [330]. In contrast, mammalian hosts have only

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cytosolic Cu- or Zn-containing SOD and mitochondrial Mncontaining SOD [331]. Two genes encoding T. cruzi Fe-SOD [332] were expressed as functional enzymes in E. coli with protective activity against paraquat (a source of O2-) [333]. Four Fe-SOD enzymatic activities (SOD I-IV) have been reported in T. cruzi extracts. SOD-I (20-kDa), SOD-II (60-kDa) and SOD-IV (25-kDa) are located mainly in the cytosol, whereas SOD-III is mitochondrial [334]. In addition, it was also shown that one of the Fe-SODs excreted into the medium has an immunogenic capacity which enables the serum diagnosis of parasite infection [335, 336]. The overexpression of one of these Fe-SOD enzymes in epimastigotes resulted in an increased sensitivity to the trypanocidal drugs, benznidazole and gentian violet [337], probably due to the fact that the increased O2-, caused by both drugs, causes an imbalance in the T. cruzi oxidative defense system in cells which over-express SOD resulting in an increased rate of H2O2 production. Additionally, a more recent work demonstrated that parasites that over-express mitochondrial T. cruzi Fe-SOD were more resistant to the PCD stimulus. Therefore, lower levels of O2- were unambiguously detected in the transfected parasites (Fe-SOD), indicating the role of mitochondrial O2- as a signaling molecule that can trigger parasite apoptosis [304]. These data are in agreement with the proteomic studies of metacyclogenesis [145], indicating that Fe-SOD over-expression may render cytoprotective effects on parasites, making them more resistant to as-yet-unidentified death stimuli present in the insect vector [304]. The research focused on the screening of compounds against the parasite ROS defense mechanisms showed that these enzymes constitute promising drug targets. For instance, the in vitro trypanocidal activity on epimastigotes and amastigotes, and SOD activity inhibition ability were evaluated for three compounds derived from 1,4-bis(alkylamino)benzo[g]phthalazines (named as C1–C3). One of them, C3, behaved as an active SOD inhibitor [338]. Other related derivatives from this compound (as described) were also tested against T. cruzi [339]. The Fe-SOD inhibition was noteworthy, whereas the effect on human Cu/Zn-SOD is negligible [339]. All of those results might indicate that one of the mechanisms of action for these compounds is by blocking the metal ion of the enzyme, leading to a substantial reduction of its activity, which can be correlated with the observed trypanocidal effect [338]. These data together provide strong structure-related evidence regarding the screening of selective trypanocidal compounds. In this context, compounds must exhibit remarkable effects against specific targets with absent or negligible activity in the mammal host cell. Tryparedoxins Tryparedoxins (TXNs) are the most predominant low molecular mass dithiol proteins (around 20-30 kDa) in trypanosomatids. TXNs are cysteine-containing oxidoreductases that can be reduced by T (SH)2 and can also reduce glutathione-protein mixed disulfides, as reported for T. brucei [340, 341]. In T. cruzi, two isoforms of TXNs, a cytosolic tryparedoxin peroxidase (CPX) and another that is located exclusively in the mitochondrion (MPX), have been described. Both belong to the 2-cysteine peroxiredoxin family and are able to detoxify the cells from H2O2, ONOO- and small chain organic hydroperoxides [342]. The presence of an MPX capable of driving crucial redox pathways is considered a requisite for normal parasite metabolism. This group of enzymes, together with trypanothione, trypanothione reductase and NADPH, are the main parasite mechanism that regulates the low steady state concentration of H2O2 [342, 343]. Furthermore, these proteins seem to also be involved in the resistance to benznidazole, as demonstrated in T. cruzi benznidazole-resistant strains [344]. Finally, when the O2- mitochondrial levels are increased, leading to ONOO- formation around the mitochondria, the parasite’s proliferative ability can be maintained by the over-expression of MPX, clearly showing a role for this mitochondrial enzyme in the detoxification of RNS [327].


Ascorbate Peroxidase Ascorbate peroxidase (APX) is a cyanide-sensitive and heatlabile enzyme, which uses ascorbate as the only substrate, producing dehydroascorbate (DHA). No enzymatic activity for this T. cruzi enzyme was detected for other typical electron donors for peroxidases such as diaminobenzidine, pyrogallol, reduced cyt c or guaiacol [293, 325]. These characteristics obscure the physiological role of ascorbate peroxidase. Thus, it was considered possible that ascorbate oxidation may be due to a protoenzyme-like complex of metal loosely bound to a protein present in the parasite [325, 329]. In most organisms, ascorbate regeneration can occur nonenzymatically by interaction with glutathione or through other activities, such as protein disulfide isomerase and DHA reductase [345]. However, in trypanosomatids both enzymatic and nonenzymatic mechanisms have been postulated for this recycling process [346, 347], the former having been confirmed in T. cruzi [341]. The APX activity was detected, firstly, in a microperoxisomal fraction, but this activity was also detected in the cytosol to a limited extent [329], more specifically in the mitochondrion and endoplasmic reticulum [341]. The peroxidase reaction requires one


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hydrogen donor molecule per H2O2 molecule utilized, with the rate of H2O2 detoxification determined by the rate of hydrogen donor production. As a consequence, catalase-lacking trypanosomes (such as T. cruzi) should have intracellular H2O2 concentrations that far exceed those of mammalian tissues [289]. In plants, the ascorbate molecules donate their electrons to the heme-containing ascorbate peroxidase, which is involved in the oxidative defense system. Nevertheless, it was seen that T. cruzi expresses an unusual plant-like ascorbate-dependent hemeperoxidase with activity that is linked to the trypanothione system. The APX (33 kDa) showed the typical absorption spectrum (531559) of a heme-binding protein. In addition, the parasites overexpressing APX showed an increased resistance to exogenous H2O2 [341]. Proteomic evidence showed that both MPX and APX are upregulated during the transformation from the proliferative epimastigote stage into the infective metacyclic trypomastigote stage [145]. These biochemical changes may pre-adapt metacyclic forms with the capacity to detoxify ROS and RNS generated by macrophages during the T. cruzi-mammalian host cell interactions [327].

Fig. (6). The T. cruzi life cycle representing the nutritional sources that the parasite encounters. Predominant forms along the parasite life cycle. The replicative epimastigotes (E) catalyze glucose aerobically when carbohydrates and amino acids are present in the mid-gut of the triatomine vector. When the parasites migrate toward the terminal portion of the intestine, the metabolism relies on amino acids such as proline, glutamate and aspartate (pro/glut/asp), which are essential to promoting the differentiation process in the infective metacyclic trypomastigotes (M). These catabolic processes produce: Krebs cycle intermediates such as succinate (complex II substrate) that can be secreted into the medium during low O2 conditions, energy in the form of ATP, reducing equivalents [NAD(P)H] reoxidized in the mitochondrion and NH3 which also can be secreted due to the absence of the urea cycle. After the vector takes a blood meal, the metacyclic forms are transmitted in feces invading the cell, in an ATP-dependent process. Metacyclic trypomastigotes escape from the phagocytic vacuole shortly after cell entry and differentiate into amastigotes (A) upon reaching the cytoplasm inside the mammal host cell. Amastigotes are intracellular dividing forms that possess an incipient flagellum. Inside this intracellular environment, the CO2 tensions are increased and proline catabolism is a relevant energy and carbon source. After amastigote proliferation, an obligatory transitional stage form, which resembles the epimastigote stage, is present. This form is named the epimastigote-intracelullar (EI) form, in which L-proline is an essential metabolite for the differentiation into the bloodstream trypomastigote form (T). Trypomastigotes eclosion occurs by cellular bursting that releases the forms which will invade other cells or tissues (via blood). This last stage metabolizes glucose to produce succinate or alanine as precursors of other metabolic pathways. The cycle is completed by a new vector feeding on the subsequent transformation in proliferative forms.


Oxidative Stress Resistance Metabolism and Enzymes as Drug Targets In spite of being compartmentalized between the mitochondria and the cytoplasm, enzymes belonging to the oxidative stress resistance machinery affect mitochondrial function as a whole. The previously discussed particularities of the redox metabolism of trypanosomatids, which is based on trypanothione (a thiol polyamine conjugate exclusively present in kinetoplastids) instead of glutathione, are being taken into account for the design of trypanocides. The most targeted enzyme in the T. cruzi redox metabolism is trypanothione reductase. Several strategies have been launched to inhibit it, including the design of inhibitors such as naphthoquinone derivatives, organometallic complexes, tricyclic compounds and polyamine derivatives (Table 1), which showed promising trypanocidal activity [348-356]. CONCLUDING REMARKS T. cruzi mitochondria possess several peculiarities that can be (and are being) used to target new drugs with the goal of obtaining more efficient therapeutic alternatives to treat Chagas disease. Most mitochondrial processes, from DNA replication, through transcription and mRNA maturation (including editing), up to their participation in energy metabolism, the maintenance of redox homeostasis and mechanism for ROS detoxification are deeply different from those of the mammalian host. As mentioned, the metabolism of this parasite seems to be highly adapted to deal with different environments and energy sources (Fig. 6). In terms of metabolism, it seems that in T. cruzi several functions are concentrated in little metabolic pathways. An example of this is the fact that the proline – glutamate pathway, for example, seems to be involved in energy supply and redox balance, resistance to oxidative stress and resistance to osmotic stress among others. The participation of parasite metabolic pathways in processes that, in the mammalian host, are distributed in a broader network of pathways, can give rise to new opportunities to identify new drug targets. ACKNOWLEDGMENTS This work was supported by grants from the Fundação de Amparo à Pesquisa do Estado de São Paulo (FAPESP grants #08/57596-4 to AMS and #09/54325-2 to CW), and Instituto Nacional de Biologia Estrutural e Química Medicinal em Doenças Infecciosas (INBEQMeDI). We are deeply grateful to Z. Yonny for suggesting helpful ideas. REFERENCES [1] [2]

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Received: June 8, 2011

Accepted: June 27, 2011


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