While a nuclear gene encoded ATE1 in eukaryotes, our search of ATE1 homologs in archaea, where most of the nuclear genome originated (Williams et al., 2013), returned no matches (Figure 1). However, sequence homologs of ATE1 are present in a large population of bacteria, including modern alpha-proteobacteria (Buffet et al., 2020), a close relative to the ancient alpha-proteobacteria that became mitochondria (Figures 1A–C). As such, it is likely that the ATE1 gene, like many mitochondria-associated genes, was transferred to the nuclear genome during mitochondrial domestication (Janeway and Medzhitov, 2002; Buffet et al., 2020). To gain further insight into the relationship between ATE1 and mitochondria from the perspective of molecular evolution, we examined the status of mitochondrial development and the presence of the ATE1 gene in several branches of eukaryotes. While almost all eukaryotes contain the ATE1 gene, two exceptions exist. One is the family of giardia, and the other is the superfamily of dinoflagellates and apicomplexan. Intriguingly, both families lack respiratory-active mitochondria. Instead, they possess mitosomes, a reduced form of mitochondria with minimal functions that cannot perform oxidative phosphorylation (Figure 1D). Since these two families are distally related and separated by many other families that possess ATE1, their loss of ATE1 is unlikely to derive from the same ancestor. For the same reason, their lack of respiratory-competent mitochondria is likely the result of convergent evolution. These data suggest that the presence of ATE1 may be essential for maintaining fully functional mitochondria.

Several studies have used fluorescent protein fusions to show the localization of ATE1 in the nucleus or the cytoplasm (Rai and Kashina, 2005; Hu et al., 2006; Rai et al., 2006; Wang et al., 2011). However, the potential localization of ATE1 to mitochondria was not examined directly. To investigate this question, we first utilized a budding yeast (Saccharomyces cerevisiae) strain with a genomically-integrated green fluorescent protein (GFP) fused to the C-terminus of the endogenous Ate1 (Ate1-GFP), which showed that a subfraction of the GFP-fused Ate1 colocalizes with Mitotracker red-stained mitochondria (Figure 2A). Next, to test if the mitochondrial localization of ATE1 was organism-specific, we fused a C-terminal GFP to the mouse ATE1, transcript variant 1, a ubiquitously-expressed isoform (Rai and Kashina, 2005; Wang et al., 2011). To avoid the competition of the endogenous protein, this recombinant ATE1 was expressed in ATE1-knockout (KO) mouse embryonic fibroblasts (MEF). We detected a fraction of the fluorescent ATE1 colocalizing with the Mitotracker red-stained branched mitochondrial network surrounding the nucleus (Figure 2B). In mice, there are six ATE1 transcript variants, four of which (transcript variants 1, 2, 3, and 4) are known to be translated into protein isoforms that are >90% identical in sequences (Rai and Kashina, 2005; Hu et al., 2006). We found that the mitochondrial localization of ATE1 is not isoform-specific, because the other three known protein isoforms also appear to localize to mitochondria (Figure 2B; see also Supplementary Figure 1A for the expression level of the tested isoforms).

To further assess the mitochondrial localization of endogenous ATE1, we purified mitochondria from MEF by differential centrifugation and probed for endogenous ATE1 by immunoblotting. We were able to detect a fraction of ATE1 associated with purified mitochondria from wildtype (WT) MEF, while the specificity of the anti-ATE1 antibody was validated by the absence of signals in mitochondria from ATE1-KO cells (Figure 2C; see also the purity of the mitochondrial fraction in Figure 2D). Furthermore, a proteinase K protection assay in purified mitochondria showed that the mitochondrion-associated ATE1 is resistant to proteinase K digestion, except when the mitochondrial membranes are disrupted by sonication (Figure 2E). These data indicated that a fraction of ATE1 localizes within mitochondria. To estimate the percentage of ATE1 that is associated with mitochondria, we used a dilution assay of the whole cell and isolated mitochondria to compare the ratios of ATE1 over Voltage-dependent anion channel (VDAC), an established mitochondrial marker. We found that only about 0.5% of total endogenous ATE1 in the cell is associated with mitochondria (Supplemental Figure 1B).

We next examined in whole cells the steady-state levels of VDAC and TOM20, two mitochondrial markers, and did not find a significant difference between WT and ATE1-KO MEF (Figure 3A). This suggested that the ATE1 does not directly affect cellular mitochondrial mass. However, when we examined mitochondrial morphology by electron microscopy, we found significant differences between WT and ATE1-KO cells (Figure 3B). Remarkably, mitochondrial filaments in ATE1-KO cells were significantly shorter and wider (Figure 3C), reminiscent of the swollen mitochondria often observed in respiratory-deficient cells, as well as typical cancer cells with a glycolytic metabolic profile (Alirol and Martinou, 2006).

To assess whether any mitochondrial functional difference accompanies the observed morphological changes, we measured cellular respiration in ATE1-KO and WT MEF. Our data showed that ATE1-KO cells had a ~30% reduction in endogenous oxygen consumption rate compared to WT MEF (Figure 3D). As an essential control, we reconstituted ATE1-KO MEF with a recombinant ATE1 (transcript variant 3, which was shown to have the most potent anti-tumor growth effect) at a level comparable to the endogenous protein. Recombinant ATE1 rescued the ATE1-KO respiratory defects (Figure 3E). Consistent with the observations in mammalian cells, we found that the deletion of ate1 in budding yeast also lead to slower growth in respiratory media (Figure 3F), reduced endogenous cell respiration by nearly 30%, and attenuated the mitochondrial membrane potential by more than 50% (Figures 3G,H).

The mitochondrial respiratory chain (MRC) is formed by four multimeric complexes (complex I to IV, CI-CIV) and two mobile electron carriers (coenzyme Q and cytochrome c). Electrons from the reducing equivalents NADH and FADH₂ enter the MRC at the level of CI and CII, respectively, and are sequentially transferred until CIV, where the final electron acceptor, molecular oxygen, is reduced to water. Additionally, MRC CI, CIII, and CIV dynamically associate in supramolecular structures known as supercomplexes. To further dissect the respiratory defect observed in the ATE1-KO cells, we first measured oxygen consumption rates in digitonin-permeabilized cells following the addition of substrates specific for CI (glutamate-malate) or CII (succinate). When we examined the overall impact of ATE1 on mitochondrial respiration, we found that the combined rate of CI- and CII substrate-driven respiration, and the maximum respiratory capacity measured in the presence of the uncloupler FCCP, are both reduced in KO cells (Figure 4A). CI does not contribute to the deleterious effect on cellular respiration since the glutamate-malate oxidation rate is slightly but significantly higher in ATE1-KO cells than in control. In contrast, the oxygen consumption rate in the presence of succinate was lowered by nearly 50% in ATE1-KO cells (Figures 4A,B and Supplementary Figure 2), indicating a specific CII activity defect.

To determine the cause of the respiratory alteration, we examined the levels of individual mitochondrial respiratory complexes and supercomplexes by Blue Native-PAGE followed by immunoblotting. Consistent with the respiratory data, the steady-state levels of mitochondrial CII in ATE1-KO cells were found to be ~50% than in WT MEFs (Figure 4C). However, we also observed a decrease in dimeric complex III (CIII₂) accumulation, while CIV was unaffected (Figure 4C). Moreover, an evident remodeling of the MRC organization was detected. In ATE1-KO cells, the levels of CI-containing supercomplexes (I+III₂ and I+III₂+IV_1−n) were increased, whereas the amount of CIII that accumulated in CIII₂ and the III₂+IV supercomplex was attenuated (Figures 4D,E). This remodeling facilitates an increase in CI stability, presumably in an attempt to compensate for the lower levels of CII.

MRC CII or succinate dehydrogenase (SDH) is also an enzyme of the TCA cycle, in which it catalyzes the conversion of succinate to fumarate. Succinate, when in excess, promotes glycolysis and therefore acts as a critical signaling component connecting the TCA cycle to glycolysis. When we measured the enzymatic activity of SDH, we found it significantly reduced in ATE1-KO cells (Figure 5A). Correspondingly, the level of succinate in ATE1-KO MEF was nearly 2-fold higher than in WT (Figure 5B). To test if this would change the balance between mitochondrial OXPHOS and glycolysis, we measured the viability of ATE1-KO MEF under glucose starvation and found that they are significantly more sensitive to the lack of glucose supply than WT cells (Figures 5C,D).

SDH is comprised of four subunits known as SDHA, B, C, and D, all of which are encoded in the nuclear genome and then imported into mitochondria (Van Vranken et al., 2015). When we examined the levels of these subunits in the mitochondrial fraction, we found that the levels of SDHB and SDHC, as well as the palmitoylation level of SDHD, are decreased in mitochondria in ATE1-KO MEF, in agreement with the lower levels of CII (Figure 6A). However, the total protein levels of these subunits in whole ATE1-KO cells were either unchanged or even slightly increased compared to WT cells (Figure 6B) while the transcription of SDH subunits is drastically increased in ATE1-KO MEF cells (Figure 6C), likely as a compensation for the functional SDH deficiency. These data indicate that ATE1 may affect the mitochondrial import and/or assembly of SDH subunits, ultimately leading to CII deficiency.

Immortalized mouse embryonic fibroblasts (MEFs) that are either wild-type or genomic knockout for ATE1 (ATE1-KO) were a gift from Dr. Anna Kashina (University of Pennsylvania), prepared as described elsewhere (Zhang et al., 2012). The human embryonic kidney cell line (HEK 293T; clone T7) was obtained from ATCC.

For routine growth and maintenance, unless otherwise indicated, mammalian cells were cultured in media containing high-glucose DMEM supplemented with 1 mM pyruvate and glutamine (Gibco, Cat# 10569) with 10% FBS (HyClone, Cat# SH30910.03). No antibiotics were used for the cell culture to minimize interference with cellular metabolism, and the cultured cells were periodically checked for bacterial or mycoplasma contamination. The cells were cultured in a 5% CO₂ incubator at 37°C, unless otherwise indicated. To minimize any effects of contact inhibition, only actively growing cells in culture density of <50% confluency were used for any test in this study, unless otherwise indicated.

The Saccharomyces cerevisiae strains used in this study include BY4741 (MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0) and W303-1A (MATa leu2-3,112 trp1-1 can1-100 ura3-1 ade2-1 his3-11,15 ybp1-1). They are both obtained from Open Biosystems. A strain carrying a null ate1Δ:KanMX cassette in the BY4741 background was obtained from Open Biosystems. To create ate1-deletion in the W303-1A yeast, we applied the knockout cassette amplified from the ate1Δ:KanMX in BY4741-strain yeast with two primers as described before (Kumar et al., 2016):

ATE1300UP: ATGGTGCTGTGCTTGTAATTGCC

ATE1300DOWN: GCTCATCAAAAACTAAGAATAAGAG

The strain with a GFP fused to the C-terminal end of endogenous Ate1 in the native chromosome locus in BY4741 genetic background was obtained from Open Biosystems. The identity of each knockout strain listed above was confirmed with PCR genotyping.

Yeast culture media were prepared as described below:

YPD: 2% glucose, 1% yeast extract, 2% peptone,

SD (Synthetic Defined) Medium (per 1,000 ml): Yeast Nitrogen Base, 1.7 g, Ammonium sulfate, 5 g, Dextrose/galactose/raffinose, 20 g, required amino acids, 50 mg, uracil (if required), 50 mg.

YPEG: 1% yeast extract, 2% peptone, 3% (v/v) glycerol, and 2% ethanol.

For solid media plates, 2% agar was added to the liquid media.

Strains grown in liquid and solid media were incubated at 30°C unless otherwise indicated.

For most serial dilution growth assays, a single colony of yeast was inoculated in liquid medium and allowed to grow to the log phase before spot platting of serial dilutions as described elsewhere (Kumar et al., 2012).

Primary antibodies used in this study are (unless otherwise indicated): rat anti ATE1 (EMD-Millipore, Billerica, MA, Cat# MABS436, clone 6F11), mouse anti-GFP (Roche Diagnostics, Indianapolis, IN, Cat# 11814460001), mouse anti porin/VDAC1 (Abcam, Cambridge, MA, Cat# ab14734), anti TOM20 (Santa Cruz Biotechnology, Dallas, TX, Cat# SC11415), anti-beta-actin (Sigma-Aldrich, St. Louis, MO, Cat# A1978), mouse anti-beta-tubulin (Sigma-Aldrich, St. Louis, MO, Cat# T5201), anti-histone H3 (Abcam, Cambridge, MA, Cat# ab1971), rabbit anti-SDHA (One World Lab, San Diego, Ca, Cat# 53800), mouse anti-SDHB (Abcam, Cambridge, MA, Cat# ab14714), rabbit anti-SDHC (Abcam, Cambridge, MA, Cat# ab155999), rabbit anti-SDHD (EMD-Millipore, Billerica, MA, Cat# ABC835), rabbit anti-Hexokinase2 (One World Lab, San Diego, Ca, Cat#55911), mouse anti-PHD2 (Santa Cruz Biotechnology, Dallas, TX, Cat# SC271835).

Secondary antibodies used in this study are (unless otherwise indicated): Anti-mouse-HRP (Pierce, now thermos Scientific, Cat # 31430), Anti-rabbit-HRP (Thermo Fisher Cat # 65-6120), Anti-Rat-HRP (BioLegand, Cat # 405405), Anti-mouse-Alexa 488 (Molecular Probe /Invitrogen, Cat # A21202), Anti-rabbit-Alexa 488 (Molecular Probes/ Invitrogen, Cat # 21206), Anti-Rat-Alexa 594 (Molecular Probe/Invitrogen, Cat # A21209).

Cell lysate or protein samples were generally prepared in SDS-loading buffer and boiled for 10 min for denaturing. For analysis of membrane-associated proteins, samples were first dissolved in 8M urea/PBS and then added SDS-loading buffer and denatured at 55°C for 10 min. The proteins were separated by electrophoresis in 4–20 or 10% SDS-PAGE (unless otherwise indicated) as needed. The proteins were then transferred to nitrocellulose or PVDF membranes for Western Blot analysis. The protein bands were examined with Chemifluorescence visualization utilizing the HRP conjugated on secondary antibodies and reagents provided in the BM Chemifluorescence Western Blotting Kit Mouse/Rabbit (Roche) or the SuperSignal West Femto Chemiluminescence Kit (Pierce). The chemiluminescent signals were either documented on film (Denville) or by GE Amersham Imager model 600. The films were scanned by an Epson Perfection 2400 photo/film scanner with at least 1,200 dpi resolution to convert into digital forms and then analyzed with Image J (NIH). For images documented by the GE imager, an ImageQuant TL software pack (v8.1) and its 1D gel analysis module were used to examine the intensity of signals by densitometry.

Cells were permeabilized in PBS with 2 mg/ml digitonin (Sigma-Aldrich, Cat# D141) at 4°C for 10 min. Cells were centrifuged at 10,000 × g for 5 min at 4°C and washed twice with PBS. Cell pellets were resuspended in 1.5 M aminocaproic acid, 50 mM Bis–Tris pH 7.0, and total protein concentration was determined by the Folin method (Lowry et al., 1951). Proteins were extracted either with 1% lauryl maltoside (for monomeric complex analysis) or 1% digitonin (for supercomplex analysis). After centrifugation at 22,000 × g for 30 min at 4°C, a buffer containing 750 mM aminocaproic acid, 50 mM Bis-Tris pH 7.0, 0.5 mM, EDTA, and 5% Serva blue G was added to the clarified extracts. Samples were loaded on a linear 3–12% polyacrylamide blue native gel (Invitrogen), transferred to PVDF membrane, and analyzed by immunoblotting with the following antibodies: anti-COX1 (Abcam, Cat#Ab14705), anti-COX3 (Abcam, Cat# Ab110259), anti-NDUFA9 (Abcam, Cat# Ab14713), anti-CORE2 (Abcam, Cat# Ab 14745), anti-SDHA (Abcam, Cat# Ab14715), anti-porin/VDAC (Abcam, Cat# Ab14734), anti-TOM20 (Santa Cruz Biotechnology, Cat# sc-11415), second antibodies anti-mouse HRP (Rockland, Cat# 610-4302), anti-rabbit HRP (Rockland, Cat# 611-1302).

ATE1 KO MEF stably transfected with ATE1-1-GFP, ATE1-2-GFP, ATE1-3-GFP, and ATE1-4-GFP was generated as described previously (Zhang et al., 2015) except cells were enriched with FACS without puromycin selection. Briefly, retroviruses carrying different C-terminal GFP tagged isoform of Ate-1 were constructed by transfecting HEK293T cells with the low-expression pBabe-Puro vectors carrying coding sequence for the desired isoform and the vectors of GAG-Pol and VSV-G vector. The viruses were then allowed to infect ATE1 KO MEF in the presence of 10 mg/mL polybrene. Successfully transfected MEF were enriched by fluorescence sorting.

The numbers of resuspended cells in solution were counted with an automated Biorad TC20 cell counter, and dead cells were excluded with trypan blue unless otherwise indicated. The diameters of the resuspended cells were also measured by the built-in feature of the cell counter. All peaks of size distributions and the associated cell numbers displayed by the counters were used to calculate the average diameter of the cell.

For the tests described below, we used actively growing cell cultures at a confluency lower than 50%. The measurements of whole-cell and mitochondrial respiration rates were done according to a protocol published elsewhere (Barrientos et al., 2009). In brief, cells were collected by trypsinization, neutralized in FBS-containing medium, and washed with DPBS. To measure whole-cell respiration, the cells were resuspended in RPMI 1640 medium with L-Glutamine and 25 mM HEPES (Sigma-Aldrich, Cat# SLM-140) at 37°C. The numbers of live cells were counted using Trypan blue on a TC-20 automated cell counter (Bio-Rad). The concentration of all cell lines used in the same test were adjusted to 4 × 10⁶ cells/ml and then transferred to the chamber of a polarographic apparatus with a Clark-type O2 electrode (Hansatech, Oxygraph plus system) for measurement of cell respiration at 37°C. At the end of the measurement, KCN (Sigma-Aldrich, Cat# 60178) was added to the cell suspension to a final concentration of 2 mM to inhibit cell respiration. KCN-sensitive cellular respiratory rates were obtained by subtracting oxygen consumption rates before and after KCN addition.

To measure substrate-driven mitochondrial respiration, the cells were resuspended in warm (37°C) permeablized-cell respiration buffer (PRB) containing 0.3 M mannitol, 10 mM KCl, 5 mM MgCl₂, 0.5 mM EDTA, 0.5 mM EGTA, 1 mg/ml BSA and 10 mM KH₃PO4 (pH 7.4). The numbers of live cells were counted using Trypan blue on a TC-20 automated cell counter (Bio-Rad). The concentrations of each cell type in the same test were adjusted to 4 × 10⁶ cells/ml. An aliquot of the cell suspension was supplemented with freshly prepared hexokinase (Sigma-Aldrich, Cat# H-5500) to 10 U/mL, and ADP to 2 mM and 1 mL of the cell suspension was transferred to the polarographic chamber/oxygraph containing a Clark-type O₂ electrode (Hansatech, Oxygraph plus system) with a temperature setting of 37°C. The whole-cell respiration was measured before digitonin was added.

To measure substrate-driven respiration, the cell membrane was permeabilized by adding freshly prepared digitonin in optimized ratios of 20 μg per 10⁶ cells for WT MEF, or 10 μg per 10⁶ cells for ATE1-KO MEF. The substrate-driven oxygen consumption rate was then examined by adding specific MRC substrates using a Hamilton microsyringe. To measure complex I-driven respiration, we added 5 mM each of glutamate (Sigma-Aldrich, Cat# G8415) and malate (Sigma-Aldrich, Cat# M1124). To measure complex II, we used 10 mM succinate (ICN Biomedicals, Cat# 102972). To measure uncoupled respiration, we added 3 μM carbonilcyanide p-triflouromethoxyphenylhydrazone (FCCP) (Sigma-Aldrich, Cat# C2920), and to assess the specificity of the measurements, we inhibited respiration with 2 mM of the CIV inhibitor potassium cyanide (KCN) (Sigma-Aldrich, Cat# 60178).

Mitochondria were purified from exponentially growing cultured cells based on the Gaines method, updated by Enriquez's group (Fernandez-Vizarra et al., 2010). In brief, cells were harvested by trypsinization and washed twice with cold DPBS before being resuspended in swelling buffer (10 mM Tris–HCl, pH 7.4, 10 mM KCl, 0.5 mM MgCl₂) and incubation on ice for 5 min. The cells were then homogenized with Teflon-glass douncer until at least 75% of the cells were broken. The homogenized lysate was then mixed with sucrose solution to reach a final concentration of 0.25 M of sucrose and then centrifuged at 600 × g for 5 min at 4°C twice to remove cell debris. The resulting supernatant was centrifuged at 7,000 × g for 10 min at 4°C to pellet the mitochondrial fraction. The pellet was washed with the STE buffer (0.32 M sucrose, 1 mM EDTA, and 10 mM Tris–HCl, pH 7.4) and centrifuged again at 7,000 × g for 10 min at 4°C. The pellet was gently resuspended in freshly prepared, ice-cold mitochondria resuspending buffer (MRB) (250 mM mannitol, 5-mM HEPES (pH 7.4), and 0.5-mM EGTA. The mixture was layered on top of Percoll medium (225-mM mannitol, 25-mM HEPES (pH 7.4), 1-mM EGTA, and 30% Percoll (vol/vol). Additional MRB buffer was added to fill the volume of the centrifuge tube before being centrifuged in a swing-bucket rotor at 95,000 × g for 30 min at 4°C. A Beckman Coulter Optima L-100 XP ultracentrifuge with an SW40 rotor (Beckman, Fullerton, CA, USA) was used in this study. The purified mitochondria, which was present in a semi-transparent layer directly above the pellet on the bottom, was then recovered.

The cell respiration of yeasts was assayed polarographically using a Clark-type oxygen electrode (Hansatech Instruments, Norfolk, UK) at 30°C as described elsewhere (Barrientos et al., 2002). The specific activities reported were corrected for KCN-insensitive respiration.

The measurement of mitochondrial membrane potential (Ψ_m) in yeast was performed as described elsewhere (Ocampo et al., 2012). In brief, yeast cells were incubated with 5 μM TMRM of Tetramethyl Rhodamine Methyl Ester (TMRM), a cell-permeant, cationic, fluorescent dye that is readily sequestered by viable mitochondria, at 30°C for 30 min, as reported (Nicholls and Ward, 2000). The cells were then washed twice in PBS and were analyzed with flow cytometry analysis on a Becton Dickinson (BD) FACSAria^TM II Flow Cytometer. Excitation was performed at 532 nm; emission was detected using a 25 nm bandpass filter centered at 575 nm (Becton Dickinson, NJ, USA). Dissipation of Ψ_m causes TMRM to leak out of mitochondria into the cytosol, where TMRM became unquenched, producing an increase in fluorescence (Nicholls and Ward, 2000). To confirm the mitochondrial specificity of the signal of TMRM, ionophore carbonyl cyanide m-chlorophenyl hydrazone (CCCP), which dissipates the membrane potential, was applied to treat the cells as controls.

Actively growing cells below 50% confluency were used. Cells were harvested by trypsin and then immediately washed three times with ice-cold DPBS. The numbers, viability, and diameters of the resuspended cells were measured on the Biorad TC20 automated cell counter. The same number of cells (40 million cells in this test) were used for each cell type. The cells were collected by centrifugation at 2,500 × g at 4°C for 3 min. The pellets were weighted and 20 volumes of ice-cold H₂O was added. The cells were then lysed by brief sonication, and the cell debris was removed by centrifugation at 20,000 × g at 4°C for 10 min. The supernatant was further centrifuged at 100,000 × g at 4°C for 1 h to remove large cellular complexes. The subsequent supernatant was then filtered through a 10K MWCO membrane (Thermo Scientific, Rockford, IL, Cat#PI88513) by centrifugation at 4°C to remove large proteins. The filtrate protein concentration was quantified by the Bradford assay using reagents from Bio-Rad (Cat# 500 0205). The cell lysate filtrate was then extracted by acetonitrile. The resulting supernatant was dried under N₂ and then derivatized by HCl n-butanol. Upon being dried under N₂, the derivative precipitation was reconstituted with 80% methanol and loaded in LC-MS/MS. The level of succinic acid in the filtrate was quantified by Agilent triple quant LC-MS/MS. The final cellular concentration of succinic acid was normalized by cell number, cell volumes, dilution factor, and the protein concentration in the filtered lysate.

The experiment was performed according to the previously published method (Clemente et al., 2013). In brief, purified mitochondria were resuspended in a buffer containing 10 mM Tris-HCl, pH 7, 10 mM KCl, 0.15 mM MgSO₄, and 0.25 M sucrose. As control, one sample was subjected to brief sonication to break the membranes. All samples were then incubated in ice for 45 min in the presence of proteinase K at a final concentration of 0, 0.32, or 0.64 μg/ml. Mitochondria were recovered by centrifugation at 8,000 × g for 15 min at 4°C and analyzed by Western blotting.

MEF cells (WT or ATE1-KO) were cultured in high-glucose DMEM containing 25 mM glucose and 1 mM pyruvate (Gibco Cat#10569) supplemented with 5% FBS (Hyclone, Cat# SH30910.03) for several generations. Immediately before the experiment, the cells were split and cultured for one generation (24 h) in the same high-glucose DMEM except with 5% dialyzed FBS (Life Technologies Cat# 26400-044). At the time of the experiment, the cells were trypsinized, washed with DPBS, and then resuspended in either starving media with glucose-free, pyruvate-free DMEM (Gibco, Cat# 11966) and 5% dialyzed FBS, or non-starving media with high-glucose, 1 mM pyruvate DMEM (Gibco Cat#10569) and 5% dialyzed FBS. The cells were then inoculated into 6-cm culture dishes with 50,000 cells per dish so that the cells would stay at a non-confluent culture density through the duration of tests. Live cells that remained attached to the plate were then counted at given time points, using trypan blue to exclude dead cells.

Optical and fluorescent images of cells were captured on a Zeiss Observer equipped with a series of objectives and Zen Pro software. Analysis of the images were performed with the Zen Pro software.

The images were processed in Adobe Photoshop by adjusting the display levels while preserving the linearity of the signals. The figures were assembled in Adobe Illustrator.

The phylogenetic tree of the ATE1 protein was calculated based on amino acid sequences with Clustal Omega program on the Uniprot website. The detailed alignment of amino acid sequences in ATE-C domain (Pfam ID: PF04377) between two species was performed with NCBI BLASTp. The sunburst graph showing the distribution of ATE-C domain (Pfam ID: PF04377) was generated with corresponding tools from pfam.xfam.org hosted by EMBL-EBI.

RNA was extracted using Quick-RNA MiniPrep Kit (Genesee Scientific, Cat #: 11-328). The corresponding cDNA was prepared by using Superscript First-strand RT-PCR kit (Invitrogen, Cat#: 11904-018). Quantitative real-time PCR was performed by using SsoAdvanced™ Universal SYBR® Green Supermix (Biorad, Cat#: 1725271) on a CFX Connect Real-Time PCR machine (Biorad). The samples were run in multiple replicates on Hard- Shell PCR 96 well plates from Biorad (Cat#: HSP9601). PCR conditions: initial denaturation for 30 s at 95°C, followed by 40 cycles with 95°C for 15 s and 58°C for 1 min. After each run, a melting curve was measured to confirm the specificity of the amplification. The relative expression of the SDH subunits were calculated by delta Ct method. The mRNA of β-actin (ACTB) was used as housekeeping gene/reference gene for loading controls.

The primers targeting different subunits of Succinate dehydrogenase and β-actin are listed below.

The statistical significance for comparison of quantitative assessments performed in WT and ATE1-KO MEF or yeast cells was estimated by the Student t-test. The data distribution was considered one-side or two-side based on the presence of the error bars in the displayed data for comparison. Error bars represent standard error of the mean (SEM) or standard deviation (SD), as indicated in the figure legends. A minimum p-value of 0.05 was considered significant.