ATP Synthase Subunit a Supports Permeability Transition in Yeast Lacking Dimerization Subunits and Modulates yPTP Conductance
Katarzyna Niedzwiecka Emilia Baranowska Chiranjit Panja Roza Kucharczyk
Institute of Biochemistry and Biophysics, Polish Academy of Sciences, Warsaw, Poland
Key Words
Mitochondria • Permeability transition pore • ATP synthase • ATP6 • Subunit a • S. cerevisiae
Abstract
Background/Aims: Mitochondrial ATP synthase, in addition to being involved in ATP synthesis, is involved in permeability transition pore (PTP) formation, which precedes apoptosis in mammalian cells and programmed cell death in yeast. Mutations in genes encoding ATP synthase subunits cause neuromuscular disorders and have been identified in cancer samples. PTP is also involved in pathology. We previously found that in Saccharomyces cerevisiae, two mutations in ATP synthase subunit a (atp6-P163S and atp6-K90E, equivalent to those detected in prostate and thyroid cancer samples, respectively) in the OM45-GFP background affected ROS and calcium homeostasis and delayed yeast PTP (yPTP) induction upon calcium treatment by modulating the dynamics of ATP synthase dimer/oligomer formation. The Om45 protein is a component of the porin complex, which is equivalent to mammalian VDAC. We aimed to investigate yPTP function in atp6-P163S and atp6-K90E mutants lacking the e and g dimerization subunits of ATP synthase. Methods: Triple mutants with the atp6-P163S or atp6-K90E mutation, the OM45-GFP gene and deletion of the TIM11 gene encoding subunit e were constructed by crossing and tetrad dissection. In spores capable of growing, the original atp6 mutations reverted to wild type, and two compensatory mutations, namely, atp6-C33S-T215C, were selected. The effects of these mutations on cellular physiology, mitochondrial morphology, bioenergetics and permeability transition (PT) were analyzed by fluorescence and electron microscopy, mitochondrial respiration, ATP synthase activity, calcium retention capacity and swelling assays. Results: The atp6-C33S-T215C mutations in the OM45-GFP background led to delayed growth at elevated temperature on both fermentative and respiratory media and increased sensitivity to high calcium ions concentration or hydrogen peroxide in the medium. The ATP synthase activity was reduced by approximately 50% and mitochondrial network was hyperfused in these cells grown at elevated temperature. The atp6-C33S-T215C stabilized ATP synthase dimers and restored the yPTP properties in Tim11∆ cells. In OM45-GFP cells, in which Tim11 is present, these mutations increased the fraction of swollen mitochondria by up to 85% vs 60% in the wild type, although the time required for calcium release doubled. Conclusion: ATP synthase subunit e is essential in the S. cerevisiae atp6-P163S and atp6-K90E mutants. In addition to subunits e and g, subunit a is critical for yPTP induction and conduction. The increased yPTP conduction decrease the S. cerevisiae cell fitness.
Introduction
Mitochondrial ATP synthase, an enzyme that provides cellular energy in the form of ATP, is composed of 17 subunits [1]. The intersubunit interactions are critical for the catalytic activity of the enzyme. These subunits form two main enzyme domains: the hydrophobic FO domain that embeds the enzyme in the inner mitochondrial membrane (IMM) and the catalytic F1 domain that is exposed in the matrix [2]. ATP synthase uses a proton gradient established by the respiratory chain across the IMM. The higher concentration of protons in the intermembrane space (IMS) than in the matrix enables ATP synthase to produce ATP from ADP and inorganic phosphate [3]. Protons pass through the channel in the FO domain formed between the c-ring (consisting of ten c subunits in Saccharomyces cerevisiae) and the hydrophilic amino acids of subunit a (which is tightly attached to the c-ring), driving the rotation of the ring. The proton enters from the IMS into the entry half-channel. It binds to an acidic residue (cE58 in S. cerevisiae) in the second helix of subunit c and, after an almost complete rotation of the ring, is liberated to the exit half-channel on the matrix side [4, 5]. The c-ring is tightly bound to the central stalk subunits γ, δ, and ε, of which subunit γ protrudes into the F1 catalytic domain (composed of a hexamer of α and β subunits). Subunit γ rotates with the ring and induces cyclic conformational changes in F1 [6, 7]. Consequently, ADP and Pi are sequentially converted to ATP at the catalytic sites of β subunits, according to the binding change mechanism. In addition to subunits a and c, the FO domain consists of hydrophobic subunits 8, i/j, and f and the membrane part of subunit b. The hydrophilic part of b protrudes into the matrix, interacts with the d, h, and OSCP (oligomycin sensitivity-conferring protein) subunits and forms the external stalk that connects FO with F1. Three additional subunits, namely, e, g, and k, stabilize the ATP synthase dimers [8] that self-assemble in longer ribbons that are important for cristae formation [9, 10].
In addition to its main function in ATP production, IMM curvature and the formation of the mitochondrial network [11], ATP synthase has been proposed to be directly involved in the permeability transition (PT) [12-14]. PT is a Ca2+-dependent increase in IMM permeability to solutes and proteins with an exclusion limit of approximately 1500 Da due to the formation of the unselective high-conductance permeability transition pore (PTP), also called the mitochondrial megachannel (MMC) [15]. The PTP can open for a short duration and assume a low conductance, probably contributing to Ca2+ homeostasis and ROS signaling [16, 17], or can open for a long period with a high conductance, contributing to cell death [15]. A molecular explanation for this complex behavior of PTP, both in regulation and channel composition, has not been provided. Understanding PTP function and regulation is a major goal for scientists performing basic research and for clinicians because its dysregulation is implicated in pathological conditions such as neurodegeneration, cancer, cardiac hypertrophy and ischemia [18, 19]. The molecular composition of the channel remains a subject of debate [20, 21]. ATP synthase dimers or tetramers (but not monomers) eluted from native gels and reconstituted into liposomes form channels characterized by different conductances depending on the model organism [14, 22-24]. Similar experiments performed on the purified enzyme embedded in vesicles showed high-conductance channel activity for the tetramers, the dimers and the monomers [25, 26]. It was also proposed that PTP opening requires the dissociation of ATP synthase dimers [27]. Well characterized modulators of the PTP have been identified to bind to and act on subunits of ATP synthase, further corroborating its role as the molecular identity of the PTP. The divalent Mg2+, Mn2+, Sr2+ or Ba2+ cations have inhibitory effects on the PTP because they compete with Ca2+ to bind the catalytic site in the β subunit [28]. The specific PTP inducer Bz423 and the PTP regulator protein CyPD bind to OSCP, and at the same time, the inhibitory effect of matrix H+ at a pH of 6.5 is achieved through the protonation of the unique histidine in the OSCP subunit [14, 29, 30]. Importantly, it was recently shown that deletion of the c subunit prevents the assembly of a functional ATP synthase [20]. This leads to loss of the PTP in mammalian cells, providing a strong argument in favor of PTP formation by ATP synthase or its specific subunits [27, 31].
Experiments in S. cerevisiae have also provided data supporting the role of ATP synthase in the formation of the yeast PTP (yPTP, also called the yeast mitochondrial unspecific channel, YMUC). Importantly, the features of the yPTP are similar to those of the mammalian PTP, i.e., regulation by calcium, ROS, and matrix cyclophilin D and involvement in cell death [22, 32, 33]. A unique feature of yeast ATP synthase is the presence of a cysteine at position 23 of the a subunit (Fig. 1). The disulfide bond formed between the cysteine residues of two neighboring a subunits stabilizes the ATP synthase dimer even in the absence of subunits e, g and k, as shown by BN-PAGE analysis of ATP synthase complexes isolated from mitochondria incubated with Cu2+ [34, 35]. The mitochondria of S. cerevisiae cells lacking subunits e and g are desensitized to calcium but are able to form a long-lasting channel with 10 times lower conductance than that of the wild-type channel. This permits calcium release but not swelling, which requires the high-conductance channel to be formed [22, 32, 35]. The first transmembrane domain of subunit b also plays a role in the stabilization of the open state of the yPTP [35].
We have previously shown that mutations in the mitochondrial ATP6 gene of S. cerevisiae, namely, atp6-P163S and atp6-K90E, leading to the substitution of proline 153 for serine or lysine 80 for glutamic acid in subunit a, respectively (positions in mature subunit a after the removal of the first ten amino acids in the presequence by the protease Atp23 [36]), desensitized the yPTP to calcium in mitochondria bearing the Om45 protein fused to GFP. Importantly, these two atp6 mutations were identified in prostate and thyroid cancer samples and may contribute to cell immortalization by facilitating escape from apoptosis [37]. The Om45 protein forms a complex with porin and interacts physically with the components of the IMM: transporters, adenine nucleotides carrier, respiratory chain complexes III and IV and ATP synthase [38]. It was proposed that this protein coordinates transport across mitochondrial membranes. In this study, we asked whether the atp6-P163S or atp6-K90E mutations influence the induction of the yPTP in mitochondria defective in ATP synthase dimerization, i.e., lacking subunits e and g. Surprisingly, we found that subunit e is essential in the atp6-P163S and atp6-K90E mutants. We show that subunit a of ATP synthase plays a role in yPTP induction and conduction and provide the first in vivo evidence that yPTP function is important for the fitness of S. cerevisiae cells.
Materials and Methods
Yeast strains
The S. cerevisiae strains used in this study were all isogenic derivatives of MR6 and are listed in Table 1. ρ+ indicates the complete wild-type mitochondrial genome (mtDNA; when followed by a mutation, it indicates the complete mtDNA with a single introduced mutation), and ρ0 indicates the strain lacking mitochondrial DNA. The RKY62α and RKY61α strains were obtained by changing the mating type using a plasmid encoding the HO endonuclease [39]. The KNY58 strain was constructed by integrating the tim11Δ::HphNTI cassette into the TIM11 locus of the MR6 strain. The cassette was amplified using the pMW77/11 plasmid as a template [40] with the following primers: 5′CACAGAAATTTTAGATAAAAGGAAGTATTATATCGGAACATAACGTATATAGGAA CTAGCGAGTGAGTTAAAGGATGGGTAAAAAGCCTGAACTCACCG3′ and 5′ATCCATCATAACTTCGTCATTCAGTG CGAGCTAATGTGCATTTTTAGTATCCTATTTATGTTGAAGCTTCTATTTTATTCCTTTGCCCTCGGACGAGTGC3′. The KNY67 strain was obtained by fusing GFP to the 3′ end of OM45 in its genomic locus in the KNY58 strain, according to [41]. Thus, all strains expressed Om45-GFP under the control of the OM45 gene promoter. The KNY58, KNY67 and MR6-OM45GFP strains were depleted of mtDNA by culturing them in liquid YPGalA medium at 37 °C and selecting respiration-deficient clones, yielding KNY59, KNY67-1 and KNY40, respectively. The absence of mtDNA in these clones was confirmed by crossing with the RKY25 MATα (a [ρ+] strain bearing a point mutation in the ATP6 gene that confers respiratory deficiency [42]). The diploid strains KNY68 and KNY70 were obtained by mating RKY61α with KNY59 and KNY67-1, respectively. The diploid strains KNY68 and KNY71 were obtained by mating RKY62α with KNY59 and KNY67-1, respectively. The KNY76-1 strain was obtained by sporulation and tetrad dissection of KNY68. The KNY88 diploid strain was obtained by mating KNY76-1 with KNY40. The KNY89-1 and KNY91-1 strains were obtained by sporulation of the KNY88 strain and subsequent tetrad dissection. The KNY103-1 strain was obtained by fusing GFP to the OM45 gene in the KNY76-1 strain [41]. The absence of the Tim11 protein and the presence of the GFP tag at the C-terminus of the Om45 protein were confirmed by Western blot analysis using an anti-Tim11 antibody and GFP fluorescence, respectively. The sequence of the ATP6 gene was verified by sequencing in all haploid strains with oAtp6-1 5′TAATATACGGGGGTGGGTCCCTCAC3′ and oATP6-10 5′GGGCCGAACTCCGAAGGAGTAAG3′, which were also used for ATP6 gene amplification.
Growth conditions
Strains were grown in rich YPGA medium (1% Bacto yeast extract, 1% Bacto peptone, 2% glucose, 40 mg/l adenine) at 28 or 36 °C with shaking at 200 rpm. For mitochondrial isolation, strains were grown in rich YPGalA medium (1% Bacto yeast extract, 1% Bacto peptone, 2% galactose, 40 mg/l adenine) to an OD600 of 4. Respiratory YPGlyA medium contained 1% Bacto yeast extract, 1% Bacto peptone, 2% glycerol, and 40 mg/l adenine (Difco, Becton Dickinson). W0 complete minimal medium contained 6.7% yeast nitrogen base w/o amino acids and 2% galactose, supplemented with appropriate drop-out amino acid stock (Sunrise) for plasmid selection. The liquid media were solidified by addition of 2% Bacto agar (Difco, Becton Dickinson). An OD600 of 1 corresponds to 1.2 x 107 cells/ml for the MR6 strain. Geneticin or hygromycin were added to YPGA at a concentration of 200 µg/ml or 300 µg/ml, respectively.
Measurement of oxygen consumption and ATP synthase activity
Mitochondria were isolated from cells grown in YPGalA medium at 28 °C according to the protocol described in reference [43]. Oxygen consumption rates were measured using a Clarke electrode in respiration buffer (0.65 M mannitol, 0.36 mM EGTA, 5 mM Tris-phosphate, 10 mM Tris-maleate pH=6.8) after consecutively adding 75 µg of mitochondrial proteins, 4 mM NADH (state 4 respiration), 150 µM ADP (state 3) or 4 µM carbonyl cyanide m-chlorophenylhydrazone (CCCP) (uncoupled respiration), as previously described [44]. The rates of ATP synthesis were determined under the same experimental conditions with 750 µM ADP; aliquots were withdrawn from the oxygraph cuvette every 15 seconds, and the reaction was stopped with 3.5% (w/v) perchloric acid and 12.5 mM EDTA. The samples were then neutralized to pH 6.5 by the addition of KOH and 0.3 M MOPS. The synthetized ATP was quantified using a luciferin/luciferase assay (Kinase-Glo Max Luminescence Kinase Assay, Promega) in a Beckman Coulter Paradigm plate reader. The participation of F1FO-ATP synthase in ATP production was assessed by measuring the sensitivity of ATP synthesis to oligomycin (3 μg/ml). The specific ATPase activity at pH 8.4 of nonosmotically protected mitochondria was measured using the procedure described by M. Somlo [45].
Measurement of the mitochondrial calcium retention capacity and swelling
A previously developed method for measurement of mitochondrial calcium retention capacity (described in detail in [37]) was used to measure the time of yPTP opening after Ca2+ addition. Briefly, mitochondria were diluted in CRC buffer (250 mM sucrose, 10 mM Tris-MOPS, 10 µM EGTA-Tris, 5 mM Pi-Tris, 1 mM NADH, 5 µM ETH129, 1 µM Calcium Green-5N (Thermo Fisher), 0.5 mg/ml BSA, pH 7.4) to a concentration of 500 µg/ml, and 50 µM CaCl2 was added. The rapid increase in the fluorescence of Calcium Green-5N was as attributed to the release of calcium ions from the mitochondrial matrix into the buffer, likely due to the PT. Matrix swelling was evaluated by measuring optical density changes at 540 nm with a Varian Cary Bio UV-Vis spectrophotometer. Mitochondria (500 µg/ml) were suspended in 2 ml of CRC buffer without Calcium Green-5N, and then, 140 µM CaCl2 and 10 µM alamethicin were added.
BN-PAGE, SDS-PAGE and Western blotting
These methods have been described previously [37]. For cross-linking experiments, mitochondria were incubated for 20 or 30 minutes at room temperature at 1 mg/ml in 250 mM sucrose, 5 mM Pi and 2 mM CuCl2. Then, 5 mM N-ethylmaleimide and 5 mM EDTA were added to block the cross-linking reaction, and the incubations were transferred to ice for 10 minutes and centrifuged. Then, the ATP synthase complexes were liberated from IMM by incubation with 1.5% digitonin in extraction buffer (30 mM HEPES, 150 mM potassium acetate, 12% glycerol, 2 mM 6-aminocaproic acid, 1 mM EGTA, protease inhibitor cocktail tablets (Roche), pH 7.4) for 20 minutes and separated by BN-PAGE. Polyclonal rabbit anti-ATP synthase subunit antibodies (a gift from Marie-France Giraud, Bordeaux, France) were used.
Morphology of the mitochondria and mitochondrial network
The mitochondrial network was labelled with RFP using the pXY142-RFP plasmid encoding RFP fused to the mitochondrial targeting sequence [46]. Cells were grown at 28 or 36 °C in W0-galactose medium lacking leucine to an OD of 1 and used directly for fluorescence microscopy. The morphology of the mitochondria was assessed in whole cells grown in YPGalA medium at 28 or 36 °C to an OD of 1 and processed for electron microscopy as described previously [47]. Briefly, cells grown in YPGalA medium were fixed by incubation overnight in 2% paraformaldehyde + 2. 5% glutaraldehyde in cacodylate buffer (pH 7.4). Then, the cells were rinsed in cacodylate buffer and deionized water. Cells were immersed in 6% KMnO4 water solution for 1 hour at room temperature, rinsed 5 times in water and dehydrated in an ethanol gradient (30-100%). Then, the material was embedded in epoxy resin (Epon 812). Thin 50-nm sections were stained with 9% uranyl acetate and lead nitrate. Images were acquired using a JEM-1200 EX (Jeol, Japan) transmission electron microscope (80 keV) equipped with a MORADA camera and iTEM 1233 software.
Statistical analyses
Unless otherwise stated in the figure legends, each experiment was repeated at least three times. The intensity of the bands was quantified using ImageJ. Data are presented as the average ± s.d. or as a representative experiment. Student's t-test was used to assess significant differences with the respective control.
Results
Tim11 is essential in atp6 mutants
Our previous study indicated that the atp6-P163S or atp6-K90E mutation in the OM45-GFP background in S. cerevisiae conferred sensitivity to high calcium concentrations or hydrogen peroxide in the medium. The yPTP was desensitized to calcium in these double mutants [37]. The time of yPTP induction by calcium was longer in atp6-P163S OM45-GFP and atp6-K90E OM45-GFP mitochondria (grown at 36 °C) than in control mitochondria and comparable to the induction time in tim11∆ mitochondria lacking ATP synthase dimers [37]. Assuming that the hypothesis regarding the formation of the PTP core by ATP synthase dimers is true, we expected to observe an effect of the atp6-P163S or atp6-K90E mutation on yPTP induction in tim11∆ cells. Deletion of the TIM11 gene not only leads to a lack of subunit e but also causes a lack of the g subunit (Supplementary Fig. S1, [22] – for all supplemental material see www.cellphysiolbiochem.com). Efforts to delete the TIM11 gene by one-step integration of the tim11::HphNTI cassette into the TIM11 locus of single atp6-P163S, atp6-K90E, and OM45-GFP mutants and double atp6-P163S OM45-GFP and atp6-K90E OM45-GFP mutants were ineffective. Using this method, we replaced the coding sequence of TIM11 with the tim11::HphNTI cassette in a strain bearing the wild-type ATP6 gene only. We hypothesized that Tim11 is essential in the single mutants atp6-P163S, atp6-K90E, and OM45-GFP and double mutants atp6-P163S OM45-GFP and atp6-K90E OM45-GFP. To verify this hypothesis, we aimed to introduce the tim11::HphNTI deletion into the above mutants by the classical genetic method of diploid sporulation and tetrad dissection. The tim11::HphNTI [ρ0] strain was crossed with the single mutants atp6-P163S and atp6-K90E and double mutants atp6-P163S OM45-GFP and atp6-K90E OM45-GFP; the obtained diploids were sporulated, and tetrads were dissected. Despite very efficient sporulation, only a few spores were able to grow on plates with twenty dissected tetrads (at least forty tetrads were analyzed per crossing (Supplementary Fig. S2)).
The presence of the tim11::HphNTI deletion or Om45-GFP in spores was analyzed by testing S. cerevisiae growth on hygromycin- or geneticin-containing plates and by evaluating GFP fluorescence. The ATP6 gene was entirely sequenced. Surprisingly, in every spore, independent of the presence of the tim11::HphNTI deletion or Om45-GFP, the TCA codon for Atp6-S153 and the GAA codon for Atp6-E80 reverted to the wild-type CCA and AAA codons for proline and lysine, respectively, but two additional mutations appeared, leading to the substitution of the cysteine in position 23 of mature Atp6p with a serine and the threonine in position 205 with a cysteine (Atp6-C23S-T205C, designated Atp6-CSTC) (Fig. 1). Although we were not able to delete the TIM11 gene by one-step transformation in the OM45-GFP strain, OM45 was fused to GFP as described in [41] in tim11::HphNTI and tim11::HphNTI atp6-CSTC mutants. The lack of subunit e or g in the analyzed strains was confirmed by Western blotting with anti-Tim11 or anti-Atp20 (Supplementary Fig. S1). We therefore concluded that the Tim11 subunit, which is nonessential for the growth of S. cerevisiae cells even in respiratory medium, is essential in cells expressing the Atp6-P153S, Atp6-K80E, Atp6-P153S and Om45-GFP or Atp6-K80E and Om45-GFP proteins. This protein nomenclature will be used throughout the remainder of this paper.
Effect of Atp6-CSTC on growth phenotypes, mitochondrial DNA stability, the mitochondrial network and morphology
We first established the impact of Atp6-CSTC substitutions on S. cerevisiae growth, the stability of the mtDNA, and the morphology of the mitochondria and the mitochondrial network. Atp6-CSTC did not disturb the growth of S. cerevisiae cells, but in the Om45-GFP background, it led to delayed growth at elevated temperature on both fermentative and respiratory media (Fig. 2A). The Atp6-P153S or Atp6-K80E substitution in Om45-GFP led to increased sensitivity to high calcium ions concentration or hydrogen peroxide in the medium [37]. Atp6-CSTC Om45-GFP cells were also hypersensitive to calcium ions and hydrogen peroxide (Fig. 2A). These growth phenotypes of Atp6-CSTC Om45-GFP cells were suppressed by deletion of subunit e.
Tim11, due to its role in the dimerization of ATP synthase, is critical for cristae invaginations [9, 10]. Thus, we examined the mitochondrial network and morphology in vivo by fluorescence and electron microscopy, respectively. The mitochondria of cells lacking Tim11 did not form cristae. However, their inner membrane formed onion-like structures, which was the phenotype observed in cells grown at normal and elevated temperatures. Additionally, the mitochondrial network in Tim11∆ cells grown at 36 °C was not tubular, reaching all cell surface, but accumulated on one or two poles of a cell, forming long tubules close to the cell membrane (Fig. 3A; at 28 °C, this phenotype is visible only in single cells) [48, 49]. Atp6-CSTC substitutions alone did not influence the mitochondrial network but restored its defect in Tim11Δ cells (also in the Om45-GFP background; Fig. 3A). However, Atp6-CSTC substitutions did not restore the onion-like cristae morphology of mitochondria caused by Tim11 deletion, and 100% of the mitochondria of cells lacking subunit e, regardless of CSTC substitutions in subunit a and the presence of Om45-GFP, had an aberrant cristae morphology (Fig. 3B). Tim11∆ cells were also genetically unstable, with up to 60% of cells grown to late logarithmic phase at permissive temperature losing mtDNA in comparison to 3% of cells of the wild-type strain. The mtDNA instability of Tim11∆ was also partially suppressed by Atp6-CSTC substitutions (Fig. 2B). Interestingly, while the cristae morphology was normal in the Atp6-CSTC Om45-GFP double mutant, the mitochondrial network, which was normal at a permissive temperature, was hyperfused during growth at elevated temperature (Fig. 3A, B) and exhibited a phenotype much stronger than that observed in the Tim11Δ cells. Strong defects in the mitochondrial network, together with increased ROS levels and deregulation of calcium homeostasis, may contribute to the temperature sensitivity phenotype of Atp6-CSTC Om45-GFP cells.
Atp6-CSTC restored respiration and ATP synthase activity in Tim11∆ cells
Normal respiratory growth in S. cerevisiae does not indicate that OXPHOS is as efficient as it is in wild-type cells. Respiratory growth defects appear only when ATP synthase activity decreases below 20% of the wild-type level [50, 51]. To explain the temperature-sensitive phenotype of the Atp6-CSTC Om45-GFP strain, we measured oxygen consumption and ATP synthesis. Mitochondria were isolated from all strains grown only at 28 °C, as culturing the Tim11∆ strain at 36 °C resulted in more than 70% petite cells. The oxygen consumption and ATP synthesis of Tim11∆ mitochondria were reduced by approximately 45 and 55%, respectively, in comparison to those of the control mitochondria, regardless of the presence of Om45-GFP (Fig. 4). Interestingly, the defect in the oxidative phosphorylation in mitochondria lacking Tim11 was suppressed by Atp6-CSTC, in both the wild-type and Om45-GFP backgrounds. Atp6-CSTC substitution alone did not influence oxygen consumption and ATP synthesis, but in combination with Om45-GFP, it reduced these activities to a level comparable to that in Tim11∆ mitochondria. Then, we measured the maximal ATPase activity in nonosmotically protected mitochondria buffered at pH 8.4 and in the presence of saturating amounts of ATP. A pH value of 8.4 is optimal for F1-ATPase activity and prevents the binding of the F1 inhibitor protein Inh1 (IF1) to ATP synthase [52]. In all strains, the maximal ATPase activity was the same (Fig. 4, gray bars), but the oligomycin-sensitive ATPase activity varied. In the mitochondria of most strains, approximately 85% of ATPase activity was sensitive to oligomycin. However, in the single mutant Tim11Δ and double mutant Tim11Δ Om45-GFP mitochondria, the sensitivity to oligomycin decreased to approximately 60%. This indicates the reduced stability of the ATP synthase complexes. We concluded that Atp6-CSTC substitutions alone had no effect on oxygen consumption, ATP synthesis and ATP hydrolysis but restored these activities to the wild-type level in mitochondria lacking Tim11. When subunit e was present in the Om45-GFP background, Atp6-CSTC substitutions decreased oxygen consumption and ATP synthesis by approximately 50%. Decreased ATP synthesis may also contribute to the temperature-sensitive phenotype of Atp6-CSTC Om45-GFP cells.
Atp6-CSTC stabilized ATP synthase dimers but not by disulfide bond formation between a subunits
Subunit e is critical for the dimerization of ATP synthase. In the absence of this subunit, the amount of fully assembled ATP synthase was normal (as assessed here based on the amount of subunit a, which is immediately degraded when not assembled into the enzyme, Supplementary Fig. S1, [42, 53, 54]). However, these complexes were exclusively found in the monomeric form. The monomer of the enzyme lacking subunit e exhibited reduced stability [48], as indicated by the presence of free F1 in the BN-PAGE analysis of the enzyme complexes extracted from the IMM (Fig. 5, left panel) and by the increase in ATPase activity in the presence of oligomycin (Fig. 4). Oligomycin inhibits only the coupled enzyme; therefore, oligomycin-insensitive ATP hydrolysis is carried out by free F1 [42]. The Atp6-CSTC substitutions increased the stability of ATP synthase dimers in Tim11∆ cells (Fig. 5A, left panel) and partially restored the oligomycin-sensitive ATPase activity (Fig. 4).
To further study the impact of Atp6-CSTC substitutions on ATP synthase dimerization, we performed an analysis of ATP synthase complexes by BN-PAGE after Cu2+ treatment, which stabilized the dimers in ∆e/∆g cells [22, 34, 35, 55]. Copper caused the cross-linking and stabilization of ATP synthase dimers in wild-type and Tim11∆ mitochondria (also in the Om45-GFP background) but not in those bearing Atp6-CSTC substitutions. This suggests that the cysteine at position 205 enables copper-dependent dimer stabilisation (Fig. 5, right panel). Thus, Atp6-CSTC substitutions stabilized ATP synthase dimers in cells lacking subunit e but not by disulfide bond formation between neighboring a subunits in these dimers.
Atp6-CSTC substitutions compensated for the lack of subunit e during yPTP induction
The absence of subunit e (and of subunit g as a consequence, Supplementary Fig. S1) decreased Ca2+ sensitivity and the conductance of the yPTP [22, 35]. We previously determined that subunit a mutations leading to Atp6-P153S or Atp6-K80E substitutions desensitized the yPTP to calcium in Om45-GFP mitochondria. This finding correlated with the sensitivity of cells to high calcium and hydrogen peroxide in the medium [37]. Atp6-CSTC Om45-GFP cells were also hypersensitive to calcium ions and hydrogen peroxide, suggesting that these substitutions may be significant for PT (Fig. 2A). We then asked how Atp6-CSTC substitutions influence the yPTP, especially in Tim11∆ and Om45-GFP mitochondria. For this purpose, we analyzed the rate of yPTP opening after high calcium addition by measuring the time necessary for calcium release into the buffer using the calcium retention capacity determination method. In this protocol, Ca2+ release occurs due to the depolarization of the membrane and therefore is compatible with the opening of a relatively small PTP [21]. Mitochondria were loaded with 50 µM CaCl2, which was completely taken up into the matrix via the calcium ionophore ETH129. After 2-3 minutes, calcium was released into the buffer through the yPTP in the mitochondria of most strains, except for the single mutant Tim11∆ and double mutant Tim11∆ Om45-GFP, as expected, as well as the double mutant Atp6-CSTC Om45-GFP. The time of calcium release in these mitochondria was two times longer than that in wild-type mitochondria (Fig. 6A). Thus, Atp6-CSTC substitutions restore yPTP properties in Tim11∆ cells, but in Om45-GFP cells, in which Tim11 is present, Atp6-CSTC substitutions have a contrasting effect. The Atp6-CSTC substitutions had a stronger effect on the yPTP induction delay in Om45-GFP mitochondria than Atp6-P153S or Atp6-K80E, as observed in mitochondria from cells grown at a physiological temperature of 28 °C, while the two latter substitutions delayed yPTP induction only in mitochondria isolated from cells grown at elevated temperatures [37].
Next, we measured the swelling of mitochondria induced by high (140 µM) Ca2+ concentrations and caused by the diffusion of sucrose into the matrix. This is only possible when a large yPTP is formed. Under this experimental condition, 60% of the wild-type mitochondria were swollen 15 minutes after calcium addition (Fig. 6B). As expected, in Tim11∆, 25-30% of the mitochondria were swollen in both the wild-type and Om45-GFP backgrounds [22, 35]. Atp6-CSTC substitutions restored the swelling of Tim11∆ mitochondria partially in the Om45 wild-type background, increasing the fraction of swollen mitochondria to approximately 35%, and fully in Om45-GFP mitochondria, increasing the fraction of swollen mitochondria up to 75% in the Atp6-CSTC Tim11∆ Om45-GFP triple mutant. Surprisingly, in the Om45-GFP background, when subunits e and g were present and participated in stabilizing the ATP synthase dimers and yPTP, Atp6-CSTC substitutions increased the fraction of swollen mitochondria by up to 85%, although the time required for calcium release doubled (Fig. 6A). We therefore concluded that Atp6-CSTC substitutions restored the yPTP in mitochondria depleted of dimerization subunits, in terms of both its induction and conductance, in an Om45-GFP-dependent manner. The increased permeability of the mitochondrial membrane in Atp6-CSTC Om45-GFP cells may have led to their decreased growth rate at elevated temperature.
Discussion
This paper extends the research on ATP synthase and its function as the PTP core, a subject that is currently under discussion in the literature [21]. We previously reported that the Atp6-P153S or Atp6-K80E substitution in Om45-GFP cells in S. cerevisiae led to calcium and ROS homeostasis deregulation and delayed the induction of yPTP by calcium, while OXPHOS functioned normally. Moreover, we have shown that the addition of calcium to mitochondria induced the formation of dimeric and oligomeric complexes of ATP synthase during yPTP formation, and P153S and K80E amino acid substitutions in subunit a modulated this process in an Om45-dependent manner [37]. We describe here another example of amino acid substitutions in the a subunit, namely, Atp6-C23S-T205C, which modulate the induction and conductance of yPTP. These two substitutions were selected in vivo as spontaneous suppressors in Atp6-P153S, Atp6-K80E, Atp6-P153S Om45-GFP and Atp6-K80E Om45-GFP cells [37], in which we additionally tried to delete subunit e. Interestingly, S. cerevisiae cells reverted the original Atp6-P153S or Atp6-K80E amino acid substitution to the wild-type amino acid and “moved” the cysteine from position 23 to position 205 of the Atp6 sequence. The absence of subunits e and g had no effect on the growth of S. cerevisiae cells but led to a lack of ATP synthase dimers and drastically reduced yPTP conductance [35]. In S. cerevisiae, ATP synthase dimers are stabilized by the formation of disulfide bonds between the cysteine residues in adjacent a subunits of two monomers, which also occurs in cells lacking the e and g subunits (Fig. 1, Fig. 5; [35]). These genetic results indicate that yPTP function is important for the fitness of S. cerevisiae cells. Complete yPTP dysfunction, a situation possible when Atp6-P153S or Atp6-K80E is combined with the absence of subunit e or g, is lethal. Unfortunately, we could not verify this hypothesis experimentally.
We analyzed the effects of Atp6-CSTC substitutions on ATP synthase activities and yPTP formation. The transfer of the cysteine from position 23 to 205 suppressed the defects caused by a lack of ATP synthase dimerization subunits. Indeed, these substitutions (i) improved the stability of the mitochondrial genome; (ii) restored the mitochondrial network but not its morphology; (iii) stabilized the ATP synthase dimers and monomers and improved enzyme function by restoring oxygen consumption and ATP synthesis to the wild-type level; and (iv) restored the yPTP induction time to the wild-type level and increased yPTP conduction. These results prove that Atp6-P153 and Atp6-K80 are, similar to subunit e, important for ATP synthase dimer stabilization.
Markedly, Atp6-CSTC substitutions have deleterious effects on S. cerevisiae cells in the presence of stable ATP synthase dimers in the Om45-GFP background, similar to Atp6-P153S and Atp6-K80E [37]. Atp6-CSTC in the Om45-GFP background had even stronger effects. These cells were not only sensitive to hydrogen peroxide or high concentrations of calcium but also temperature sensitive when grown on fermentative medium. In these cells, the oxygen consumption and ATP synthesis was reduced to 50% at physiological growth temperature, while in Atp6-P153S Om45-GFP mitochondria only at elevated growth temperature [37]. Interestingly, although yPTP opening is desensitized to calcium in Atp6-CSTC Om45-GFP cells, even more than in Atp6-P153S Om45-GFP and Atp6-K80E Om45-GFP, its conductance for sucrose is higher in Atp6-CSTC Om45-GFP than in the wild-type and Atp6-P153S Om45-GFP cells (Supplementary Fig. S3B). Taking into account the fact that the mitochondrial network at elevated growth temperatures is aggregated in Atp6-CSTC Om45-GFP, as in Atp6-P153S Om45-GFP (Supplementary Fig. S3A), while Atp6-P153S Om45-GFP mitochondria swell normally at permissive temperatures (Supplementary Fig. S3B) and slightly slower than wild-type mitochondria at elevated temperatures [37, 56], we propose that the increased conductance of the yPTP in Atp6-CSTC Om45-GFP cells contributes to the temperature-sensitive growth phenotype, which is absent in Atp6-P153S Om45-GFP cells. The yPTP function is therefore critical for S. cerevisiae cell fitness when overactivated (especially at elevated temperature). The results presented here argue that subunit a has a direct role in yPTP formation together with subunits e and g and the first transmembrane helix of subunit b [35]. Subunit a modulates yPTP induction by calcium ions, which we have shown previously [37], and yPTP conductance, which we show here (Fig. 6B, Fig. 7).
The results presented in this paper are very important in the current debate regarding the role of ATP synthase in PT, particularly because these results have been obtained in vivo. There is evidence that PT is an evolutionarily conserved process between yeasts and mammals [32, 57]. Understanding how the channel operates and its composition and regulation are important objectives for medicine. Indeed, there is much data in the literature showing that abnormal PTP function plays a role in diseases and pathological conditions, including myopathies, Parkinson’s disease, diabetic encephalopathy, cardiomyopathy, and ischemia/reperfusion [58-63]. Furthermore, mutations in the MT-ATP6 gene, encoding ATP synthase subunit a, in addition to their presence in cancer samples [64, 65], are the cause of myopathies, such as neurogenic ataxia retinitis pigmentosa (NARP; [66, 67]), maternally inherited Leigh syndrome (MILS; [68-70]), mitochondrial encephalomyopathy, lactic acidosis, stroke-like episodes (MELAS; [71]), and other diseases, including cardiomyopathies [72]. Whether the role of subunit a in PTP regulation described here plays a role in the pathogenesis of this group of diseases is an intriguing possibility that warrants further study.
Acknowledgements
RK thanks Prof. P. Bernardi for critically reading the manuscript. The authors thank the Laboratory of Electron Microscopy of Mossakowski Medical Research Centre of PAS for the mitochondrial morphology analysis.
Statement of Ethics
The permission number for work with genetically modified microorganisms (GMM I) for RK is 01.2-28/201.
Funding
This work was supported by a grant from the National Science Center of Poland (2016/23/B/NZ3/02098) to RK.
Author Contributions
KN isolated suppressors, constructed the strains, isolated mitochondria, performed the CRC and swelling experiments and protein electrophoresis, and prepared figures; EB isolated mitochondria and performed respiration and ATP synthase activity assays and protein electrophoresis; CP performed phenotypic analysis and respiration measurements and prepared figures; RK designed the work, wrote the original draft of the manuscript, and prepared figures; all authors participated in the review and editing of the manuscript.
Disclosure Statement
The authors declare that they have no competing or financial interests.
References
|
1
Guo H, Bueler SA, Rubinstein JL: Atomic model for the dimeric FO region of
mitochondrial ATP synthase. Science 2017;358:936-940. |
|
|
|
|
|
2 Rubinstein JL, Walker JE, Henderson R:
Structure of the mitochondrial ATP synthase by electron cryomicroscopy. EMBO
J 2003;22:6182-6192. |
|
|
|
|
|
3 Mitchell P, Moyle J: Chemiosmotic hypothesis
of oxidative phosphorylation. Nature 1967;213:137-139. |
|
|
|
|
|
4 Pogoryelov D, Krah A, Langer JD, Yildiz O,
Faraldo-Gomez JD, Meier T: Microscopic rotary mechanism of ion translocation
in the F(o) complex of ATP synthases. Nature Chem Biol 2010;6:891-899. |
|
|
|
|
|
5 Vik SB, Antonio BJ: A mechanism of proton translocation by F1F0 ATP synthases suggested by double mutants of the a subunit. J Biol Chem 1994;269:30364-30369. |
|
|
|
|
|
6 Boyer PD: The binding change mechanism for ATP
synthase--some probabilities and possibilities. Biochim Biophys Acta
1993;1140:215-250. |
|
|
|
|
|
7 Abrahams JP, Leslie AG, Lutter R, Walker JE:
Structure at 2.8 A resolution of F1-ATPase from bovine heart mitochondria.
Nature 1994;370:621-628. |
|
|
|
|
|
8 Hahn A, Parey K, Bublitz M, Mills DJ,
Zickermann V, Vonck J, Kuhlbrandt W, Meier T: Structure of a Complete ATP
Synthase Dimer Reveals the Molecular Basis of Inner Mitochondrial Membrane
Morphology. Mol Cell 2016;63:445-456. |
|
|
|
|
|
9 Davies KM, Anselmi C, Wittig I, Faraldo-Gomez
JD, Kuhlbrandt W: Structure of the yeast F1Fo-ATP synthase dimer and its role
in shaping the mitochondrial cristae. Proc Natl Acad Sci U S A
2012;109:13602-13607. |
|
|
|
|
|
10 Paumard P, Vaillier J, Coulary B, Schaeffer
J, Soubannier V, Mueller DM, Brethes D, di Rago JP, Velours J: The ATP
synthase is involved in generating mitochondrial cristae morphology. EMBO J
2002;21:221-230. |
|
|
|
|
|
11 Habersetzer J, Larrieu I, Priault M, Salin B,
Rossignol R, Brethes D, Paumard P: Human F1F0 ATP synthase, mitochondrial
ultrastructure and OXPHOS impairment: a (super-)complex matter? PloS One
2013;8:e75429. |
|
|
|
|
|
12 Alavian KN, Beutner G, Lazrove E, Sacchetti
S, Park HA, Licznerski P, Li H, Nabili P, Hockensmith K, Graham M, Porter GA,
Jr., Jonas EA: An uncoupling channel within the c-subunit ring of the F1FO
ATP synthase is the mitochondrial permeability transition pore. Proc Natl
Acad Sci U S A 2014;111:10580-10585. |
|
|
|
|
|
13 Bonora M, Bononi A, De Marchi E, Giorgi C,
Lebiedzinska M, Marchi S, Patergnani S, Rimessi A, Suski JM, Wojtala A,
Wieckowski MR, Kroemer G, Galluzzi L, Pinton P: Role of the c subunit of the
FO ATP synthase in mitochondrial permeability transition. Cell Cycle 2013;12:674-683. |
|
|
|
|
|
14 Giorgio V, von Stockum S, Antoniel M, Fabbro
A, Fogolari F, Forte M, Glick GD, Petronilli V, Zoratti M, Szabo I, Lippe G,
Bernardi P: Dimers of mitochondrial ATP synthase form the permeability
transition pore. Proc Natl Acad Sci U S A 2013;110:5887-5892. |
|
|
|
|
|
15 Bernardi P, Rasola A, Forte M, Lippe G: The
Mitochondrial Permeability Transition Pore: Channel Formation by F-ATP
Synthase, Integration in Signal Transduction, and Role in Pathophysiology.
Physiol Rev 2015;95:1111-1155. |
|
|
|
|
|
16 Ichas F, Jouaville LS, Mazat JP: Mitochondria
are excitable organelles capable of generating and conveying electrical and
calcium signals. Cell 1997;89:1145-1153. |
|
|
|
|
|
17 Zorov DB, Juhaszova M, Sollott SJ:
Mitochondrial reactive oxygen species (ROS) and ROS-induced ROS release.
Physiol Rev 2014;94:909-950. |
|
|
|
|
|
18 Maximilian Buja L: Mitochondria in Ischemic
Heart Disease. Adv Exp Med Biol 2017;982:127-140. |
|
|
|
|
|
19 Rasola A, Bernardi P: The mitochondrial
permeability transition pore and its adaptive responses in tumor cells. Cell
Calcium 2014;56:437-445. |
|
|
|
|
|
20 He J, Ford HC, Carroll J, Ding S, Fearnley
IM, Walker JE: Persistence of the mitochondrial permeability transition in
the absence of subunit c of human ATP synthase. Proc Natl Acad Sci U S A
2017;114:3409-3414. |
|
|
|
|
|
21 Bernardi P: Why F-ATP Synthase Remains a
Strong Candidate as the Mitochondrial Permeability Transition Pore. Front Physiol
2018;9:1543. |
|
|
|
|
|
22 Carraro M, Giorgio V, Sileikyte J, Sartori G,
Forte M, Lippe G, Zoratti M, Szabo I, Bernardi P: Channel formation by yeast
F-ATP synthase and the role of dimerization in the mitochondrial permeability
transition. J Biol Chem 2014;289:15980-15985. |
|
|
|
|
|
23 von Stockum S, Giorgio V, Trevisan E, Lippe
G, Glick GD, Forte MA, Da-Re C, Checchetto V, Mazzotta G, Costa R, Szabo I,
Bernardi P: F-ATPase of Drosophila melanogaster forms 53-picosiemen (53-pS)
channels responsible for mitochondrial Ca2+-induced Ca2+ release. J Biol Chem
2015;290:4537-4544. |
|
|
|
|
|
24 Szabo I, Zoratti M: Mitochondrial channels:
ion fluxes and more. Physiol Rev 2014;94:519-608. |
|
|
|
|
|
25 Urbani A, Giorgio V, Carrer A, Franchin C,
Arrigoni G, Jiko C, Abe K, Maeda S, Shinzawa-Itoh K, Bogers JFM, McMillan
DGG, Gerle C, Szabo I, Bernardi P: Purified F-ATP synthase forms a
Ca(2+)-dependent high-conductance channel matching the mitochondrial
permeability transition pore. Nat Commun 2019;10:4341. |
|
|
|
|
|
26 Mnatsakanyan N, Llaguno MC, Yang Y, Yan Y,
Weber J, Sigworth FJ, Jonas EA: A mitochondrial megachannel resides in
monomeric F1FO ATP synthase. Nat Commun 2019;10:5823. |
|
|
|
|
|
27 Bonora M, Morganti C, Morciano G, Pedriali G,
Lebiedzinska-Arciszewska M, Aquila G, Giorgi C, Rizzo P, Campo G, Ferrari R,
Kroemer G, Wieckowski MR, Galluzzi L, Pinton P: Mitochondrial permeability
transition involves dissociation of F1FO ATP synthase dimers and C-ring
conformation. EMBO Rep 2017;18:1077-1089. |
|
|
|
|
|
28 Giorgio V, Burchell V, Schiavone M, Bassot C,
Minervini G, Petronilli V, Argenton F, Forte M, Tosatto S, Lippe G, Bernardi
P: Ca(2+) binding to F-ATP synthase beta subunit triggers the mitochondrial
permeability transition. EMBO Rep 2017;18:1065-1076. |
|
|
|
|
|
29 Johnson KM, Chen X, Boitano A, Swenson L,
Opipari AW, Jr., Glick GD: Identification and validation of the mitochondrial
F1F0-ATPase as the molecular target of the immunomodulatory benzodiazepine
Bz-423. Chem Biol 2005;12:485-496. |
|
|
|
|
|
30 Antoniel M, Jones K, Antonucci S, Spolaore B,
Fogolari F, Petronilli V, Giorgio V, Carraro M, Di Lisa F, Forte M, Szabo I,
Lippe G, Bernardi P: The unique histidine in OSCP subunit of F-ATP synthase
mediates inhibition of the permeability transition pore by acidic pH. EMBO
Rep 2018;19:257-268. |
|
|
|
|
|
31 Neginskaya MA, Solesio ME, Berezhnaya EV,
Amodeo GF, Mnatsakanyan N, Jonas EA, Pavlov EV: ATP Synthase
C-Subunit-Deficient Mitochondria Have a Small Cyclosporine A-Sensitive
Channel, but Lack the Permeability Transition Pore. Cell Rep 2019;26:11-17
e12. |
|
|
|
|
|
32 Kamei Y, Koushi M, Aoyama Y, Asakai R: The
yeast mitochondrial permeability transition is regulated by reactive oxygen
species, endogenous Ca(2+) and Cpr3, mediating cell death. Biochim Biophys
Acta Bioenerg 2018;1859:1313-1326. |
|
|
|
|
|
33 Gutierrez-Aguilar M, Uribe-Carvajal S: The
mitochondrial unselective channel in Saccharomyces cerevisiae. Mitochondrion
2015;22:85-90. |
|
|
|
|
|
34 Velours J, Stines-Chaumeil C, Habersetzer J,
Chaignepain S, Dautant A, Brethes D: Evidence of the proximity of ATP
synthase subunits 6 (a) in the inner mitochondrial membrane and in the
supramolecular forms of Saccharomyces cerevisiae ATP synthase. J Biol Chem
2011;286:35477-35484. |
|
|
|
|
|
35 Carraro M, Checchetto V, Sartori G,
Kucharczyk R, di Rago JP, Minervini G, Franchin C, Arrigoni G, Giorgio V,
Petronilli V, Tosatto SCE, Lippe G, Szabo I, Bernardi P: High-Conductance
Channel Formation in Yeast Mitochondria is Mediated by F-ATP Synthase e and g
Subunits. Cell Physiol Biochem 2018;50:1840-1855. |
|
|
|
|
|
36 Zeng X, Neupert W, Tzagoloff A: The
metalloprotease encoded by ATP23 has a dual function in processing and
assembly of subunit 6 of mitochondrial ATPase. Mol Biol Cell 2007;18:617-626. |
|
|
|
|
|
37 Niedzwiecka K, Tisi R, Penna S, Lichocka M,
Plochocka D, Kucharczyk R: Two mutations in mitochondrial ATP6 gene of ATP
synthase, related to human cancer, affect ROS, calcium homeostasis and
mitochondrial permeability transition in yeast. Biochim Biophys Acta Mol Cell
Res 2018;1865:117-131. |
|
|
|
|
|
38 Lauffer S, Mabert K, Czupalla C, Pursche T,
Hoflack B, Rodel G, Krause-Buchholz U: Saccharomyces cerevisiae porin pore
forms complexes with mitochondrial outer membrane proteins Om14p and Om45p. J
Biol Chem 2012;287:17447-17458. |
|
|
|
|
|
39 Herskowitz I, Jensen RE: Putting the HO gene
to work: practical uses for mating-type switching. Meth Enzym 1991;194:132-146. |
|
|
|
|
|
40 Wysocka-Kapcinska M, Torocsik B, Turiak L,
Tsaprailis G, David CL, Hunt AM, Vekey K, Adam-Vizi V, Kucharczyk R, Chinopoulos
C: The suppressor of AAC2 Lethality SAL1 modulates sensitivity of
heterologously expressed artemia ADP/ATP carrier to bongkrekate in yeast.
PloS One 2013;8:e74187. |
|
|
|
|
|
41 Kanki T, Kang D, Klionsky DJ: Monitoring
mitophagy in yeast: the Om45-GFP processing assay. Autophagy
2009;5:1186-1189. |
|
|
|
|
|
42 Kucharczyk R, Salin B, di Rago JP:
Introducing the human Leigh syndrome mutation T9176G into Saccharomyces
cerevisiae mitochondrial DNA leads to severe defects in the incorporation of
Atp6p into the ATP synthase and in the mitochondrial morphology. Hum Mol
Genet 2009;18:2889-2898. |
|
|
|
|
|
43 Guerin B, Labbe P, Somlo M: Preparation of
yeast mitochondria (Saccharomyces cerevisiae) with good P/O and respiratory
control ratios. Meth Enzym 1979;55:149-159. |
|
|
|
|
|
44 Rigoulet M, Guerin B: Phosphate transport and
ATP synthesis in yeast mitochondria: effect of a new inhibitor: the
tribenzylphosphate. FEBS Lett 1979;102:18-22. |
|
|
|
|
|
45 Somlo M: Induction and repression of mitochondrial
ATPase in yeast. Eur J Biochem 1968;5:276-284. |
|
|
|
|
|
46 Westermann B, Neupert W:
Mitochondria-targeted green fluorescent proteins: convenient tools for the
study of organelle biogenesis in Saccharomyces cerevisiae. Yeast
2000;16:1421-1427. |
|
|
|
|
|
47 Rak M, Tetaud E, Godard F, Sagot I, Salin B,
Duvezin-Caubet S, Slonimski PP, Rytka J, di Rago JP: Yeast cells lacking the
mitochondrial gene encoding the ATP synthase subunit 6 exhibit a selective
loss of complex IV and unusual mitochondrial morphology. J Biol Chem
2007;282:10853-10864. |
|
|
|
|
|
48 Everard-Gigot V, Dunn CD, Dolan BM, Brunner
S, Jensen RE, Stuart RA: Functional analysis of subunit e of the F1Fo-ATP
synthase of the yeast Saccharomyces cerevisiae: importance of the N-terminal
membrane anchor region. Eukaryot Cell 2005;4:346-355. |
|
|
|
|
|
49 Westermann B: Bioenergetic role of
mitochondrial fusion and fission. Biochim Biophys Acta 2012;1817:1833-1838. |
|
|
|
|
|
50 Mukhopadhyay A, Uh M, Mueller DM: Level of
ATP synthase activity required for yeast Saccharomyces cerevisiae to grow on
glycerol media. FEBS Lett 1994;343:160-164. |
|
|
|
|
|
51 Kucharczyk R, Ezkurdia N, Couplan E,
Procaccio V, Ackerman SH, Blondel M, di Rago JP: Consequences of the
pathogenic T9176C mutation of human mitochondrial DNA on yeast mitochondrial
ATP synthase. Biochim Biophys Acta 2010;1797:1105-1112. |
|
|
|
|
|
52 Venard R, Brethes D, Giraud MF, Vaillier J,
Velours J, Haraux F: Investigation of the role and mechanism of IF1 and STF1
proteins, twin inhibitory peptides which interact with the yeast
mitochondrial ATP synthase. Biochemistry 2003;42:7626-7636. |
|
|
|
|
|
53 Lefebvre-Legendre L, Vaillier J, Benabdelhak
H, Velours J, Slonimski PP, di Rago JP: Identification of a nuclear gene
(FMC1) required for the assembly/stability of yeast mitochondrial F(1)-ATPase
in heat stress conditions. J Biol Chem 2001;276:6789-6796. |
|
|
|
|
|
54 Rak M, Zeng X, Briere JJ, Tzagoloff A:
Assembly of F0 in Saccharomyces cerevisiae. Biochim Biophys Acta
2009;1793:108-116. |
|
|
|
|
|
55 Fronzes R, Weimann T, Vaillier J, Velours J,
Brethes D: The peripheral stalk participates in the yeast ATP synthase
dimerization independently of e and g subunits. Biochemistry
2006;45:6715-6723. |
|
|
|
|
|
56 Niedzwiecka K, Kabala AM, Lasserre JP,
Tribouillard-Tanvier D, Golik P, Dautant A, di Rago JP, Kucharczyk R: Yeast
models of mutations in the mitochondrial ATP6 gene found in human cancer
cells. Mitochondrion 2016;29:7-17. |
|
|
|
|
|
57 Azzolin L, von Stockum S, Basso E, Petronilli
V, Forte MA, Bernardi P: The mitochondrial permeability transition from yeast
to mammals. FEBS Lett 2010;584:2504-2509. |
|
|
|
|
|
58 Sileikyte J, Forte M: The Mitochondrial
Permeability Transition in Mitochondrial Disorders. Oxid Med Cell Longev
2019;2019:3403075. |
|
|
|
|
|
59 Ludtmann MHR, Angelova PR, Horrocks MH, Choi
ML, Rodrigues M, Baev AY, Berezhnov AV, Yao Z, Little D, Banushi B,
Al-Menhali AS, Ranasinghe RT, Whiten DR, Yapom R, Dolt KS, Devine MJ, Gissen
P, Kunath T, Jaganjac M, et al.: alpha-synuclein oligomers interact with ATP
synthase and open the permeability transition pore in Parkinson's disease.
Nat Commun 2018;9:2293. |
|
|
|
|
|
60 Yan S, Du F, Wu L, Zhang Z, Zhong C, Yu Q,
Wang Y, Lue LF, Walker DG, Douglas JT, Yan SS: F1F0 ATP Synthase-Cyclophilin
D Interaction Contributes to Diabetes-Induced Synaptic Dysfunction and
Cognitive Decline. Diabetes 2016;65:3482-3494. |
|
|
|
|
|
61 Fernandez-Sanz C, Ruiz-Meana M, Castellano J,
Miro-Casas E, Nunez E, Inserte J, Vazquez J, Garcia-Dorado D: Altered FoF1
ATP synthase and susceptibility to mitochondrial permeability transition pore
during ischaemia and reperfusion in aging cardiomyocytes. Throm Hem
2015;113:441-451. |
|
|
|
|
|
62 Halestrap AP, Richardson AP: The
mitochondrial permeability transition: a current perspective on its identity
and role in ischaemia/reperfusion injury. J Mol Cell Cardiol 2015;78:129-141. |
|
|
|
|
|
63 Angelin A, Bonaldo P, Bernardi P: Altered
threshold of the mitochondrial permeability transition pore in Ullrich
congenital muscular dystrophy. Biochim Biophys Acta 2008;1777:893-896. |
|
|
|
|
|
64 Jarviaho T, Hurme-Niiranen A, Soini HK,
Niinimaki R, Mottonen M, Savolainen ER, Hinttala R, Harila-Saari A, Uusimaa
J: Novel non-neutral mitochondrial DNA mutations found in childhood acute
lymphoblastic leukemia. Clin Genet 2018;93:275-285. |
|
|
|
|
|
65 Li L, Chen L, Li J, Zhang W, Liao Y, Chen J,
Sun Z: Correlational study on mitochondrial DNA mutations as potential risk
factors in breast cancer. Oncotarget 2016;7:31270-31283. |
|
|
|
|
|
66 Duno M, Wibrand F, Baggesen K, Rosenberg T,
Kjaer N, Frederiksen AL: A novel mitochondrial mutation m.8989G>C
associated with neuropathy, ataxia, retinitis pigmentosa - the NARP syndrome.
Gene 2013;515:372-375. |
|
|
|
|
|
67 Baracca A, Barogi S, Carelli V, Lenaz G,
Solaini G: Catalytic activities of mitochondrial ATP synthase in patients
with mitochondrial DNA T8993G mutation in the ATPase 6 gene encoding subunit
a. J Biol Chem 2000;275:4177-4182. |
|
|
|
|
|
68 Schon EA, Santra S, Pallotti F, Girvin ME:
Pathogenesis of primary defects in mitochondrial ATP synthesis. Semin Cell
Dev Biol 2001;12:441-448. |
|
|
|
|
|
69 Uittenbogaard M, Brantner CA, Fang Z, Wong
LC, Gropman A, Chiaramello A: Novel insights into the functional metabolic
impact of an apparent de novo m.8993T>G variant in the MT-ATP6 gene
associated with maternally inherited form of Leigh Syndrome. Mol Genet Metab
2018;124:71-81. |
|
|
|
|
|
70 Akagi M, Inui K, Tsukamoto H, Sakai N,
Muramatsu T, Yamada M, Matsuzaki K, Goto Y, Nonaka I, Okada S: A point
mutation of mitochondrial ATPase 6 gene in Leigh syndrome. Neuromuscul Disord
2002;12:53-55. |
|
|
|
|
|
71 Aure K, Dubourg O, Jardel C, Clarysse L,
Sternberg D, Fournier E, Laforet P, Streichenberger N, Petiot P,
Gervais-Bernard H, Vial C, Bedat-Millet AL, Drouin-Garraud V, Bouillaud F,
Vandier C, Fontaine B, Lombes A: Episodic weakness due to mitochondrial DNA
MT-ATP6/8 mutations. Neurology 2013;81:1810-1818. |
|
|
|
|
|
72 Dautant A, Meier T, Hahn A,
Tribouillard-Tanvier D, di Rago JP, Kucharczyk R: ATP Synthase Diseases of
Mitochondrial Genetic Origin. Front Physiol 2018;9:329. |
|