Key words: ATP synthase, cryo-EM, mitochondria, mitochondrial membrane potential, oxidative phosphorylation
Abstract
Mitochondrial F-type adenosine triphosphate (ATP) synthases are commonly introduced as highly conserved membrane-embedded rotary machines generating the majority of cellular ATP. This simplified view neglects recently revealed striking compositional diversity of the enzyme and the fact that in specific life stages of some parasites, the physiological role of the enzyme is to maintain the mitochondrial membrane potential at the expense of ATP rather than to produce ATP. In addition, mitochondrial ATP synthases contribute indirectly to the organelle's other functions because they belong to major determinants of submitochondrial morphology. Here, we review current knowledge about the trypanosomal ATP synthase composition and architecture in the context of recent advances in the structural characterization of counterpart enzymes from several eukaryotic supergroups. We also discuss the physiological function of mitochondrial ATP synthases in three trypanosomatid parasites, Trypanosoma cruzi, Trypanosoma brucei and Leishmania, with a focus on their disease-causing life cycle stages. We highlight the reversed proton-pumping role of the ATP synthase in the T. brucei bloodstream form, the enzyme's potential link to the regulation of parasite's glycolysis and its role in generating mitochondrial membrane potential in the absence of mitochondrial DNA.
Introduction
F-type adenosine triphosphate (ATP) synthases (also called F-ATPases of F1F0-ATPases) are bidirectional turbine-like enzymes coupling ATP synthesis or hydrolysis with proton translocation through biological membranes in bacteria and their endosymbiotic descendants, mitochondria and chloroplasts (for recent reviews see Junge and Nelson, 2015; Walker, 2017; Kuhlbrandt, 2019). When operating in the forward mode, ATP synthases utilize the proton motive force to produce ATP and thus represent fundamental constituents of the oxidative phosphorylation pathway. In the reverse mode, they act as ATP-consuming proton pumps contributing to the generation of electrochemical membrane potential. Bacteria employ both modes depending on the species and growth conditions (Cotter and Hill, 2003). By contrast, mitochondrial ATP synthases in most aerobic eukaryotes function as a major source of cellular ATP and the reversal of their activity is a manifestation of pathophysiological conditions (Campanella et al., 2009). Unlike the monomeric bacterial and chloroplastic counterparts, all mitochondrial ATP synthases characterized so far occur in dimers (Arnold et al., 1998; Dudkina et al., 2005). The wide phylogenetic distribution of the ATP synthase dimers suggests that the enzyme's dimerization is a common feature of all aerobic eukaryotes with oxidative phosphorylation and was possibly present already in the last eukaryotic common ancestor. Dimerization of ATP synthases induces curvature of the inner mitochondrial membrane (Dudkina et al., 2006; Davies et al., 2012) and the intrinsic propensity of dimers to self-assemble into rows facilitates membrane shaping and governs cristae formation (Anselmi et al., 2018; Blum et al., 2019). Spatial separation of ATP synthases and electron transport chain (ETC) complexes on rims and flat regions of cristae, respectively, creates a local proton concentration gradient in cristae lumen, enhancing oxidative phosphorylation (Davies et al., 2011). Therefore, apart from the enzymatic function, ATP synthases affect mitochondrial physiology by aiding in the determination of submitochondrial ultrastructure.
In Trypanosoma brucei, a medically relevant digenetic parasite, the mitochondrial ATP synthase exhibits a 2-fold deviation from its counterparts from traditional model organisms. First, its architecture differs markedly from canonical ATP synthases. Second, its role switches during the parasite's life cycle, from being an ATP producer in the insect form to an ATP consumer maintaining the vital mitochondrial membrane potential in the mammalian bloodstream form (BSF). This switch is associated with major rearrangement of mitochondrial morphology, possibly involving altered ATP synthase oligomerization. Unlike in T. brucei, in Trypanosoma cruzi and Leishmania, ATP synthase acts true to its name and participates in the oxidative phosphorylation providing ATP in all life forms of the parasite. In the first part of this review, we discuss the recent progress in our understanding of ATP synthase architecture based on structural characterization of enzymes from various eukaryotic supergroups with a focus on trypanosomal ATP synthase complex. In the second part, we summarize our knowledge of the ATP synthase importance for the disease-causing forms of medically relevant trypanosomatid parasites.
Unexpected diversity of mitochondrial ATP synthases
Structural studies of mitochondrial ATP synthases have been key for the understanding of molecular mechanisms of ATP synthesis. The seminal research on the structure of the matrix-facing soluble F1-ATPase subcomplex isolated from the bovine heart (Abrahams et al., 1994) initiated two decades of dominance of X-ray crystallography in the field. Numerous crystal structures of subcomplexes of bovine and yeast ATP synthases provided atomic models of the enzyme's rotor and the extrinsic part of the peripheral stalk, and revealed the mechanism of the conversion of the rotational force into the cyclic catalysis of ATP formation (reviewed in Junge and Nelson, 2015; Walker, 2017). Due to the striking similarity of the blueprints for bacterial and opisthokontal mitochondrial F-type ATP synthases and their fundamental role in cellular energetics, it was assumed that the mitochondrial enzymes do not differ substantially between eukaryotic lineages. However, only the advance of cryo-electron microscopy (cryo-EM) techniques in the last few years allowed researchers to build atomic models of complete mitochondrial ATP synthases not only from conventional model organisms [mammals (Gu et al., 2019; Pinke et al., 2020; Spikes et al., 2020) and Saccharomyces cerevisiae (Guo et al., 2017)], but also from protists representing other eukaryotic phyla [Polytomella (Murphy et al., 2019), Euglena (Muhleip et al., 2019), Tetrahymena (Flygaard et al., 2020) and Toxoplasma (Mühleip et al., 2021)]. From the mechanistic point of view, structures revealed by cryo-EM elucidated how the proton motive force across biological membranes is converted into the torque of the enzyme's rotor (Klusch et al., 2017; Hahn et al., 2018) and how the symmetry mismatch of the rotor and catalytic subcomplexes is compensated by the flexibility of the peripheral stalk (Sobti et al., 2019). From the evolutionary perspective, the structures demonstrated that although the functional core is universally conserved in all F-type ATP synthases, peripheral parts and the dimerization interface of mitochondrial ATP synthases have diverged substantially among eukaryotic lineages. Notably, cryo-EM structures not only allowed identification of novel lineage-specific components, but in some species also revealed conserved subunits, which had not been previously identified by traditional homology searches using known genomes and proteomes (Fig. 1).
All F-type ATP synthases are composed of two subcomplexes, the membrane-embedded F0 and the soluble F1, connected by a rotary central stalk and a stationary peripheral stalk (Fig. 2). The globular F1 subcomplex, also referred to as F1-ATPase, is a pseudo-symmetrical assembly of three heterodimers of α- and β-subunits organized around single γ-subunit, the main constituent of the central stalk. The functional core of F0 consists of the a-subunit and a ring of several identical c-subunits (c-ring), providing together a path for protons across the membrane. Proton translocation drives the rotation of the c-ring and the tightly attached central stalk. The torque of the asymmetric γ-subunit induces cyclic conformational changes of three active sites on the interfaces of α- and β-subunits, resulting in the stepwise binding of ADP and phosphate, their condensation into ATP, and its release, emptying the nucleotide binding site for the next catalytic cycle (Abrahams et al., 1994). The (αβ)3-hexamer is prevented from rotation by interaction with the oligomycin sensitivity-conferring protein (OSCP), the uppermost constituent of the extrinsic part of the peripheral stalk.
The entire F1-ATPase consisting of subunits α3, β3, γ, δ and ɛ (stoichiometry of subunits indicated by the subscripts), OSCP and both proton translocating components (a-subunit and c-ring) are present in all reported mitochondrial ATP synthases. The rest of the enzyme is markedly less conserved (Fig. 2). The highest diversity is observed in the soluble part of the peripheral stalk and in the lumenal and peripheral F0 regions, often composed of lineage-specific subunits or lineage-specific extensions of conserved proteins. Subunits involved in the anchoring of the peripheral stalk to the membrane and in clamping the a-subunit to the c-ring (b, d, e, f, g, i/j, k and 8), originally identified in opisthokonts, appear to be conserved in several other eukaryotic groups (Fig. 1). In many cases, their proposed homology is based largely on tertiary and quaternary structure resemblances revealed by cryo-EM. Low sequence similarity precludes identification of the respective homologs in lineages without structurally characterized ATP synthases. Although dimerization is a general hallmark of all reported mitochondrial ATP synthases, subunits constituting the dimer interface differ between eukaryotic lineages, likely due to divergent evolution. Consequently, the shape of dimers and topography of oligomeric assemblies vary in the characterized enzymes, which is reflected in diverse cristae morphology (Fig. 1; Dudkina et al., 2010; Davies et al., 2012; Muhleip et al., 2016; Muhleip et al., 2017; Mühleip et al., 2021).
Unique structural features of ATP synthases in Trypanosomatida
The knowledge about the architecture of mitochondrial ATP synthases in Trypanosomatida, a group of protozoan parasites with monoxenous or dixenous life cycles, is based mostly on studies performed with cultured insect trypomastigotes of T. brucei. The composition of the mitochondrial ATP synthase in these cells was determined by mass spectrometry (MS) identification of proteins co-immunoprecipitated with F1 and tandem affinity purified using TAP-tagged conserved or newly identified subunits as baits. These biochemical analyses complemented by a comprehensive genome search revealed 23 subunits (Zikova et al., 2009), including the a-subunit detected by MS later (Skodova-Sverakova et al., 2015). Characterization of the trypanosomal F1-ATPase released by chloroform from mitochondrial membrane fragments and purified by two-step chromatography (Gahura and Zikova, 2019) showed that the catalytic subcomplex contains three copies of the euglenozoan-specific p18-subunit in addition to the universally conserved subunits α, β, γ, δ and ɛ (Gahura et al., 2018b). The F1-ATPase structure determined by X-ray crystallography further revealed that each copy of p18-subunit, an α-helical protein with three pentatricopeptide repeats, associates peripherally with one of the three α chains (Fig. 3A), and does not contribute directly to the catalytic mechanism (Montgomery et al., 2018). Although the atomic structure did not explain the function of p18, the elaboration of the F1 head in Euglenozoa by this additional subunit is extraordinary, because the α3β3γ subcomplex is invariant in all other known F-type ATP synthases. In addition, the α-subunit was found to be proteolytically cleaved at two sites separated by eight amino acid residues, producing two fragments α1–127 and α135–560, both stably associated with the complex. The split of α-subunit into two parts was also demonstrated in other euglenozoan protists Crithidia fasciculata (Speijer et al., 1997), Leishmania tarentolae (Nelson et al., 2004) and Euglena gracilis (Sathish Yadav et al., 2017). The structure of the trypanosomal F1-ATPase showed that the cleavage occurs at the region corresponding to a loop on the surface of F1-ATPases from other organisms and splits the N-terminal β-barrel domain from the rest of the protein (Montgomery et al., 2018; Fig. 3A and B). Nevertheless, the architecture of the entire (αβ)3-headpiece, including the nucleotide binding pockets, is highly similar to the prototypical bovine F1-ATPase (Bowler et al., 2007) and the biological relevance of the α-subunit cleavage remains unclear.
So far, there is no available structure of a complete ATP synthase from any trypanosomatid species. However, low-resolution structures obtained by subtomogram averaging documented general morphological similarity between ATP synthase dimers from T. brucei and E. gracilis, a representative of euglenids, a lineage closely related to kinetoplastids (Muhleip et al., 2017). An atomic model of dimeric ATP synthase from E. gracilis has recently been determined by cryo-EM (Muhleip et al., 2019). Apart from the universally conserved components of mitochondrial ATP synthases (α, β, γ, δ, ɛ, a, c and OSCP), the E. gracilis enzyme contains seven subunits with previously recognized homology (Sathish Yadav et al., 2017) to proteins found in the complex from T. brucei (Zikova et al., 2009). To get a deeper insight into the trypanosomal ATP synthase structure, we performed systematic pairwise sequence comparison of all remaining subunits in the complex from E. gracilis with all non-conserved trypanosomal ATP synthase components, and propose homology of four additional protein pairs, which exhibit >20% sequence identity (Fig. 3C). Overall, 19 of 23 subunits of the trypanosomal ATP synthase have homologs found in the structure from E. gracilis. Taken together, although the ATP synthases from E. gracilis and T. brucei are not compositionally identical, it is conceivable to assume that the structural features shaped by the conserved subunits are shared between the two species (Fig. 3C), and possibly among Euglenozoa in general.
The peripheral stalk in euglenozoan ATP synthases differs markedly from other lineages. The universally conserved OSCP features a long and partially flexible C-terminal extension, which bridges the core of the protein on the top of F1 with ATPTB2, a highly divergent and extended homolog of the d-subunit from opisthokontal ATP synthases anchoring the peripheral stalk to the membrane. The interaction of ATPTB2 with the extension of OSCP is clamped between two laterally positioned lineage-specific globular proteins ATPTB3 and ATPTB4 on one side and one copy of p18 on the other side. The contacts of the peripheral stalk with p18 contribute to the immobilization of the (αβ)3-hexamer (Fig. 3D). However, the reinforcement of the interaction between the F1 head and the peripheral stalk is unlikely the only role of p18, because the protein is essential for the integrity of the F1-ATPase (Gahura et al., 2018b), in contrast to the peripheral stalk components OSCP (Hierro-Yap et al., 2021) and ATPTB2/d-subunit (Subrtova et al., 2015). In E. gracilis, the extensions of OSCP and d-subunit compensate for the reduction of the b-subunit, the major constituent of the peripheral stalk in bacteria and mitochondria of mammals and yeast. Sequence comparison did not reveal a homolog of the b-subunit in the ATP synthase of T. brucei, suggesting that it might be further reduced or completely absent in Trypanosomatida.
In all reported F-type ATP synthases, the proton channel-forming a-subunit contains two horizontal membrane-embedded helices H5 and H6, which are directly involved in proton translocation (Guo et al., 2017; Murphy et al., 2019). In mammals (Spikes et al., 2020) and yeast (Guo et al., 2017) the two helices are lined by a series of six parallel transmembrane helices, contributed by one helix of each of subunits b, f, 8, i/j and k, and by a transmembrane helix of the a-subunit itself. The same configuration is observed in E. gracilis (Fig. 3E), suggesting the conservancy of the contributing subunits. Although the ATP synthase in T. brucei contains homologs of E. gracilis subunits k and i/j, the homologs of subunits b, f and 8 were not identified. The dimer interface of the ATP synthase in E. gracilis is formed largely by subunits ATPEG1 and ATPEG2 without discernible homology outside Euglenozoa, but with apparent homologs in Trypanosomatida (called ATPTB9 and ATPTB8 in T. brucei; Perez et al., 2014; Fig. 3C and F). The presence of these two proteins in T. brucei together with the fact that the dimers from both species arrange into short left-handed helices, as showed by cryo-electron tomography (cryo-ET; Muhleip et al., 2017), strongly suggests that the dimerization is mediated by the same elements. The dimer in E. gracilis is further stabilized by a homotypic interaction of extensions of d-subunit adopting a ferredoxin-like fold on the matrix side of the membrane. This extension is specific to euglenids and the interaction is therefore completely missing in T. brucei, possibly explaining the poor stability of its ATP synthase dimers (our observation). Notably, the dimer interface in Euglena contains several ordered phospholipid molecules, predominantly cardiolipins (Muhleip et al., 2019; Fig. 3F). Molecules of cardiolipin are presumably integrated also in the trypanosomal ATP synthase, as cardiolipin plays a vital role in the stability of the complex (Serricchio et al., 2020).
Role of the ATP synthase in trypanosomatid parasites
The function of the ATP synthase in trypanosomatid parasites depends on the species and their life cycle stage. Digenetic trypanosomatid parasites (e.g. T. brucei, T. cruzi and Leishmania) possess a complex life cycle as they alternate between the insect vector and a mammalian host. In humans, they cause dreadful diseases (African trypanosomiasis, Chagas disease and leishmaniases). The scarcity of effective treatments and weak vaccine prospects drive an ongoing urgent need to identify new therapeutic targets. Considering the ATP synthase's central position in energy metabolism and its structural and compositional divergence from the mammalian counterpart, one may consider this complex as a promising drug target. The feasibility of targeting the ATP synthase is underscored by the recent approval of bedaquiline, a specific F-ATP synthase inhibitor that blocks the c-ring rotation, to treat multi-drug resistant tuberculosis (Preiss et al., 2015). Whereas the function of the ATP synthase in T. brucei parasites is well understood, the exact role of the ATP synthase and its requirement for virulence and survival of Leishmania and T. cruzi parasites are not well defined yet.
Role of the ATP synthase in intracellular amastigotes of Leishmania and T. cruzi parasites
Leishmania parasites infect a mammalian host when an infected sand fly takes a blood meal. The promastigote form is released to the skin and is rapidly internalized by macrophages. Inside the macrophages, promastigotes differentiate to aflagellated and non-motile amastigotes, which multiply in the acidic environment of the phagolysosome-like parasitophorous vacuole, whose exact metabolic content has not been determined yet. The environment most likely offers a wide range of carbon sources (sugars, amino acids and fatty acids) for the parasite to salvage due to macrophage polarization to the M2 state, which exhibits increased oxidative metabolism and promotes Leishmania parasites growth (Saunders and McConville, 2020). Despite this seemingly plentiful environment, intracellular amastigotes do not proliferate rapidly (Kloehn et al., 2015), but rather activate a stringent metabolic response, a protective mechanism to confer resistance to multiple cellular stresses (oxidative, nutritional and pH). This programmed response is associated with reduced rates of glucose uptake and with low metabolic activity to possibly minimize reactive oxygen species (ROS)-sensitive processes and to lessen the levels of endogenously generated ROS (McConville et al., 2015). The glucose-sparing metabolism of the intracellular amastigotes questions the importance of oxidative phosphorylation, and hence ATP production by the ATP synthase, for the parasite. Nevertheless, amastigotes show increased reliance on mitochondrial metabolism, including tricarboxylic acid (TCA) cycle and fatty acid β-oxidation (Saunders et al., 2014). These pathways generate reduced electron carriers NADH and FADH2 that are reoxidized by the ETC coupled to ATP synthase (Dey et al., 2010; Saunders et al., 2014). The ablation of aconitase, a TCA cycle enzyme, cytochrome c oxidase, or ATP synthase is predicted to be lethal by an in silico-generated model of the metabolic network (Subramanian et al., 2015). Indeed, the inhibition of ATP synthase by the cell-permeable antimicrobial peptide histatin 5 arrests of Leishmania amastigotes' growth (Luque-Ortega et al., 2008). Nevertheless, to prove that oxidative phosphorylation is critical for the Leishmania amastigote forms, further studies are necessary (Fig. 4).
Trypanosoma cruzi parasites are transmitted to humans by a blood-sucking triatomine bug through contact with its infected feces. In the skin wound, metacyclic trypomastigotes invade various types of cells, escape the lysosome-derived parasitophorous vacuole and establish infection in the host cell cytoplasm as intracellular amastigotes (de Souza et al., 2010). During T. cruzi trypomastigotes' transition to amastigotes, there is a dramatic shift from sugar-based metabolism to catabolism of amino and fatty acids manifested by repression of glucose transporters and increased levels of TCA cycle enzymes and proteins involved in fatty acid oxidation and oxidative phosphorylation. The oxidative metabolism of amino and fatty acids suggests a crucial role for oxidative phosphorylation to generate ATP (Atwood et al., 2005; Silber et al., 2009; Li et al., 2016; Shah-Simpson et al., 2016). This is further supported by the identification of highly selective compounds (i.e. GNF7686 and ELQ271) that target complex III and inhibit the growth of intracellular amastigotes in a dose-dependent manner (Khare et al., 2015).
After several rounds of intracellular replication, amastigotes differentiate to trypomastigotes, which are released from the cell by its rupture. Extracellular trypomastigotes disseminate and infect other cells or they are ingested by the triatomine bug to conclude the life cycle. Interestingly, the extracellular bloodstream trypomastigotes switch back to glycolysis to meet their energy demands but still employ ETC complexes to maintain the mitochondrial membrane potential. Moreover, disproportional changes in expression of ETC complexes II, III and IV create a bottleneck forcing electrons to prematurely reduce molecules of oxygen and generate significant levels of ROS (Goncalves et al., 2011). This induced production of ROS probably provides a redox-mediated pre-conditioning to evoke protection against host-induced oxidative challenge once the parasite enters the cell again. In this particular case, the ETC linked to ATP synthase also serves as an adaptive process allowing parasites to survive redox challenges imposed by the host. Presumably, inhibition of the ATP synthase would cause a decrease in ATP levels, which by itself can be lethal, but it would also further increase the already elevated ROS levels. Although there is evidence that the oxidative environment stimulates the parasite's growth and ROS are critical for successful infection of T. cruzi in various cell types (Paiva and Bozza, 2014), very high levels of ROS reverse this favourable effect and are harmful for the parasite (Goes et al., 2016). Therefore, an additive effect of endogenous and exogenous ROS bursts may induce deleterious changes resulting in parasites' elimination by oxidative damage (Fig. 4).
Despite some evidence that mitochondrial ETC and ATP synthase might be essential entities for the clinically relevant infectious stages of Leishmania and T. cruzi parasites, the knowledge of the mitochondrial energy metabolism in these parasites remains largely incomplete.
The F-ATPase in the bloodstream form of T. brucei
The disease-causing form of T. brucei is an extracellular parasite inhabiting various niches of its mammalian host, including the bloodstream, adipose tissue, skin or central nervous system (Silva Pereira et al., 2019). Since these environments offer different nutrients, the BSF mitochondrion maintains its complexity and plasticity despite its largely reduced size compared to the insect stage organelle (Zikova et al., 2017). Uniquely, the parasite's mitochondrion employs ATP synthase as an ATP-consuming proton pump (Fig. 4).
The discovery of the oligomycin-sensitive F-ATP synthase in BSF by Opperdoes et al. (1977), elicited the curiosity of scientists, who back then considered the BSF mitochondrion a vestigial organelle with no obvious role in cellular metabolism. Seeking if this ‘promitochondrion’ possesses mitochondrial membrane potential, Nolan and Voorheis reported the existence of electrochemical gradients in BSF trypanosomes, documented by the accumulation of the radiolabelled lipophilic cation methyltriphenylphosphonium. One year after, the same authors reported that oligomycin caused a collapse of the mitochondrial membrane potential identical in magnitude to that achieved by the proton uncoupler FCCP, concluding that in BSF cells, the nature of the mitochondrial electrical gradient was exclusively attributable to the proton-pumping activity, or reverse mode, of the ATP synthase (Nolan and Voorheis, 1992). In more than 10 years, RNA interference silencing of two F1-ATPase subunits further corroborated the previous findings and confirmed the unusual and essential function of this enzyme for the survival of BSF trypanosomes (Schnaufer et al., 2005; Brown et al., 2006).
The role of the F-ATPase in BSF T. brucei strikingly contrasts with that of other eukaryotes, where the reverse operation of the ATP synthase is a rare and short-term phenomenon associated with hypoxia or anoxia. For example, under ischaemic conditions in mammalian tissues, respiration is halted and the ATP synthase reverses in response to decreased mitochondrial membrane potential, partially compensating for its loss by utilizing ATP produced by glycolysis. To limit the detrimental depletion of cellular ATP caused by an ATP-consuming F-ATP synthase (Rouslin et al., 1986; Jennings et al., 1991; St-Pierre et al., 2000), the activity of this enzyme is regulated by a short naturally occurring protein, the inhibitory factor 1 (IF1) (Pullman and Monroy, 1963; Walker, 1994). IF1 specifically inhibits the ATPase activity with no effect on ATP synthesis (Pullman and Monroy, 1963; Asami et al., 1970), although some authors propose an inhibitory effect of IF1 on both ATP synthase modes (Harris et al., 1979; Schwerzmann and Pedersen, 1981; Formentini et al., 2012; Garcia-Bermudez and Cuezva, 2016). IF1 has also been implicated in cristae biogenesis, dimerization of the ATP synthase and metabolic reprogramming of cancer cells (Campanella et al., 2009). This regulatory protein has been identified in T. brucei (TbIF1) and related trypanosomatid parasites. In T. brucei the expression of TbIF1 is tightly regulated throughout the life cycle as it is expressed only in the insect stages, but not in BSF (Panicucci et al., 2017). In T. cruzi and Leishmania parasites, regulation of IF1 expression is reflected in its slight upregulation in the mammalian intracellular stages compared to the respective insect stages in agreement with increased mitochondrial activity of T. cruzi and Leishmania amastigotes (Li et al., 2016; Inbar et al., 2017). As in other organisms, TbIF1 inhibits the hydrolytic but not the synthetic activity of the mitochondrial ATP synthase in vitro (Panicucci et al., 2017; Gahura et al., 2018a). The artificial expression of TbIF1 is lethal for BSF trypanosomes, due to the abolishment of the mitochondrial membrane potential (Panicucci et al., 2017).
In addition to its role in the maintenance of mitochondrial membrane potential, a functional F-ATPase is also required by BSF parasites to prevent the intramitochondrial accumulation of ATP. Elevated ATP levels lead to inhibition of the alternative oxidase, the only terminal oxidase in BSF parasites, and consequently to a reduced respiration rate (Luevano-Martinez et al., 2020). This represents a novel and unique role of the F-ATPase connecting this enzyme to the regulation of the glycolytic pathway, which is intimately linked to respiration via the alternative oxidase (Bakker et al., 1999). Despite these fundamental roles, BSF trypanosomes can withstand a loss of approximately 90% of the membrane-bound F-ATPase complexes with no obvious deleterious effect on their viability in culture nor on the mitochondrial membrane potential in live cells (Hierro-Yap et al., 2021). We speculate that the long-term tolerance of reduced levels of F-ATPase complexes might be favourable for the emergence of trypanosomes lacking mitochondrial genome, as it might provide the time frame for the parasites to gain key nuclear mutation(s) in the F-ATPase that enable(s) generation of mitochondrial membrane potential in the absence of mitochondrial-encoded a-subunit.
The vestigial F-ATPase in trypanosomes without mitochondrial genome
The mitochondrial DNA of trypanosomatids, which is arranged in a distinct submitochondrial structure termed kinetoplast (hence kinetoplast DNA, kDNA), encodes mostly for the components of the oxidative phosphorylation pathway, key for the survival of T. brucei's insect stages (Stuart, 1983; Schnaufer et al., 2002). In BSF trypanosomes, which do not carry out oxidative phosphorylation, kDNA is indispensable due to the requirement of the F-ATP synthase a-subunit involved in proton translocation across the inner mitochondrial membrane. To synthesize the a-subunit, BSF mitochondria express additional two kDNA-encoded proteins, uS3m and uS12m, core components of mitochondrial ribosomes (Ramrath et al., 2018). However, there are BSF trypanosomes that partially or completely lack their kDNA and they are referred to as dyskinetoplastic (Dk) subspecies (Agbe and Yielding, 1995). The Dk trypanosomes thriving in nature, T. brucei evansi (Hoare, 1937) and T. brucei equiperdum (Tobie, 1951), cause surra and dourine, respectively, devastating diseases that affect a broad range of domestic and wild animals in Africa, Asia and South America (Brun et al., 1998). Stable kDNA-deficient strains can also be generated artificially using compounds targeting the kDNA network, such as ethidium bromide or acriflavine (Stuart, 1971; Riou et al., 1980; Riou and Benard, 1980). Although Dk trypanosomes were thought to be locked in the long slender BSF stage, the absence of kDNA does not impair their differentiation into the stumpy form, a transmission pre-adapted stage with partially activated mitochondrial functions. However, the absence of kDNA does reduce the lifespan of the stumpy form (Dewar et al., 2018), hampering their differentiation into the procyclic form (Timms et al., 2002). Therefore, Dk trypanosomes are transmitted between hosts during sexual intercourse or mechanically via blood-sucking insects or vampire bats (Brun et al., 1998), which facilitated their spread outside the area of the tsetse fly belt (Lun and Desser, 1995).
Because the ATP synthase a-subunit is absent in Dk trypanosomes, the molecular mechanism for the generation of mitochondrial membrane potential is analogous to that of mammalian ρ0 cells and yeast petite mutants, both of which lack mitochondrial genome. The mechanism involves ATP hydrolysis by the vestigial ATP synthase, which provides the substrate for the electrogenic exchange of mitochondrial ADP3− for cytosolic ATP4− by the ATP/ADP carrier (AAC). Accordingly, oligomycin, an inhibitor that blocks the c-ring rotation, has virtually no effect on the growth of T. b. evansi (Schnaufer et al., 2005). Moreover, Dk strains contain one of several point mutations in the nuclear-encoded subunits of the F1-ATPase, most often the L262P or A273P substitutions in the γ-subunit (Lai et al., 2008; Dean et al., 2013). These substitutions are sufficient to allow the loss of kDNA and enable the generation of mitochondrial membrane potential in an F0-independent manner, perhaps by enhancing the hydrolytic activity of the F1-ATPase (Dean et al., 2013).
The functional independence from the F0 section led to the long-standing belief that the F1 moiety exists as a soluble entity detached from the inner mitochondrial membrane in Dk trypanosomes (Schnaufer et al., 2005; Jensen et al., 2008). However, the finding that the silenced expression of the peripheral stalk subunit ATPTB2/d-subunit slowed the growth of Dk trypanosomes in vitro (Subrtova et al., 2015) indicates that the complex might be attached to the membrane. Considering the presence of a membrane-bound F-ATP synthase in cells devoid of mitochondrial genome in other organisms (Wittig et al., 2010; He et al., 2017), it is likely that the vestigial F-ATPase in Dk trypanosomes also remains associated with the membrane to increase the spatial proximity between the AAC and the F1-ATPase, and ensure an efficient exchange of substrates between the two entities.
Perspectives
Although many architectural features of ATP synthase from Euglena can be extrapolated to Trypanosomatida, a high-resolution cryo-EM structure of a trypanosomatid ATP synthase will be necessary for an unbiased and complete description of the enzyme in this parasitic lineage. Such research will reveal structural divergency, predominantly in the membrane-embedded part of the enzyme, as suggested by the absence of apparent homologs of some conserved ATP synthase subunits in T. brucei (Fig. 1). If several rotational states of the enzyme are characterized, the study would explain how the apparently flexible peripheral stalk immobilizes the catalytic subcomplex and what is the contribution of its unprecedented contacts with the F1-ATPase (Fig. 3D). An atomic model of trypanosomal ATP synthase could also reveal details with potential relevance to structure-based drug development. For example, the IF1-F1-ATPase interaction interface can be exploited by peptidomimetics to develop inhibitors selective for ATP hydrolytic activity of the enzyme. In the insect stage of T. brucei, the dimers of ATP synthase contribute to the shaping of the inner mitochondrial membrane by assembling into short helices on the rims of discoidal cristae (Muhleip et al., 2017). However, it is not clear how the dimers arrange in the BSF cells, which exhibit reduced complexity of mitochondrial ultrastructure. The understanding of the molecular mechanisms of membrane reshaping during the T. brucei life cycle, which can now be mimicked in vitro (Kolev et al., 2012; Dolezelova et al., 2020), will further illuminate the principles underlying mitochondrial dynamics in eukaryotes in general. Furthermore, recent data suggested interaction of the ATP synthase complex with mitochondrial calcium uniporter complex, AAC and phosphate carrier (Huang and Docampo, 2020). Further studies scrutinizing this putative megacomplex may reveal ATP synthase regulation by Ca2+ and strengthen the importance of the functional interplay between ATP synthase and both carriers in BSF and Dk cells. Considering our detailed knowledge of ATP synthase's role in T. brucei parasites, it is surprising that very little is known about its function in medically important parasites, Leishmania and T. cruzi. Although this gap is caused most likely by the fact that these parasites are less amenable to genetic manipulation, this can now be overcome by using subgenomic CRISPR/Cas9 libraries to dissect the role of oxidative phosphorylation complexes in different life cycle forms of these parasites. This will lead to a better understanding of the parasites' metabolic flexibility and capacity, ultimately advancing our efforts to target mitochondrial essential metabolic pathways for drug development.
Financial support
This study was supported by the Czech Science Foundation (18-17529S) and ERD fund and Ministry of Education, Youth and Sports (CZ.02.1.01/0.0/0.0/16_019/0000759).
Conflict of interest
The authors declare no competing interests.
References
- Abrahams JP, Leslie AG, Lutter R and Walker JE (1994) Structure at 2.8 Å resolution of F1-ATPase from bovine heart mitochondria. Nature 370, 621–628. [DOI] [PubMed] [Google Scholar]
- Agbe A and Yielding KL (1995) Kinetoplasts play an important role in the drug responses of Trypanosoma brucei. Journal of Parasitology 81, 968–973. [PubMed] [Google Scholar]
- Anselmi C, Davies KM and Faraldo-Gomez JD (2018) Mitochondrial ATP synthase dimers spontaneously associate due to a long-range membrane-induced force. The Journal of General Physiology 150, 763–770. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Arnold I, Pfeiffer K, Neupert W, Stuart RA and Schagger H (1998) Yeast mitochondrial F1Fo-ATP synthase exists as a dimer: identification of three dimer-specific subunits. The EMBO Journal 17, 7170–7178. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Asami K, Juniti K and Ernster L (1970) Possible regulatory function of a mitochondrial ATPase inhibitor in respiratory chain-linked energy transfer. Biochimica et Biophysica Acta 205, 307–311. [DOI] [PubMed] [Google Scholar]
- Atwood JA III, Weatherly DB, Minning TA, Bundy B, Cavola C, Opperdoes FR, Orlando R and Tarleton RL (2005). The Trypanosoma cruzi proteome. Science, 309, 473-476. [DOI] [PubMed] [Google Scholar]
- Bakker BM, Michels PA, Opperdoes FR and Westerhoff HV (1999) What controls glycolysis in bloodstream form Trypanosoma brucei? Journal of Biological Chemistry 274, 14551–14559. [DOI] [PubMed] [Google Scholar]
- Blum TB, Hahn A, Meier T, Davies KM and Kühlbrandt W (2019) Dimers of mitochondrial ATP synthase induce membrane curvature and self-assemble into rows. Proceedings of the National Academy of Sciences 116, 4250–4255. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Bowler MW, Montgomery MG, Leslie AG and Walker JE (2007) Ground state structure of F1-ATPase from bovine heart mitochondria at 1.9 Å resolution. Journal of Biological Chemistry 282, 14238–14242. [DOI] [PubMed] [Google Scholar]
- Brown SV, Hosking P, Li J and Williams N (2006) ATP synthase is responsible for maintaining mitochondrial membrane potential in bloodstream form Trypanosoma brucei. Eukaryotic Cell 5, 45–53. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Brun R, Hecker H and Lun ZR (1998) Trypanosoma evansi and T. equiperdum: distribution, biology, treatment and phylogenetic relationship (a review). Veterinary Parasitology 79, 95–107. [DOI] [PubMed] [Google Scholar]
- Bultema JB, Braun HP, Boekema EJ and Kouril R (2009) Megacomplex organization of the oxidative phosphorylation system by structural analysis of respiratory supercomplexes from potato. Biochimica et Biophysica Acta 1787, 60–67. [DOI] [PubMed] [Google Scholar]
- Burki F, Roger AJ, Brown MW and Simpson AGB (2020) The new tree of eukaryotes. Trends in Ecology & Evolution 35, 43–55. [DOI] [PubMed] [Google Scholar]
- Campanella M, Parker N, Tan CH, Hall AM and Duchen MR (2009) IF1: setting the pace of the F1Fo-ATP synthase. Trends in Biochemical Sciences 34, 343–350. [DOI] [PubMed] [Google Scholar]
- Cotter PD and Hill C (2003) Surviving the acid test: responses of gram-positive bacteria to low pH. Microbiology and Molecular Biology Reviews 67, 429–453. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Davies KM, Strauss M, Daum B, Kief JH, Osiewacz HD, Rycovska A, Zickermann V and Kuhlbrandt W (2011) Macromolecular organization of ATP synthase and complex I in whole mitochondria. Proceedings of the National Academy of Sciences 108, 14121–14126. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Davies KM, Anselmi C, Wittig I, Faraldo-Gomez JD and Kuhlbrandt W (2012) Structure of the yeast F1Fo-ATP synthase dimer and its role in shaping the mitochondrial cristae. Proceedings of the National Academy of Sciences 109, 13602–13607. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Dean S, Gould MK, Dewar CE and Schnaufer AC (2013) Single point mutations in ATP synthase compensate for mitochondrial genome loss in trypanosomes. Proceedings of the National Academy of Sciences 110, 14741–14746. [DOI] [PMC free article] [PubMed] [Google Scholar]
- de Souza W, de Carvalho TM and Barrias ES (2010) Review on Trypanosoma cruzi: host cell interaction. International Journal of Cell Biology 2010, 295394. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Dewar CE, MacGregor P, Cooper S, Gould MK, Matthews KR, Savill NJ and Schnaufer A (2018) Mitochondrial DNA is critical for longevity and metabolism of transmission stage Trypanosoma brucei. PLoS Pathogens 14, e1007195. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Dey R, Meneses C, Salotra P, Kamhawi S, Nakhasi HL and Duncan R (2010) Characterization of a Leishmania stage-specific mitochondrial membrane protein that enhances the activity of cytochrome c oxidase and its role in virulence. Molecular Microbiology 77, 399–414. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Dolezelova E, Kunzova M, Dejung M, Levin M, Panicucci B, Regnault C, Janzen CJ, Barrett MP, Butter F and Zikova A (2020) Cell-based and multi-omics profiling reveals dynamic metabolic repurposing of mitochondria to drive developmental progression of Trypanosoma brucei. PLoS Biology 18, e3000741. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Dudkina NV, Heinemeyer J, Keegstra W, Boekema EJ and Braun HP (2005) Structure of dimeric ATP synthase from mitochondria: an angular association of monomers induces the strong curvature of the inner membrane. FEBS Letters 579, 5769–5772. [DOI] [PubMed] [Google Scholar]
- Dudkina NV, Sunderhaus S, Braun HP and Boekema EJ (2006) Characterization of dimeric ATP synthase and cristae membrane ultrastructure from Saccharomyces and Polytomella mitochondria. FEBS Letters 580, 3427–3432. [DOI] [PubMed] [Google Scholar]
- Dudkina NV, Oostergetel GT, Lewejohann D, Braun HP and Boekema EJ (2010) Row-like organization of ATP synthase in intact mitochondria determined by cryo-electron tomography. Biochimica et Biophysica Acta 1797, 272–277. [DOI] [PubMed] [Google Scholar]
- Flygaard RK, Mühleip A, Tobiasson V and Amunts A (2020) Type III ATP synthase is a symmetry-deviated dimer that induces membrane curvature through tetramerization. Nature Communications 11, 1–11. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Formentini L, Sanchez-Arago M, Sanchez-Cenizo L and Cuezva JM (2012) The mitochondrial ATPase inhibitory factor 1 triggers a ROS-mediated retrograde prosurvival and proliferative response. Molecular Cell 45, 731–742. [DOI] [PubMed] [Google Scholar]
- Gahura O and Zikova A (2019) Isolation of F1-ATPase from the parasitic protist Trypanosoma brucei. Journal of Visualized Experiments 143, e58334. doi: 10.3791/58334 [DOI] [PubMed] [Google Scholar]
- Gahura O, Panicucci B, Vachova H, Walker JE and Zikova A (2018a) Inhibition of F1-ATPase from Trypanosoma brucei by its regulatory protein inhibitor TbIF1. The FEBS Journal 285, 4413–4423. [DOI] [PubMed] [Google Scholar]
- Gahura O, Subrtova K, Vachova H, Panicucci B, Fearnley IM, Harbour ME, Walker JE and Zikova A (2018b) The F1-ATPase from Trypanosoma brucei is elaborated by three copies of an additional p18-subunit. The FEBS Journal 285, 614–628. [DOI] [PubMed] [Google Scholar]
- Garcia-Bermudez J and Cuezva JM (2016) The ATPase inhibitory factor 1 (IF1): a master regulator of energy metabolism and of cell survival. Biochimica et Biophysica Acta 1857, 1167–1182. [DOI] [PubMed] [Google Scholar]
- Goes GR, Rocha PS, Diniz AR, Aguiar PH, Machado CR and Vieira LQ (2016) Trypanosoma cruzi needs a signal provided by reactive oxygen species to infect macrophages. PLoS Neglected Tropical Diseases 10, e0004555. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Goncalves RL, Barreto RF, Polycarpo CR, Gadelha FR, Castro SL and Oliveira MF (2011) A comparative assessment of mitochondrial function in epimastigotes and bloodstream trypomastigotes of Trypanosoma cruzi. Journal of Bioenergetics and Biomembranes 43, 651–661. [DOI] [PubMed] [Google Scholar]
- Gu J, Zhang L, Zong S, Guo R, Liu T, Yi J, Wang P, Zhuo W and Yang M (2019) Cryo-EM structure of the mammalian ATP synthase tetramer bound with inhibitory protein IF1. Science 364, 1068–1075. [DOI] [PubMed] [Google Scholar]
- Guo H, Bueler SA and Rubinstein JL (2017) Atomic model for the dimeric Fo region of mitochondrial ATP synthase. Science 358, 936–940. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Hahn A, Vonck J, Mills DJ, Meier T and Kuhlbrandt W (2018) Structure, mechanism, and regulation of the chloroplast ATP synthase. Science 360, eaat4318. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Harris DA, von Tscharner V and Radda GK (1979) The ATPase inhibitor protein in oxidative phosphorylation. The rate-limiting factor to phosphorylation in submitochondrial particles. Biochimica et Biophysica Acta 548, 72–84. [DOI] [PubMed] [Google Scholar]
- He J, Carroll J, Ding S, Fearnley IM and Walker JE (2017) Permeability transition in human mitochondria persists in the absence of peripheral stalk subunits of ATP synthase. Proceedings of the National Academy of Sciences 114, 9086–9091. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Hierro-Yap C, Subrtová K, Gahura O, Panicucci B, Dewar C, Chinopoulos C, Schnaufer A and Zikova A (2021) Bioenergetic consequences of FoF1-ATP synthase/ATPase deficiency in two life cycle stages of Trypanosoma brucei. Journal of Biological Chemistry 1, 100357. doi: 10.1016/j.jbc.2021.100357. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Hoare CABSCJ (1937) Spontaneous occurrence of strains of Trypanosoma evansi devoid of kinetonucleus. Parasitology 29, 43–56. [Google Scholar]
- Huang G and Docampo R (2020) The mitochondrial calcium uniporter interacts with subunit c of the ATP synthase of trypanosomes and humans. mBio 11, e00168–20. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Huet D, Rajendran E, van Dooren GG and Lourido S (2018) Identification of cryptic subunits from an apicomplexan ATP synthase. eLife 7, e38097. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Inbar E, Hughitt VK, Dillon LA, Ghosh K, El-Sayed NM and Sacks DL (2017) The transcriptome of Leishmania major developmental stages in their natural sand fly vector. mBio 8, e00029–17. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Jennings RB, Reimer KA and Steenbergen C (1991) Effect of inhibition of the mitochondrial ATPase on net myocardial ATP in total ischemia. Journal of Molecular and Cellular Cardiology 23, 1383–1395. [DOI] [PubMed] [Google Scholar]
- Jensen RE, Simpson L and Englund PT (2008) What happens when Trypanosoma brucei leaves Africa. Trends in Parasitology 24, 428–431. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Junge W and Nelson N (2015) ATP synthase. Annual Reviews of Biochemistry 84, 631–657. [DOI] [PubMed] [Google Scholar]
- Khare S, Roach SL, Barnes SW, Hoepfner D, Walker JR, Chatterjee AK, Neitz RJ, Arkin MR, McNamara CW, Ballard J, Lai Y, Fu Y, Molteni V, Yeh V, McKerrow JH, Glynne RJ and Supek F (2015) Utilizing chemical genomics to identify cytochrome b as a novel drug target for Chagas disease. PLoS Pathogens 11, e1005058. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Klodmann J, Senkler M, Rode C and Braun HP (2011) Defining the protein complex proteome of plant mitochondria. Plant Physiology 157, 587–598. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Kloehn J, Saunders EC, O'Callaghan S, Dagley MJ and McConville MJ (2015) Characterization of metabolically quiescent Leishmania parasites in murine lesions using heavy water labeling. PLoS Pathogens 11, e1004683. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Klusch N, Murphy BJ, Mills DJ, Yildiz O and Kuhlbrandt W (2017) Structural basis of proton translocation and force generation in mitochondrial ATP synthase. eLife 6, e33274. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Kolev NG, Ramey-Butler K, Cross GA, Ullu E and Tschudi C (2012) Developmental progression to infectivity in Trypanosoma brucei triggered by an RNA-binding protein. Science 338, 1352–1353. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Kuhlbrandt W (2019) Structure and mechanisms of F-type ATP synthases. Annual Reviews of Biochemistry 88, 515–549. [DOI] [PubMed] [Google Scholar]
- Lai DH, Hashimi H, Lun ZR, Ayala FJ and Lukes J (2008) Adaptations of Trypanosoma brucei to gradual loss of kinetoplast DNA: Trypanosoma equiperdum and Trypanosoma evansi are petite mutants of T. brucei. Proceedings of the National Academy of Sciences 105, 1999–2004. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Li Y, Shah-Simpson S, Okrah K, Belew AT, Choi J, Caradonna KL, Padmanabhan P, Ndegwa DM, Temanni MR, Corrada Bravo H, El-Sayed NM and Burleigh BA (2016) Transcriptome remodeling in Trypanosoma cruzi and human cells during intracellular infection. PLoS Pathogens 12, e1005511. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Luevano-Martinez LA, Girard R, Alencar MB and Silber AM (2020) ATP Regulates the activity of an alternative oxidase in Trypanosoma brucei. FEBS Letters 594, 2150–2158. [DOI] [PubMed] [Google Scholar]
- Lun ZR and Desser SS (1995) Is the broad range of hosts and geographical distribution of Trypanosoma evansi attributable to the loss of maxicircle kinetoplast DNA? Parasitology Today 11, 131–133. [DOI] [PubMed] [Google Scholar]
- Luque-Ortega JR, van't Hof W, Veerman EC, Saugar JM and Rivas L (2008) Human antimicrobial peptide histatin 5 is a cell-penetrating peptide targeting mitochondrial ATP synthesis in Leishmania. The FASEB Journal 22, 1817–1828. [DOI] [PubMed] [Google Scholar]
- McConville MJ, Saunders EC, Kloehn J and Dagley MJ (2015) Leishmania carbon metabolism in the macrophage phagolysosome- feast or famine? F1000Research 4, 938. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Montgomery MG, Gahura O, Leslie AGW, Zikova A and Walker JE (2018) ATP Synthase from Trypanosoma brucei has an elaborated canonical F1-domain and conventional catalytic sites. Proceedings of the National Academy of Sciences 115, 2102–2107. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Muhleip AW, Joos F, Wigge C, Frangakis AS, Kuhlbrandt W and Davies KM (2016) Helical arrays of U-shaped ATP synthase dimers form tubular cristae in ciliate mitochondria. Proceedings of the National Academy of Sciences 113, 8442–8447. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Muhleip AW, Dewar CE, Schnaufer A, Kuhlbrandt W and Davies KM (2017) In situ structure of trypanosomal ATP synthase dimer reveals a unique arrangement of catalytic subunits. Proceedings of the National Academy of Sciences 114, 992–997. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Muhleip A, McComas SE and Amunts A (2019) Structure of a mitochondrial ATP synthase with bound native cardiolipin. eLife 8, e51179. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Mühleip A, Kock Flygaard R, Ovciarikova J, Lacombe A, Fernandes P, Sheiner L and Amunts A (2021) ATP Synthase hexamer assemblies shape cristae of Toxoplasma mitochondria. Nature Communications 12, 1–13. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Murphy BJ, Klusch N, Langer J, Mills DJ, Yildiz O and Kuhlbrandt W (2019) Rotary substates of mitochondrial ATP synthase reveal the basis of flexible F1–Fo coupling. Science 364, eaaw9128. [DOI] [PubMed] [Google Scholar]
- Nelson RE, Aphasizheva I, Falick AM, Nebohacova M and Simpson L (2004) The I-complex in Leishmania tarentolae is an uniquely-structured F1-ATPase. Molecular and Biochemical Parasitology 135, 221–224. [DOI] [PubMed] [Google Scholar]
- Nolan DP and Voorheis HP (1992) The mitochondrion in bloodstream forms of Trypanosoma brucei is energized by the electrogenic pumping of protons catalysed by the F1Fo-ATPase. European Journal of Biochemistry 209, 207–216. [DOI] [PubMed] [Google Scholar]
- Opperdoes FR, Borst P and Spits H (1977) Particle-bound enzymes in the bloodstream form of Trypanosoma brucei. European Journal of Biochemistry 76, 21–28. [DOI] [PubMed] [Google Scholar]
- Paiva CN and Bozza MT (2014) Are reactive oxygen species always detrimental to pathogens? Antioxidants & Redox Signaling 20, 1000–1037. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Panek T, Elias M, Vancova M, Lukes J and Hashimi H (2020) Returning to the fold for lessons in mitochondrial Crista diversity and evolution. Current Biology 30, R575–R588. [DOI] [PubMed] [Google Scholar]
- Panicucci B, Gahura O and Zikova A (2017) Trypanosoma brucei TbIF1 inhibits the essential F1-ATPase in the infectious form of the parasite. PLoS Neglected Tropical Diseases 11, e0005552. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Perez E, Lapaille M, Degand H, Cilibrasi L, Villavicencio-Queijeiro A, Morsomme P, Gonzalez-Halphen D, Field MC, Remacle C, Baurain D and Cardol P (2014) The mitochondrial respiratory chain of the secondary green alga Euglena gracilis shares many additional subunits with parasitic Trypanosomatidae. Mitochondrion 19, 338–349. [DOI] [PubMed] [Google Scholar]
- Pinke G, Zhou L and Sazanov LA (2020) Cryo-EM structure of the entire mammalian F-type ATP synthase. Nature Structural & Molecular Biology 27, 1077–1085. [DOI] [PubMed] [Google Scholar]
- Preiss L, Langer JD, Yildiz O, Eckhardt-Strelau L, Guillemont JE, Koul A and Meier T (2015) Structure of the mycobacterial ATP synthase Fo rotor ring in complex with the anti-TB drug bedaquiline. Science Advances 1, e1500106. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Pullman ME and Monroy GC (1963) A naturally occurring inhibitor of mitochondrial adenosine triphosphatase. Journal of Biological Chemistry 238, 3762–3769. [PubMed] [Google Scholar]
- Ramrath DJF, Niemann M, Leibundgut M, Bieri P, Prange C, Horn EK, Leitner A, Boehringer D, Schneider A and Ban N (2018) Evolutionary shift toward protein-based architecture in trypanosomal mitochondrial ribosomes. Science 362, eaau7735. [DOI] [PubMed] [Google Scholar]
- Riou G and Benard J (1980) Berenil induces the complete loss of kinetoblast DNA sequences in Trypanosoma equiperdum. Biochemical and Biophysical Research Communications 96, 350–354. [DOI] [PubMed] [Google Scholar]
- Riou GF, Belnat P and Benard J (1980) Complete loss of kinetoplast DNA sequences induced by ethidium bromide or by acriflavine in Trypanosoma equiperdum. Journal of Biological Chemistry 255, 5141–5144. [PubMed] [Google Scholar]
- Rouslin W, Erickson JL and Solaro RJ (1986) Effects of oligomycin and acidosis on rates of ATP depletion in ischemic heart muscle. American Journal of Physiology 250, H503–H508. [DOI] [PubMed] [Google Scholar]
- Salunke R, Mourier T, Banerjee M, Pain A and Shanmugam D (2018) Highly diverged novel subunit composition of apicomplexan F-type ATP synthase identified from Toxoplasma gondii. PLoS Biology 16, e2006128. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Sathish Yadav KN, Miranda-Astudillo HV, Colina-Tenorio L, Bouillenne F, Degand H, Morsomme P, Gonzalez-Halphen D, Boekema EJ and Cardol P (2017) Atypical composition and structure of the mitochondrial dimeric ATP synthase from Euglena gracilis. Biochimica et Biophysica Acta 1858, 267–275. [DOI] [PubMed] [Google Scholar]
- Saunders EC and McConville MJ (2020) Immunometabolism of Leishmania granulomas. Immunology and Cell Biology 98, 832–844. [DOI] [PubMed] [Google Scholar]
- Saunders EC, Ng WW, Kloehn J, Chambers JM, Ng M and McConville MJ (2014) Induction of a stringent metabolic response in intracellular stages of Leishmania mexicana leads to increased dependence on mitochondrial metabolism. PLoS Pathogens 10, e1003888. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Schnaufer A, Domingo GJ and Stuart K (2002) Natural and induced dyskinetoplastic trypanosomatids: how to live without mitochondrial DNA. International Journal of Parasitology 32, 1071–1084. [DOI] [PubMed] [Google Scholar]
- Schnaufer A, Clark-Walker GD, Steinberg AG and Stuart K (2005) The F1-ATP synthase complex in bloodstream stage trypanosomes has an unusual and essential function. The EMBO Journal 24, 4029–4040. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Schwerzmann K and Pedersen PL (1981) Proton-adenosine triphosphatase complex of rat liver mitochondria: effect of energy state on its interaction with the adenosine triphosphatase inhibitory peptide. Biochemistry 20, 6305–6311. [DOI] [PubMed] [Google Scholar]
- Senkler J, Senkler M, Eubel H, Hildebrandt T, Lengwenus C, Schertl P, Schwarzlander M, Wagner S, Wittig I and Braun HP (2017) The mitochondrial complexome of Arabidopsis thaliana. The Plant Journal 89, 1079–1092. [DOI] [PubMed] [Google Scholar]
- Serricchio M, Hierro-Yap C, Schadeli D, Ben Hamidane H, Hemphill A, Graumann J, Zikova A and Butikofer P (2020) Depletion of cardiolipin induces major changes in energy metabolism in Trypanosoma brucei bloodstream forms. The FASEB Journal 1–16. [DOI] [PubMed] [Google Scholar]
- Shah-Simpson S, Pereira CF, Dumoulin PC, Caradonna KL and Burleigh BA (2016) Bioenergetic profiling of Trypanosoma cruzi life stages using seahorse extracellular flux technology. Molecular and Biochemical Parasitology 208, 91–95. [DOI] [PubMed] [Google Scholar]
- Silber AM, Tonelli RR, Lopes CG, Cunha-e-Silva N, Torrecilhas AC, Schumacher RI, Colli W and Alves MJ (2009) Glucose uptake in the mammalian stages of Trypanosoma cruzi. Molecular and Biochemical Parasitology 168, 102–108. [DOI] [PubMed] [Google Scholar]
- Silva Pereira S, Trindade S, De Niz M and Figueiredo LM (2019) Tissue tropism in parasitic diseases. Open Biology 9, 190036. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Skodova-Sverakova I, Horvath A and Maslov DA (2015) Identification of the mitochondrially encoded subunit 6 of F1Fo ATPase in Trypanosoma brucei. Molecular and Biochemical Parasitology 201, 135–138. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Sobti M, Ishmukhametov R, Bouwer JC, Ayer A, Suarna C, Smith NJ, Christie M, Stocker R, Duncan TM and Stewart AG (2019) Cryo-EM reveals distinct conformations of E. coli ATP synthase on exposure to ATP. eLife 8, e43864. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Speijer D, Breek CK, Muijsers AO, Hartog AF, Berden JA, Albracht SP, Samyn B, Van Beeumen J and Benne R (1997) Characterization of the respiratory chain from cultured Crithidia fasciculata. Molecular and Biochemical Parasitology 85, 171–186. [DOI] [PubMed] [Google Scholar]
- Spikes TE, Montgomery MG and Walker JE (2020) Structure of the dimeric ATP synthase from bovine mitochondria. Proceedings of the National Academy of Sciences 117, 23519–23526. [DOI] [PMC free article] [PubMed] [Google Scholar]
- St-Pierre J, Brand MD and Boutilier RG (2000) Mitochondria as ATP consumers: cellular treason in anoxia. Proceedings of the National Academy of Sciences 97, 8670–8674. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Stuart KD (1971) Evidence for the retention of kinetoplast DNA in an acriflavine-induced dyskinetoplastic strain of Trypanosoma brucei which replicates the altered central element of the kinetoplast. Journal of Cell Biology 49, 189–195. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Stuart K (1983) Mitochondrial DNA of an African trypanosome. Journal of Cellular Biochemistry 23, 13–26. [DOI] [PubMed] [Google Scholar]
- Subramanian A, Jhawar J and Sarkar RR (2015) Dissecting Leishmania infantum energy metabolism – a systems perspective. PLoS ONE 10, e0137976. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Subrtova K, Panicucci B and Zikova A (2015) ATPasetb2, a unique membrane-bound FoF1-ATPase component, is essential in bloodstream and dyskinetoplastic trypanosomes. PLoS Pathogens 11, e1004660. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Timms MW, van Deursen FJ, Hendriks EF and Matthews KR (2002) Mitochondrial development during life cycle differentiation of African trypanosomes: evidence for a kinetoplast-dependent differentiation control point. Molecular Biology of the Cell 13, 3747–3759. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Tobie EJ (1951) Loss of the kinetoplast in a strain of Trypanosoma equiperdum. Transactions of the American Microscopical Society 70, 251–254. [Google Scholar]
- Vazquez-Acevedo M, Cardol P, Cano-Estrada A, Lapaille M, Remacle C and Gonzalez-Halphen D (2006) The mitochondrial ATP synthase of chlorophycean algae contains eight subunits of unknown origin involved in the formation of an atypical stator-stalk and in the dimerization of the complex. Journal of Bioenergetics and Biomembranes 38, 271–282. [DOI] [PubMed] [Google Scholar]
- Walker JE (1994) The regulation of catalysis in ATP synthase. Current Opinion in Structural Biology 4, 912–918. [DOI] [PubMed] [Google Scholar]
- Walker JE (2017) Structure, mechanism and regulation of ATP synthases. In Wikstrom M (ed.), Mechanisms of Primary Energy Transduction in Biology. Cambridge, UK: The Royal Society of Chemistry, pp. 338–373. [Google Scholar]
- Waterhouse A, Bertoni M, Bienert S, Studer G, Tauriello G, Gumienny R, Heer FT, de Beer TAP, Rempfer C, Bordoli L, Lepore R and Schwede T (2018) Swiss-model: homology modelling of protein structures and complexes. Nucleic Acids Research 46, W296–W303. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Wittig I, Meyer B, Heide H, Steger M, Bleier L, Wumaier Z, Karas M and Schagger H (2010) Assembly and oligomerization of human ATP synthase lacking mitochondrial subunits a and A6L. Biochimica et Biophysica Acta 1797, 1004–1011. [DOI] [PubMed] [Google Scholar]
- Zikova A, Schnaufer A, Dalley RA, Panigrahi AK and Stuart KD (2009) The FoF1-ATP synthase complex contains novel subunits and is essential for procyclic Trypanosoma brucei. PLoS Pathogens 5, e1000436. [DOI] [PMC free article] [PubMed] [Google Scholar]
- Zikova A, Verner Z, Nenarokova A, Michels PAM and Lukes J (2017) A paradigm shift: the mitoproteomes of procyclic and bloodstream Trypanosoma brucei are comparably complex. PLoS Pathogens 13, e1006679. [DOI] [PMC free article] [PubMed] [Google Scholar]