Mitochondrial protein import clogging as a mechanism of disease

  1. Liam P Coyne
  2. Xiaowen Wang
  3. Jiyao Song
  4. Ebbing de Jong
  5. Karin Schneider
  6. Paul T Massa
  7. Frank A Middleton
  8. Thomas Becker
  9. Xin Jie Chen  Is a corresponding author
  1. Department of Biochemistry and Molecular Biology, State University of New York Upstate Medical University, United States
  2. Institute of Biochemistry and Molecular Biology, Faculty of Medicine, University of Freiburg, Germany
  3. Institute of Biochemistry and Molecular Biology, Faculty of Medicine, University of Bonn, Germany
  4. Proteomics and Mass Spectrometry Core Facility, State University of New York Upstate Medical University, United States
  5. Department of Microbiology and Immunology, State University of New York Upstate Medical University, United States
  6. Department of Neurology, State University of New York Upstate Medical University, United States
  7. Department of Neuroscience and Physiology, State University of New York Upstate Medical University, United States

Abstract

Mitochondrial biogenesis requires the import of >1,000 mitochondrial preproteins from the cytosol. Most studies on mitochondrial protein import are focused on the core import machinery. Whether and how the biophysical properties of substrate preproteins affect overall import efficiency is underexplored. Here, we show that protein traffic into mitochondria can be disrupted by amino acid substitutions in a single substrate preprotein. Pathogenic missense mutations in ADP/ATP translocase 1 (ANT1), and its yeast homolog ADP/ATP carrier 2 (Aac2), cause the protein to accumulate along the protein import pathway, thereby obstructing general protein translocation into mitochondria. This impairs mitochondrial respiration, cytosolic proteostasis, and cell viability independent of ANT1’s nucleotide transport activity. The mutations act synergistically, as double mutant Aac2/ANT1 causes severe clogging primarily at the translocase of the outer membrane (TOM) complex. This confers extreme toxicity in yeast. In mice, expression of a super-clogger ANT1 variant led to neurodegeneration and an age-dependent dominant myopathy that phenocopy ANT1-induced human disease, suggesting clogging as a mechanism of disease. More broadly, this work implies the existence of uncharacterized amino acid requirements for mitochondrial carrier proteins to avoid clogging and subsequent disease.

Editor's evaluation

This manuscript describes important insight into the molecular mechanism by which destabilized mitochondrial proteins 'clog' import channels and contribute to the pathologic mitochondrial and cellular dysfunction implicated in human disease. The evidence supporting this conclusion is convincing, utilizing yeast, mammalian cell culture, and mouse models. This work, which defines an interesting mechanism of disease pathogenesis, will be of broad interest to researchers in the fields of mitochondrial biology, protein quality control and proteostasis.

https://doi.org/10.7554/eLife.84330.sa0

eLife digest

Inside our cells, compartments known as mitochondria generate the chemical energy required for life processes to unfold. Most of the proteins found within mitochondria are manufactured in another part of the cell (known as the cytosol) and then imported with the help of specialist machinery. For example, the TOM and TIM22 channels provide a route for the proteins to cross the two membrane barriers that separate the cytosol from the inside of a mitochondrion.

ANT1 is a protein that is found inside mitochondria in humans, where it acts as a transport system for the cell’s energy currency. Specific mutations in the gene encoding ANT1 have been linked to degenerative conditions that affect the muscles and the brain. However, it remains unclear how these mutations cause disease.

To address this question, Coyne et al. recreated some of the mutations in the gene encoding the yeast equivalent of ANT1 (known as Aac2). Experiments in yeast cells carrying these mutations showed that the Aac2 protein accumulated in the TOM and TIM22 channels, creating a ‘clog’ that prevented other essential proteins from reaching the mitochondria. As a result, the yeast cells died. Mutant forms of the human ANT1 protein also clogged up the TOM and TIM22 channels of human cells in a similar way.

Further experiments focused on mice genetically engineered to produce a “super-clogger” version of the mouse equivalent of ANT1. The animals soon developed muscle and neurological conditions similar to those observed in human diseases associated with ANT1.

The findings of Coyne et al. suggest that certain genetic mutations in the gene encoding the ANT1 protein cause disease by blocking the transport of other proteins to the mitochondria, rather than by directly affecting ANT1’s nucleotide trnsport role in the cell. This redefines our understanding of diseases associated with mitochondrial proteins, potentially altering how treatments for these conditions are designed.

Introduction

Mitochondria are essential organelles responsible for a wide range of cellular functions. To carry out these functions, they are equipped with a proteome of 1,000–1,500 proteins (Sickmann et al., 2003; Pagliarini et al., 2008; Morgenstern et al., 2017; Morgenstern et al., 2021). The vast majority of these proteins are encoded by the nuclear genome, synthesized in the cytosol, and sorted into one of the four mitochondrial sub-compartments, namely the outer mitochondrial membrane (OMM), intermembrane space (IMS), inner mitochondrial membrane (IMM), and the matrix (Neupert and Herrmann, 2007; Endo and Yamano, 2009; Chacinska et al., 2009; Wiedemann and Pfanner, 2017). The entry gate by which >90% of mitochondrial proteins enter mitochondria is the translocase of the outer membrane (TOM) complex. Therefore, proper function of the TOM complex is paramount for mitochondrial function and cell viability.

After passage through the TOM complex, specialized protein translocases transport preproteins into the mitochondrial sub-compartments (Neupert and Herrmann, 2007; Endo and Yamano, 2009; Chacinska et al., 2009; Wiedemann and Pfanner, 2017; de Marcos-Lousa et al., 2006). The import of mitochondrial carrier proteins to the protein-dense IMM is particularly challenging. Molecular chaperones like Hsp70 and Hsp90 target these highly hydrophobic proteins through the cytosol to the Tom70 receptor, which is associated with the TOM complex on the mitochondrial surface (Young et al., 2003; Bhangoo et al., 2007). After transport through the channel-forming Tom40 subunit of the TOM complex, the heterohexameric small TIM chaperones (i.e. the Tim9-Tim10 complex) transport the preprotein to the carrier translocase of the inner membrane (TIM22 complex) (Koehler et al., 1998; Sirrenberg et al., 1998; Wiedemann et al., 2001; Truscott et al., 2002; Webb et al., 2006; Weinhäupl et al., 2018; Ellenrieder et al., 2019). The carrier translocase is a multisubunit complex that inserts carrier proteins into the IMM in a membrane potential (Δψ)-dependent manner (Sirrenberg et al., 1996; Kerscher et al., 1997; Kovermann et al., 2002; Rehling et al., 2003; Zhang et al., 2021; Qi et al., 2021). The central subunit Tim22 integrates carrier proteins into the IMM (Rehling et al., 2003; Peixoto et al., 2007). It associates with different partner proteins in yeast and human mitochondria. In yeast, Tim18 and Sdh3 are required for assembly and stability of the carrier translocase, whereas Tim54 tethers the small TIM chaperones (Tim9-Tim10-Tim12 complex) to the translocase (Wagner et al., 2008; Gebert et al., 2011). In human cells, TIM22 associates with the acylglycerol kinase (AGK) and TIM29, which are both required for full import capacity (Kang et al., 2016; Callegari et al., 2016; Kang et al., 2017; Vukotic et al., 2017). Human TIM22 also associates with the TIM9-TIM10 complex (Qi et al., 2021).

Several diseases are associated with mutations directly affecting the protein import machinery of the carrier pathway. For example, mutations in TIMM8A have been found in patients suffering Mohr-Tranebjaerg syndrome/deafness dystonia syndrome (Koehler et al., 1999). Mutations in TOMM70 and TIMM22 have been linked to a devastating neurological syndrome and a mitochondrial myopathy, respectively (Pacheu-Grau et al., 2018; Dutta et al., 2020; Wei et al., 2020). Mutations in another TIM22 complex subunit, AGK, has been associated with the Senger’s syndrome marked by cataracts, hypertrophic cardiomyopathy, skeletal myopathy, and exercise intolerance (Kang et al., 2017; Vukotic et al., 2017; Mayr et al., 2012). Clearly, defective protein import is linked to neurological and musculoskeletal diseases. However, whether protein import defects contribute to diseases not directly related to mutations in the core protein import machinery is unclear.

Here, we show that pathogenic missense mutations in a mitochondrial carrier protein, ADP/ATP translocase 1 (ANT1) or ADP/ATP carrier 2 (Aac2) in yeast, cause arrest of the protein at the translocases during import into mitochondria. This effectively ‘clogs’ the protein import pathway to obstruct general protein import and is associated with muscle and neurological disease in mice. Our findings demonstrate that global protein import is vulnerable to missense mutations in mitochondrial preproteins, and also provide strong evidence that protein import clogging contributes to neurological and muscular syndromes caused by dominant mutations in Ant1 (Kaukonen et al., 2000; Siciliano et al., 2003; Simoncini et al., 2017).

Results

Super-toxic Aac2p mutants dominantly kill cells

We previously showed that four pathogenic ANT1 variants modeled in the yeast Aac2p (Figure 1A) share numerous dominant phenotypes including cold sensitivity, mitochondrial DNA (mtDNA) instability, a propensity to misfold inside mitochondria, and hypersensitivity to low Δψ conditions (Wang et al., 2008; Liu et al., 2015; Coyne and Chen, 2019). We reasoned that if the mutant proteins share a common mechanism of toxicity that drives these phenotypes, then combining mutations into a single protein may enhance toxicity. To test this, we transformed the wild-type M2915-6A yeast strain with centromeric plasmids expressing wild-type, single and double mutant aac2 alleles, and selected for Ura+ transformants on glucose medium. We found that transformants expressing aac2M114P,A128P and aac2M114P,A137D formed smaller colonies on the selective medium at 25°C relative to wild-type AAC2 and single mutant alleles (Figure 1B). Strikingly, transformants expressing aac2A106D,M114P, aac2A106D,A128P, aac2A106D,A137D, and aac2A128P,A137D were unable to form visible colonies at 25°C. Growth of transformants expressing some of the double mutants were improved at 30°C (Figure 1—figure supplement 1A). These data suggest that combining missense mutations into a single Aac2 protein increases toxicity, even when expressed from a centromeric vector.

Figure 1 with 1 supplement see all
Super-toxic Aac2p mutants dominantly kill cells.

(A) Schematic showing the location of pathogenic mutations in transmembrane α-helices 2 and 3 of the ADP/ATP translocator in human (ANT1) compared with mouse (Ant1) and yeast (Aac2p). adPEO, autosomal dominant progressive external ophthalmoplegia. (B) Expression of double mutant aac2 alleles is highly toxic. The yeast M2915-6A strain was transformed with the centromeric vector pRS416 (URA3) expressing wild-type or mutant aac2 alleles and transformants were grown on selective glucose medium lacking uracil at 25°C for 3 days. (C) Growth of yeast cells after serial dilution, showing dominant toxicity of aac2A128P,A137D that is integrated into the genome in the W303-1B strain background. YPD, yeast peptone dextrose medium; YPGE, yeast peptone glycerol ethanol medium. (D) The aac2A128P,A137D allele dominantly increases the frequency of ‘petite’ colonies, which are white. This indicates mitochondrial DNA (mtDNA) destabilization. (E) ‘Petite’ frequencies of yeast strains expressing the mutant alleles of aac2. The strains were first grown in YPD medium for 24 hr before being plated on YPD medium for scoring ‘petite’ colonies. (F) Immunoblot analysis showing extremely low levels of Aac2pA128P,A137D (lower panels) Ilv5p was used as a loading control for mitochondrial protein. Total protein determined with total protein stain (LI-COR). Short, short exposure; Long, long exposure. Upper panel, quantitation from three independent experiments. Aac2p values were normalized by Ilv5p to control for mitochondrial content, and data were represented as relative to wild-type; * indicates p<0.05, ***p<0.001, ****p<0.0001 from one-way ANOVA with Tukey’s multiple comparisons test. Data represented as mean ± SEM.

We integrated a single copy of aac2A128P,A137D into the genome of both AAC2 and aac2Δ strains in the W303-1B background. We chose this strain background for two reasons: first, it is more tolerant of mutant aac2 expression (Wang et al., 2008); and second, mutant aac2 expression is not ρ0-lethal in this background (like it is in M2915-6A and BY4741), meaning that cells expressing mutant aac2 alleles can survive the loss of mtDNA. This allows us to score mtDNA instability via quantitation of smaller white colonies (‘petites’) that form when mtDNA is mutated or depleted (Chen and Clark-Walker, 2000). We found that growth of cells co-expressing aac2A128P,A137D and AAC2 is reduced on glucose medium (Figure 1C), and they form petites at a much higher frequency compared with those expressing the aac2A128P and aac2A137D single mutant alleles (Figure 1D–E). In the aac2Δ background, neither Aac2pA137D nor Aac2pA128P,A137D supported respiratory growth (Figure 1C), consistent with the A123D/A137D mutation eliminating nucleotide transport activity (Palmieri et al., 2005). Cells expressing only aac2A128P,A137D were barely viable, and cell growth was completely inhibited at 25°C even on glucose medium (Figure 1C). This is in sharp contrast to the AAC2-null strain, which strongly supports a dominant effect of aac2A128P,A137D on cell viability.

Interestingly, we found that Aac2pA128P,A137D accumulates to just 4.7% of wild-type Aac2 levels (Figure 1F). We did not observe any accumulation of aggregated Aac2pA128P,A137D (Figure 1—figure supplement 1B–C), indicating that Aac2pA128P,A137D degradation (see below), rather than aggregation, is the likely explanation for low protein recovery. Taken together, the data indicate that aac2A128P,A137D imparts potent toxicity through a mechanism that is independent of nucleotide transport.

Super-toxic Aac2 proteins clog the TOM complex

We hypothesized that the highly toxic double mutant Aac2p is arrested during mitochondrial protein import thereby clogging protein import. To test this, we imported 35S-labeled Aac2p variants into wild-type mitochondria and analyzed the import reaction by blue native polyacrylamide gel electrophoresis (BN-PAGE) followed by autoradiography (Figure 2A; Ellenrieder et al., 2019; Ryan et al., 1999). We did not observe a significant reduction in the amount of IMM-inserted ‘mature’ Aac2pA128P or Aac2pA137D compared with wild-type Aac2p, suggesting no obvious defect for importing into wild-type mitochondria that are fully energized in vitro. In contrast, integration of the double mutant Aac2pA128P,A137D into the IMM was reduced by >70% (Figure 2A–B). To determine whether the mutant preprotein can enter mitochondria, we treated the import reaction with proteinase K to digest non-imported preproteins. We found that a significant portion of mutant Aac2p, particularly Aac2pA128P,A137D, is sensitive to proteinase K digestion (Figure 2C–D). These data suggest that either the Aac2pA128P,A137D preprotein is not transported to the TOM complex, or it fails to fully traverse the TOM complex. To decipher between these possibilities, we imported Aac2p variants into mitochondria containing hemagglutinin (HA)-tagged Tom40 for affinity purification of the TOM complex. Indeed, Aac2pA128P,A137D had increased association with Tom40-HA compared with wild-type (Figure 2E–F). These observations suggest that a significant fraction of Aac2pA128P,A137D is arrested at the TOM complex during import into fully energized wild-type mitochondria.

Figure 2 with 1 supplement see all
Super-toxic ADP/ATP carrier 2 (Aac2) proteins clog the translocase of the outer membrane (TOM) complex.

(A) In vitro protein import assay. 35S-labeled Aac2p and mutant variants were imported into wild-type mitochondria for 10 or 20 min and analyzed by blue native electrophoresis and autoradiography. (B) Quantitation from three independent experiments depicted in (A). p-Value from two-way repeated measures ANOVA with Sidak’s multiple comparisons test. (C) 35S-labeled Aac2p and mutant variants were imported into wild-type mitochondria without (upper) or with (lower panel) subsequent proteinase K treatment to degrade non-imported preproteins. Reaction analyzed by SDS-PAGE and autoradiography. (D) Quantitation from three independent experiments depicted in (C). p-Values were calculated as in (B). (E) Preferential association of mutant Aac2p with Tom40-HA. 35S-labeled Aac2p and mutant variants were imported into Tom40-HA mitochondria, followed by anti-HA immunoprecipitation and analysis by SDS-PAGE and autoradiography. (F) Quantitation from three independent experiments depicted in (E). p-Value was calculated with a one-way ANOVA and Dunnett’s multiple comparisons test. (G) Immunoblot analysis showing accumulation of the un-cleaved precursor of Hsp60p (p) in cells expressing chromosomally integrated aac2A128P,A137D. Cells were grown in YPD at 30°C. m, mature (i.e. cleaved). (H) Immunoblot analysis showing accumulation of un-cleaved Hsp60p precursor (p) in cells expressing aac2 alleles from a centromeric vector. Cells were grown in yeast nitrogen-based dextrose media with supplemented casamino acids, lacking uracil at 25°C. (I) Quantitation from three replicates of (H). Aac2p values normalized by the mitochondrial protein Ilv5p, and then normalized to vector-transformed samples. p-Values were calculated as in (F). Data represented as mean ± SEM.

Next we wondered whether Aac2pA128P,A137D clogs the TOM complex in vivo. If it does, we would expect the accumulation of unprocessed mitochondrial preproteins that travel through the TOM complex, which would include the vast majority of intramitochondrial proteins including both TIM22 and TIM23 substrates. Indeed, precursor of the mitochondrial matrix protein Hsp60p was readily detectable in cells expressing Aac2pA128P,A137D despite extremely low levels of the mutant protein (Figure 2G). We also acutely expressed aac2 from galactose-inducible GAL10 promoter to limit possible indirect effects on protein import efficiency. As expected, expression of mutant aac2 from the GAL10- promoter was highly toxic (Figure 2—figure supplement 1A). Importantly, the timing of Hsp60p precursor accumulation (Figure 2—figure supplement 1B) was coupled with the induction of the mutant Aac2p (Figure 2—figure supplement 1C). The data suggest that the mutant Aac2p directly affects the import of other mitochondrial proteins in vivo.

We extended our analysis to include additional single and double mutant aac2 alleles (see Figure 1A). In transformants expressing double mutant aac2 alleles from a centromeric vector, the level of un-cleaved Hsp60p correlated with toxicity (Figure 2H, see also Figure 1B and Figure 1—figure supplement 1A). Moreover, total Aac2p levels were reduced by all double mutant Aac2p variants except the least toxic Aac2pM114P,A137D (Figure 2H–I). This suggests impaired biogenesis of endogenous wild-type Aac2p. The extent of endogenous Aac2 reduction also correlated with cell toxicity. These data further support the idea that protein import clogging underlies the super-toxicity of Aac2p double mutant.

Yme1p, but not the proteasome, contributes substantially to Aac2pA128P,A137D degradation

We hypothesized that the proteasome should degrade Aac2A128P,A137D as it clogs at the TOM complex. To test this, we first generated strains lacking the drug efflux pump Pdr5p, which allows accumulation of the proteasome inhibitor MG132 in yeast cells. Proteasome inhibition by MG132 was confirmed by the accumulation of Sml1p, a labile proteasome substrate (Figure 3A; Andreson et al., 2010). The experiment was performed in aac2Δ background to facilitate Aac2pA128P,A137D detection. Interestingly, we found that MG132 failed to increase the steady-state level of Aac2pA128P,A137D (Figure 3A). As an orthogonal approach, we tested whether proteasome dysfunction induced by two temperature-sensitive mutants, ump1Δ and pre9Δ, could increase Aac2pA128P,A137D levels, and observed no effect (Figure 3B–C). Cyclohexamide chase following a galactose-induced Aac2pA128P,A137D synthesis confirmed that the mutant protein is unstable compared with the wild-type Aac2p (Figure 3D–E). Consistently, MG132 failed to stabilize acutely induced Aac2pA128P,A137D. These data suggest that little, if any, Aac2pA128P,A137D is degraded by the proteasome.

Degradation of Aac2pA128P, A137D by Yme1p.

(A) Effect of MG132 on the steady-state level of Aac2A128P, A137D in a strain disrupted of PDR5 and AAC2. Cells were first grown in YPD medium at 30°C for 4 hr before MG132 was added at the indicated concentrations. Cells were cultured for another 2 or 4 hr before being harvested for western blot analysis. Sml1 was used as a control for proteasome inhibition. (B) Temperature-sensitive phenotype of ump1Δ and pre9Δ cells. Cells were grown at the indicated temperatures for 2 days before being photographed. (C) Western blot analysis showing that ump1Δ and pre9Δ do not affect the steady-state level of Aac2pA128P, A137D. Cells were grown in YPD medium for 2 and 4 hr at the restrictive temperature (37°C) before being analyzed for Aac2pA128P, A137D levels. TPS, total protein staining. (D) Western blot showing the stability of Aac2pA128P, A137D after cycloheximide (Cyh) chase in cells disrupted of YME1 or treated with MG132, following GAL10-induced synthesis of Aac2pA128P, A137D in galactose medium at 30°C for 4 hr. (E) Quantification of data for the turnover rate of Aac2pA128P, A137D (Mut) and its wild-type control (WT) depicted in (D). Aac2 levels were first normalized by total protein stain and then plotted as values relative to time zero. Depicted are mean values ± SEM from three independent experiments. p-Value was calculated with a two-way repeated measures ANOVA with Tukey’s multiple comparisons test to compare genotypes at time = 120 min.

Next we tested whether an intramitochondrial proteolytic system is responsible for Aac2pA128P,A137D degradation. One likely candidate is Yme1p, which is an IMS-facing AAA protease whose genetic deletion is synthetically lethal with single mutant aac2 variants (Wang et al., 2008). Indeed, we found that Aac2pA128P,A137D is significantly but not fully stabilized in cells disrupted of YME1 (Figure 3D–E). Inhibition of proteasomal function with MG132 does not significantly stabilize Aac2pA128P,A137D in yme1Δ cells. These data strongly suggest that Yme1p plays an important role in the degradation of Aac2pA128P,A137D, possibly serving as a quality control mechanism for degrading stalled import substrates in the vicinity of TIM22 and/or the TOM complex (Wu et al., 2018).

Aac2pA128P accumulates along the carrier import pathway and induces protein import stress

The single mutant Aac2p variants did not show significantly reduced import in vitro and only a mild accumulation at the TOM complex (Figure 2A–B; E-F). However, acute overexpression of the single mutant Aac2pA128P led to accumulation of the precursor form of Hsp60p, consistent with clogging of the translocation apparatus (Figure 2—figure supplement 1B). To explore an arrest of Aac2pA128P at protein translocases, we performed affinity purification Aac2p-HIS6 and Aac2pA128P-HIS6 followed by a quantitative proteomic comparison of the co-purified proteins (Figure 4A–B; Figure 4—figure supplement 1A–C). Numerous proteins preferentially co-purify with Aac2pA128P-HIS6 over Aac2p-HIS6 (Figure 4—source data 1), which were enriched for chaperones involved in targeting of mitochondrial preproteins through the cytosol (Figure 4C; Figure 4—figure supplement 1D–F). Aac2pA128P also has increased association with the TOM complex, as well as the translocase of the inner membrane (TIM22) complex that is responsible for the Δψ-dependent insertion of carrier proteins into the IMM (Figure 4C). The association with Tim22p was confirmed by immunoblotting (Figure 4—figure supplement 1G–H). Expectedly, there is no increased association of Aac2pA128P with Tim23p, which is known to associate with wild-type Aac2p and is not involved in its import (Figure 4—figure supplement 1I; Dienhart and Stuart, 2008). Repeat affinity purification, except with double the ionic strength in the binding buffer, reproduced these data (Figure 4—figure supplement 2A–E, Figure 4—figure supplement 2—source data 1).

Figure 4 with 3 supplements see all
Aac2pA128P accumulates along the carrier import pathway and induces protein import stress.

(A) Growth of cells after serial dilution showing that aac2A128P-HIS6 is toxic at 25°C on glucose medium in an M2915-6A-derived strain. (B) Schematic of our approach to identify aberrant protein-protein interactions of Aac2pA128P-HIS6. (C) Co-purified proteins significantly enriched in Aac2pA128P-HIS6 eluate compared with Aac2p-HIS6. See Materials and methods for details on abundance value calculation. FDR-corrected p-values depicted are from multiple t test analysis. Lower panel is a schematic of the mitochondrial carrier protein import pathway. (D) Absolute quantities of Aac2p and Tim22p in Aac2p-HIS6 and Aac2pA128P-HIS6 eluates, as determined by parallel reaction monitoring (PRM) mass spectrometry. p-Value was calculated with Student’s t test. (E–F) Tetrad analysis demonstrating lethality of aac2A128P expression with genetic defects in carrier protein import in the M2915-6A strain background. Cell were grown on YPD at 30°C. Data depicted as mean ± SEM.

Figure 4—source data 1

Proteins that preferentially interact with Aac2pA128P-HIS6, compare with Aac2-HIS6 in low-salt conditions.

https://cdn.elifesciences.org/articles/84330/elife-84330-fig4-data1-v2.xlsx

To quantitate the increased association of Aac2pA128P to Tim22p relative to the wild-type Aac2p, we performed targeted quantitative proteomics (Figure 4—figure supplement 2F). We found that ~5 mmoles of Tim22p are pulled down per mole of Aac2pA128P-HIS6 (Figure 4D). If 5 mmoles per mole of Aac2pA128P molecules are bound to Tim22p in vivo, this could theoretically occupy ~85% of Tim22p channels, as Aac2p is present at ~188,000 molecules per cell and Tim22p at just 1100 (Morgenstern et al., 2017). Taken together, the data suggest that Aac2pA128P is stalled at both the TOM and TIM22 complexes during import.

Genetic and transcriptional analyses provided additional support for a clogging activity associated with Aac2pA128P. First, we observed that aac2A128P-expressing cells are sensitive to genetic perturbation to the carrier import pathway. TOM70 and TIM18 are two non-essential genes directly involved in the import of mitochondrial carrier proteins such as Aac2, serving as an import receptor at the TOM complex and a component of the TIM22 complex, respectively (Young et al., 2003; Wagner et al., 2008). We found that meiotic segregants combining aac2A128P expression with tom70Δ or tim18Δ form barely visible microcolonies at 30°C on glucose medium (Figure 4E–F). It was recently proposed that Tom70p’s primary function is to recruit chaperones to the mitochondrial surface to prevent proteotoxicity in this protein-dense region (Backes et al., 2021). Thus, deletion of TOM70 may affect aac2A128P cells by two mechanisms: first, by further reducing carrier protein import and subsequently by impairing cytosolic proteostasis in the context of mitochondrial preprotein overaccumulation in the cytosol (Wang and Chen, 2015).

Next, we wondered whether the cellular response to acute Aac2pA128P expression could be indicative of protein import stress. To test this, we surveyed expression levels of a panel of genes known to be upregulated by clogging the TOM complex. These experiments were performed in the BY4742 strain background, in which expression of aac2A128P is incompatible with mtDNA loss (Figure 4—figure supplement 3A), thus minimizing any transcriptional effects secondary to mtDNA loss. We found that CIS1 transcript levels are acutely increased upon expression of aac2A128P, but not of AAC2, from a GAL10 promoter (Figure 4—figure supplement 3B). CIS1 upregulation is the hallmark of the mitochondrial compromised protein import response (mitoCPR) (Weidberg and Amon, 2018). We also found that RPN4, HSP82, SSA3, and SSA4 are transiently upregulated (Figure 4—figure supplement 3C–F), consistent with previously published gene activation patterns induced by a synthetic mitochondrial protein import clogger (Boos et al., 2019). Thus, transcriptional responses further suggest that Aac2pA128P clogs the protein import machinery.

Accumulating evidence suggests a functional crosstalk between the mitochondrial protein translocases and phospholipid biosynthesis/trafficking pathways (Hoffmann and Becker, 2022; Garg et al., 2022). Interestingly, we found that aac2A128P,A137D expression renders cells hypersensitive to defects in phospholipid homeostasis. We expressed aac2A128P,A137D in cells impaired in production of cardiolipin (CL) (pel1Δ) and phosphatidylethanolamine (PE) (psd1Δ). Expression of aac2A128P,A137D strongly inhibited the growth of cells in these genetic backgrounds (Figure 4—figure supplement 3G–H). The data are consistent with previous observations that CL and PE facilitate mitochondrial protein import (Becker et al., 2013; Gebert et al., 2009). Alternatively, mitochondrial protein import clogging may affect phospholipid homoeostasis, which synergizes with pel1Δ and psd1Δ to cause cell lethality.

ANT1A114P and ANT1A114P,A123D clog mitochondrial protein import in human cells

We introduced equivalent mutations in human SLC25A4 (encoding the ANT1) with a C-terminal HA-tag, and transiently expressed the mutant proteins in HeLa cells (see Figure 1A). Like in yeast, combining missense mutations in Ant1 dramatically reduced steady-state protein levels, suggesting impaired ANT1 biogenesis (Figure 5A). Most strikingly, the level of ANT1A114P,A123D, equivalent to the yeast Aac2pA128P,A137D, is only 2.2% of the wild-type. Three lines of evidence suggest that ANT1A114P and ANT1A114P,A123D obstruct protein import into mitochondria. First, immunoprecipitation of ANT1A114P-HA demonstrated that the mutant protein has increased interactions with components of both TOM and TIM22 complexes (Figure 5B–C), as observed with its yeast ortholog Aac2pA128P. A meaningful immunoprecipitation of Aac2pA128P,A137D-HA was precluded by the extremely low-level accumulation of the mutant protein. Second, protease protection assay demonstrated that 42% and 52% of total ANT1A114P and ANT1A114P,A123D proteins are exposed on the OMM, whereas the wild-type ANT1 is protected from proteinase K digestion (Figure 5D–E). This is consistent with in vitro import studies of the Aac2p variants in yeast mitochondria (Figure 2A–F). Third, proteomics demonstrated that mitochondrial proteins accumulate in the cytosol of SLC25A4 p.A114P and SLC25A4 p.A114P,A123D -transfected cells, compared with wild-type Ant1-transfected cells (Figure 5F). Moreover, mitochondrial matrix proteins were significantly enriched in the cytosol of ANT1A114P,A123D versus ANT1-expressing cells (FDR<3×10–6) (Figure 5G; Figure 5—source data 1), corroborating that the double mutant Aac2p impairs global protein import. We therefore conclude that ANT1A114P and ANT1A114P,A123D clog protein import in human cells, with ANT1A114P,A123D having enhanced clogging activity despite low protein levels.

Figure 5 with 1 supplement see all
ANT1A114P and ANT1A114P,A123D clog mitochondrial protein import in human cells.

(A) Combining pathogenic mutations in ADP/ATP translocase 1 (ANT1) strongly reduces protein levels, as indicated by immunoblot analysis of ANT1-hemagglutinin (HA) levels 24 hr after transfecting HeLa cells. ANT1 variant levels were normalized by TFAM, then plotted as relative to wild-type level. * indicates p<0.05, ****p<0.0001 from one-way ANOVA with Dunnett’s multiple comparisons test. (B) Immunoprecipitation (IP) of ANT1-HA and ANT1A114P-HA from transiently transfected HeLa cells followed by immunoblot analysis, showing that ANT1A114P has increased interaction with the protein import machinery like its yeast ortholog Aac2pA128P. (C) Quantitation from four independent IP, one of which is depicted in (B). p-Values were calculated with a Student’s t test. (D) Immunoblot analysis following protease protection assay showing that ANT1A114P and ANT1A114P,A123D are sensitive to proteinase K (PK) in isolated mitochondria. Swelling in hypotonic buffer was used to burst the outer membrane, and Triton X-100 was used to disrupt all membranes. OM, outer membrane; IMS, intermembrane space; IM, inner membrane. (E) Quantitation of the wild-type and mutant ANT1 pools that are protected from PK degradation in intact mitochondria. All HA levels were normalized by TFAM, then plotted as relative to its untreated sample. Replicates from three independent transfections. p-Values were calculated with a one-way ANOVA and Holm-Sidak’s multiple comparisons test. (F) ANT1A114P and ANT1A114P,A123D obstruct general mitochondrial protein import. Proteomics of the cytosolic fraction of transfected HeLa cells reveals increase in mitochondrial proteins caused by ANT1A114P-HA and ANT1A114P,A123D-HA expression relative to ANT1-HA. p-Values were calculated with a Student’s t test of the average abundance levels of each mitochondrial protein. (G) Volcano plot comparing the cytosolic proteome of SLC25A4 p.A114P,A123D vs SLC25A4-transfected HeLa cells. Data represented as mean ± SEM. (H) Enrichment analysis of proteins significantly increased in the cytosol of SLC25A4 p.A114P,A123D-transfected HeLa cells. Depicted are the most significant enriched protein groups generated from three different databases: GO: Biological Process (top), KEGG pathway (middle), and GO: Molecular Function (bottom).

Figure 5—source data 1

Proteomic comparison of the cytosolic fraction of SLC25A4 p.A114P,A123D- versus SLC25A4-transfected HeLa cells.

https://cdn.elifesciences.org/articles/84330/elife-84330-fig5-data1-v2.xlsx
Figure 5—source data 2

Uncropped Western blots from Figure 5A,B,D.

https://cdn.elifesciences.org/articles/84330/elife-84330-fig5-data2-v2.zip

Regarding the function of the non-mitochondrial proteins increased in the cytosol of ANT1A114P,A123D-expressing cells, we found that the most significantly enriched Biological Process, Molecular Function, and KEGG pathway were ‘chaperone’, ‘stress response’, and ‘protein processing in the ER’ (Figure 5H). This may represent a stress response directed toward the increased mitochondrial protein burden in the cytosol.

An alternative explanation for reduced protein import in mutant Ant1-transfected cells is general mitochondrial damage and/or apoptosis activation leading to reduced Δψ, which would also cause a reduction in protein import. However, neither ANT1A114P,A123D nor its single mutant counterparts reduced Δψ or significantly increased cell death compared with wild-type (Figure 5—figure supplement 1). Thus, the effect on protein import by ANT1A114P and ANT1A114P,A123D is likely due to their physical retention in the import pathway rather than due to reduction of Δψ. It is interesting that ANT1A114P,A123D does not increase apoptosis in the immortalized HeLa cells, despite clearly clogging protein import. This may be related to the transient nature of ANT1 expression or an inherent resistance of the immortalized cells to cell death.

Ant1A114P,A123D causes dominant muscle and neurological disease in mice

We generated a knock-in Slc25a4 p.A114P,A123D/+ (expressing Ant1A114P,A123D) mouse line to model protein import clogging in vivo (Figure 6A–B). We observed a neurodegeneration phenotype that culminates in paralysis in some Slc25a4 p.A114P,A123D/+ mice (Figure 6C; Video 1; Video 2). This phenotype occurred in only four Slc25a4 p.A114P,A123D/+ mice, with a penetrance of 3.4% among the heterozygous mice that have reached 15 months of age. The presenting symptom is typically altered gait after the age of 11 months, followed by weight loss and death within 2–3 weeks of symptom onset. Histological analysis of the lumbar spinal cord in a symptomatic Slc25a4 p.A114P,A123D/+ mouse demonstrated dissolution of Nissl substance in the cell bodies of ventral horn neurons (Figure 6D), consistent with motor neuron degeneration (Bodian and Mellors, 1945). Neurodegeneration in the spinal cord was also indicated by GFAP accumulation by immunofluorescence and immunoblotting (Figure 6E–G). Transmission electron microscopy of ventral horn neurons revealed loss of cristae density of mitochondria, which suggests defects in mitochondrial biogenesis (Figure 6H).

Slc25a4 p.A114P,A123D/+ mouse generation and neurodegeneration.

(A) Schematic of the strategy by which the knock-in Slc25a4 p.A114P,A123D/+ knock-in mice were generated. E1–E4, exons 1–4 of Slc25a4. In gray is the inserted cDNA containing two missense mutations in exon 2, followed by the endogenous 3’ UTR. Lox gtF and gtR indicate genotyping primers. (B) Agarose gel electrophoresis of PCR genotyping using genotyping primers indicated in (A). Fl, floxed. (C) Ascending paralytic phenotype of an Slc25a4 p.A114P,A123D/+ mouse at 11 months of age, and its wild-type littermate. Arrows point to paralyzed hindlimbs. (D) Nissl-stained lumbar spinal cord neuron of a symptomatic Slc25a4 p.A114P,A123D/+ mouse and wild-type littermate. This neuron shows loss of Nissl substance and blurring of nuclear boundaries, process known as ‘chromatolysis’, which indicates neuron degeneration. (E) Indirect immunofluorescence for the astrocyte marker glial fibrillary acidic protein (GFAP) indicating spinal cord gliosis in a symptomatic Slc25a4 p.A114P,A123D/+ mouse. G, gray matter; W, white matter. (F) Immunoblot analysis of lumbar spinal cord lysate confirmed increase in GFAP in symptomatic Slc25a4 p.A114P,A123D/+ mice. (G) Quantitation from (F) showing significant increase in GFAP in the spinal cord of a symptomatic Slc25a4 p.A114P,A123D/+ mouse indicating neuroinflammation. p-Value was calculated from Student’s t test. (H) Transmission electron microscopy of a ventral horn neuron of a symptomatic Slc25a4 p.A114P,A123D/+ mouse and wild-type littermate control.

Video 1
Paralytic phenotype of an Slc25a4 p.A114P,A123D/+ mouse at the age of 12 months.
Video 2
Paralytic phenotype of an Slc25a4 p.A114P,A123D/+ mouse at the age of 16 months.

Dominant SLC25A4-induced diseases primarily affect skeletal muscle in addition to low-penetrant neurological involvement (Kaukonen et al., 2000; Siciliano et al., 2003; Simoncini et al., 2017; Kaukonen et al., 1999; Napoli et al., 2001; Deschauer et al., 2005). Key muscle features include mildly reduced mitochondrial respiratory function, COX-deficient muscle fibers, and muscle weakness. Consistent with clinical phenotypes, we found that the maximal respiratory rate (state 3) of Slc25a4 p.A114P,A123D/+ muscle mitochondria was reduced by ~20% and ~30% when utilizing complex I in 9- and 24-month-old mice, respectively (Figure 7A). The respiratory control ratio, which is the single most useful and sensitive general measure of energy coupling efficiency (Brand and Nicholls, 2011), was reduced by ~31% and ~42% in young and old mice, respectively. Surprisingly, when complex I is inhibited and complex II substrate (succinate) is present, maximal respiration is increased by ~23% in the mutant mice at 9 months of age (Figure 7B). This result is important because it confirms that ATP/ADP transport is not a limiting factor for sustaining a high respiratory rate in Slc25a4 p.A114P,A123D/+ mice.

Ant1A114P,A123D (encoded by Slc25a4 p.A114P,A123D) causes a dominant mitochondrial myopathy in mice.

(A) Respirometry of isolated skeletal muscle mitochondria with complex I stimulated by glutamate (glu) and malate (mal). State 3, maximal respiratory rate after addition of ADP; state 4, oligomycin (oligo)-inhibited respiratory rate; respiratory control ratio = state 3/state 4. N=6 mice/genotype at 9 months of age; n=4 mice per genotype at 24 months of age. Three measurements were taken per mouse. p-Values were derived from repeated measures ANOVA with measurement order as the within-subjects variable. Data from two age groups were analyzed independently. FCCP, trifluoromethoxy carbonylcyanide phenylhydrazone. (B) Respirometry of isolated skeletal muscle mitochondria with complex II stimulated by succinate and complex I inhibited by rotenone. N=2 mice/genotype at 9 months of age, 4 measurements/mouse; n=4 mice/genotype at 24 months of age, 3 measurements/mouse. Data analyzed as in (A). (C) Soleus muscles stained with hematoxylin and eosin (H&E) showing smaller myofibers in 30-month-old Slc25a4 p.A114P,A123D/+ mice. (D) Feret’s diameter analysis of H&E stained soleus in (C) reveals atrophy in Slc25a4 p.A114P,A123D/+ mice. At least 340 myofibers were measured per soleus. Myofiber diameters were binned into 5 μm ranges and plotted as % of total. N=3 mice/genotype. Data analyzed by two-way ANOVA with Sidak’s multiple comparisons test. (E) Succinate dehydrogenase (SDH) histochemical activity staining of the soleus showing abnormal fibers that stain for SDH peripherally but are pale internally (arrows). (F) Histochemical cytochrome c oxidase (COX) and SDH sequential staining of the soleus shows abnormal fibers that stain for COX peripherally, but do not stain for COX or SDH internally. (G) Quantitation of abnormal COX fibers shown in (F). p-Value was calculated from Student’s t test. (H) Forelimb grip strength is reduced in 30-month-old Slc25a4 p.A114P,A123D/+ mice. p-Value from Student’s t test. (I) Maximal forelimb grip strength is reduced in 30-month-old Slc25a4 p.A114P,A123D/+ mice. p-Value from Student’s t test. Data represented as mean ± SEM.

In addition to mild bioenergetic defect, we detected significant reduction in myofiber diameter in the skeletal muscle from aged Slc25a4 p.A114P,A123D/+ mice (Figure 7C–D). Sequential COX/SDH histochemical assay failed to detect any COX-negative/SDH-positive fibers in the aged Slc25a4 p.A114P,A123D/+ muscles. Instead, we found that ~3.5% of myofibers have reduced COX and SDH activity centrally, but remained COX and SDH-positive in the periphery (Figure 7E–G). Finally, we found that the Slc25a4 p.A114P,A123D/+ mice display muscle weakness (Figure 7H–I). These data demonstrate that Slc25a4 p.A114P,A123D induces a dominant myopathic phenotype.

Ant1A114P,A123D clogs protein import in vivo

For biochemical characterization of the mutant protein, we turned to homozygous Slc25a4 p.A114P,A123D/Slc25a4 p.A114P,A123D mice as to eliminate the detection of confounding wild-type Ant1. We found that the Ant1A114P,A123D protein is virtually undetectable in the total lysates of mouse tissues, suggesting a rapid degradation of the mutant protein as observed in yeast and human cells. (Figure 8A). Only in isolated skeletal muscle mitochondria were we able to detect Ant1A114P,A123D, which was present at ~0.1% of wild-type level (Figure 8B–C). Protease protection assay on isolated muscle mitochondria demonstrated that Ant1A114P,A123D is more sensitive to proteinase K digestion compared with wild-type Ant1 (Figure 8D–E), consistent with import clogging in vivo. Moreover, Smac also appears to be more sensitive to proteinase K in Slc25a4 p.A114P,A123D/Slc25a4 p.A114P,A123D mitochondria, while Mdh2 and Tim23 do not (Figure 8F–H). While this could indicate instability or partial rupture of the OMM in mutant mitochondria, it could also be a result of clogging and imply that clogging does not affect all mitochondrial preproteins equally. Consistent with this concept, our experiments in yeast showed minimal effects of clogging on Ilv5p, a mitochondrial matrix protein, in contrast to Hsp60p (Figure 1F; Figure 2H–I). This could be explained by the observation that the Tom40 β-barrel pore contains several distinct protein paths (Shiota et al., 2015; Araiso et al., 2019).

Figure 8 with 2 supplements see all
Ant1A114P,A123D clogs protein import in vivo.

(A) Immunoblot analysis of tissue lysate showing low Ant1A114P,A123D protein levels in heterozygous and homozygous mice. (B) Immunoblot analysis of isolated muscle mitochondria demonstrating low Ant1A114P,A123D protein levels. (C) Quantitation of Ant1 levels in isolated muscle mitochondria from three mice per genotype, as determined by immunoblotting. Values were normalized to total protein stain and shown as relative to wild-type. (D) Ant1A114P,A123D is more sensitive to proteinase K (PK) than wild-type Ant1 in intact mitochondria. Immunoblot analysis after PK protection assay of isolated muscle mitochondria in isotonic buffer. Ant1A114P,A123D was detected using SuperSignal West Femto Maximum Sensitivity Substrate (top right panel). (E–H) Quantitation from protease protection assay, as shown in (D). n=3 mice per genotype. p-Values were calculated with a two-way ANOVA, showing significant main effect of genotype. Data represented as mean ± SEM. (I) Schematic of tandem mass tagged (TMT) quantitative proteomic analysis. (J) Volcano plot comparing the cytosolic proteome of Slc25a4 p.A114P,A123D/+ vs wild-type skeletal muscle, with mitochondrial proteins highlighted in blue.

Figure 8—source data 1

Proteomic comparison of the cytosolic fraction of 30-month-old skeletal muscle from Slc25a4 p.A114P,A123D/+ versus wild-type mice.

https://cdn.elifesciences.org/articles/84330/elife-84330-fig8-data1-v2.xlsx
Figure 8—source data 2
https://cdn.elifesciences.org/articles/84330/elife-84330-fig8-data2-v2.zip

Finally, to test if Ant1A114P,A123D obstructs general mitochondrial protein import in vivo, we compared the cytosolic proteomes of aged muscle from wild-type and Slc25a4 p.A114P,A123D/+ mice using tandem mass tagged (TMT) quantitative proteomics (Figure 8I). We found a striking global increase in mitochondrial proteins in the cytosol of Slc25a4 p.A114P,A123D/+ muscle (Figure 8J). Among the 75 proteins increased by at least 25% in the cytosol (p<0.05), proteins assigned to the mitochondrion accounted for 45 of them (Figure 8—source data 1). This enrichment was highly significant (FDR<10–33). Taken together, the data suggest that Ant1A114P,A123D clogs general protein import in vivo. Supporting this is a trend of increase in proteasomal subunits and Hsp70 proteins in the cytosol of Slc25a4 p.A114P,A123D/+muscle, which suggests an anti-mPOS response (Figure 8—figure supplement 1A–B).

To evaluate the impact of Slc25a4 p.A114P,A123D expression on mitochondrial proteostasis, we assessed the assembly state of protein complexes and supercomplexes in the inner membrane with native gel electrophoresis. The respiratory complexes and supercomplexes were minimally affected (Figure 8—figure supplement 1C–D), as was the TIM23 complex (Figure 8—figure supplement 1E). To gain a global view of mitochondrial proteostasis, we performed TMT labeling quantitative proteomics on mitochondrial fractions from skeletal muscle. We found minimal changes in the steady-state levels of mitochondrial proteins, including other Ant isoforms (Figure 8—figure supplement 1F, Figure 8—figure supplement 1—source data 1). This may reflect highly efficient degradation of Ant1A114P,A123D, which is suggested by its steady-state level being 0.1% of wild-type (Figure 8B–C). It is also consistent with the subtle bioenergetic defects and the overall mild neuronal and muscular phenotypes associated with the Slc25a4 p.A114P,A123D/+ mice. Interestingly, a focused analysis on mitochondrial proteases and chaperones suggested a subtle but statistically significant increase in the mitochondrial proteostatic machinery (p=2.7 × 10–14, analyzed with a two-way ANOVA; Figure 8—figure supplement 1G), which supports proteostatic stress inside mitochondria.

Another possible explanation for minimal changes in mitochondrial proteins in the context of protein import clogging could be increased protein import TIM22 pathway. Our mitochondrial proteomics dataset suggested global increase in TIM22 pathway components (Figure 8—figure supplement 1H). Immunoblot validation of Tim22 showed an ~40% increase (Figure 8—figure supplement 1I–J). These data may represent chronic adaptation to carrier protein import clogging, though more work is required to rigorously test this hypothesis.

Transcriptional response induced by Slc25a4 p.A114P,A123D expression in mouse muscle

The integrated stress response (ISR) is activated by a diverse range of mitochondrial insults. Surprisingly, we found no evidence of ISR activation downstream of Slc25a4 p.A114P,A123D expression. First, eIF2α phosphorylation, a hallmark of the ISR, was not increased in Slc25a4 p.A114P,A123D/+ mouse skeletal muscle at multiple ages (Figure 8—figure supplement 2A–B). Second, transcriptomic analysis in Slc25a4 p.A114P,A123D/+ skeletal muscle failed to show induction of ISR target genes (Figure 8—figure supplement 2C). Instead, transcription factor enrichment analysis of significantly upregulated genes (q<0.05) revealed activation of pathways controlled by novel transcriptional factors, including FOXO1 and FOXO3 that are involved in metabolic homeostasis and autophagy (Figure 8—figure supplement 2D–E). It appeared that Slc25a4 p.A114P,A123D induces an entirely unique transcriptional signature (Figure 8—figure supplement 2F; Figure 8—figure supplement 2—source data 1), although it remains to be determined whether severe mitochondrial protein import clogging relative to Ant1A114P,A123D induces similar stress responses and/or activates ISR. Among the upregulated genes, Depp1 is known to activate autophagy (Stepp et al., 2014). More importantly, when directly compared to Slc25a4 knockout mice (Morrow et al., 2017), there is very limited overlap in the enrichment profile among significantly altered genes (Figure 8—figure supplement 2G). This indicates that the primary stress in Slc25a4 p.A114P,A123D/+ skeletal muscle is distinct from that in Ant1 knockout mice.

In summary, we found that Ant1A114P,A123D dominantly causes muscle and low-penetrant neurological disease in mice that recapitulates pathological and molecular phenotypes of dominant Ant1-induced diseases in humans. This corroborates the toxicity of Aac2A128P,A137D in yeast (Figure 1), and supports the idea that mitochondrial protein import clogging by a mutant substrate preprotein is pathogenic.

Discussion

Failure in mitochondrial protein import has severe physiological consequences. In addition to defective mitochondrial biogenesis and energy metabolism, it also causes toxic accumulation and aggregation of mitochondrial preproteins in the cytosol, a process termed mitochondrial Precursor Overaccumulation Stress (mPOS) (Wang and Chen, 2015; Coyne and Chen, 2018; Song et al., 2021). To prevent these consequences, many cellular safeguards have been identified that can maintain protein import efficiency and/or mitigate mPOS (Backes et al., 2021; Weidberg and Amon, 2018; Boos et al., 2019; Izawa et al., 2012; Wrobel et al., 2015; Nargund et al., 2015; Izawa et al., 2017; Hansen et al., 2018; Zurita Rendón et al., 2018; Liu et al., 2019; Mårtensson et al., 2019; Su et al., 2019; Ordureau et al., 2020; Phu et al., 2020; Xiao et al., 2021; Shakya et al., 2021; Xin et al., 2022; Sam et al., 2021; Schulte et al., 2023; Dewar et al., 2022; Krämer et al., 2023). So far, studies on mitochondrial stressors that impair protein import have been mainly focused on mutations that directly affect the core protein import machinery, synthetic clogger proteins, or on pharmacological interventions that reduce Δψ (Song et al., 2021). In this report, we have uncovered the first example of naturally occurring missense mutations in an endogenous mitochondrial protein causing toxic protein import clogging. We showed that a single protein without overexpression is sufficient to clog import, thereby inducing robust protein import stress responses, bioenergetic defects, and cytosolic proteostatic stress. The data supports a model in which import clogging kills yeast cells and contributes to muscle and neurological degeneration in mice.

Mitochondrial protein import can be clogged by a mutant mitochondrial preprotein

The mitochondrial carrier protein family is the largest of the transporter families and has highly conserved domain and sequence features across all eukaryotes. Their import into mitochondria is unique in that they assume partially folded conformations called ‘hairpin loops’ that place adjacent hydrophobic α-helices antiparallel with one another (de Marcos-Lousa et al., 2006; Wiedemann et al., 2001). There are three ‘hairpin loop’ modules per carrier preprotein that transit through the Tom40 β-barrel in a loop-first topology, such that the C- and N-termini initially remain exposed to the cytosol. This leaves little wiggle room for the carrier preprotein considering the Tom40 β-barrel can only accommodate the width of up to two α-helical segments at a time (Wiedemann et al., 2001). To get through the Tom40 β-barrel pore, the hydrophobic hairpin loops are thought to be guided by an array of hydrophobic patches on the inside of the pore (Shiota et al., 2015; Araiso et al., 2019). It is in the hydrophobic α-helices of the ADP/ATP translocase ANT1 (Aac2p in yeast) that dominant pathogenic mutations are found, which introduce either a proline or an aspartic acid (see Figure 1). Here, we show that these mutations cause the protein to arrest at or in the TOM complex, thereby partially obstructing general protein import. There are multiple potential mechanisms. First, as the mutations introduce a proline or aspartic acid into the hydrophobic α-helix, it may simply be loss of the necessary hydrophobic interaction with the inner surface of the Tom40 β-barrel pore. Another possibility is that the hairpin loop structure is disrupted such that it becomes too wide to fit through the narrow pore. This may be particularly relevant for the proline mutants which would introduce a kink into the α-helix. A third possibility is that mutant ANT1/Aac2p is not impaired in transit through the pore, but instead is inefficiently chaperoned by the small TIM chaperones in the IMS, leading to ‘backpressure’ in the pathway. Interestingly, carrier preproteins appear to form partial α-helical secondary structures while bound to the Tim9-Tim10 chaperone (Weinhäupl et al., 2018). Future work will be essential to rigorously decipher between these potential mechanisms of clogging.

The data strongly suggest that double mutant Aac2p preferentially clogs the TOM complex (Figure 2). In contrast, co-immunoprecipitation of single mutants Aac2pA128P and ANT1A114P showed increased association with both the TOM and TIM22 complexes. We interpret these data with caution, as it would be unexpected that single mutants would clog at multiple locations. However, it is possible that hairpin loops and/or hydrophobic α-helices are crucial for interaction with both complexes. As the precise mechanism of import and IMM insertion by the TIM22 complex remains unresolved, it would be premature to speculate on potential clogging mechanisms.

One of the most surprising observations of this study was the extreme toxicity of the double mutant clogger proteins that arrest at the TOM complex. The deleterious stressors downstream of clogging are likely to be multifactorial and include respiratory defects and cytosolic protestatic stress via mPOS. While it is possible that general proteostasis is perturbed by Aac2pA128P,A137D saturating proteolytic machinery, our data suggest this is unlikely. The IMM-associated AAA protease Yme1p (and not the proteasome) appears to be a major proteolytic pathway for clogged Aac2pA128P,A137D. That genetic deletion of YME1 is well tolerated in yeast cells suggests that loss of Yme1-based proteolysis does not underlie toxicity of aac2A128P,A137D expression. We propose that clogging of the TOM complex is the primary mechanism of Aac2pA128P,A137D-induced protein import defects and the ensuing cell stress. Importantly, the former is strongly supported by in vitro experiments showing that Aac2pA128P,A137D is defective in translocating through the TOM complex in fully energized, wild-type mitochondria.

In the context of low Aac2pA128P,A137D levels in cells (just 4.7% of wild-type), it is important to note that the ADP/ATP carrier is one of the most abundant proteins in mitochondria, outnumbering the Tom40 pore-forming protein by almost an order of magnitude (Morgenstern et al., 2017; Pfanner et al., 2019). Thus, if ~70% of the reduced level of Aac2pA128P,A137D is actively clogging TOM complexes, this would still occupy ~30% of the Tom40 channels, which may have a considerable effect on general protein import. This reflects the central importance of proper TOM complex function to mitochondrial and cell homeostasis, and also provides physiological justification for the existence of multiple pathways dedicated to unclogging the TOM complex (Weidberg and Amon, 2018; Mårtensson et al., 2019).

It will be interesting to determine whether and how Yme1 plays a general role in quality control of the TIM22 import pathway. Yme1 degradation of Aac2pA128P,A137D could occur at the TOM complex, in association with the small TIMs or at the TIM22 complex. A functional crosstalk between Yme1p and the TIM22 complex has been proposed (Hwang et al., 2007; Kumar et al., 2023). Detailed mechanistic studies are required.

Mitochondrial protein import clogging as a mechanism of disease

A long-standing mystery in the mitochondrial disease field is how dominant mutations in SLC25A4 (encoding adenine nucleotide translocase 1 [Ant1]) cause such a wide spectrum of clinical and molecular phenotypes that are not observed in patients with homozygous null SLC25A4 (Kaukonen et al., 2000; Siciliano et al., 2003; Simoncini et al., 2017; Palmieri et al., 2005; Napoli et al., 2001; Deschauer et al., 2005; Thompson et al., 2016; Echaniz-Laguna et al., 2012; Tosserams et al., 2018; Kashiki et al., 2022). The implication is that the dominant pathogenic ANT1 proteins must have an unknown gain-of-function pathogenic mechanism. Similarly, the phenotypes observed in Slc25a4 p.A114P,A123D/+ mice cannot be explained by haploinsufficiency or a dominant negative effect on ADP/ATP transport for the following reasons. First, as in humans, the neurological and muscle phenotypes in Slc25a4 p.A114P,A123D/+ mice have never been reported in heterozygous Slc25a4 knockout mice despite 25 years of characterization by Doug Wallace and colleagues. Second, mitochondria from Slc25a4 p.A114P,A123D/+ mouse muscle are able to exceed the maximal respiration rate of wild-type mitochondria when complex II is stimulated (Figure 6B), indicating that one functional copy of Ant1 is sufficient to support high respiratory rates without compensatory increase in other Ant isoforms (see Figure 8—figure supplement 1F; Figure 8—figure supplement 1—source data 1; Figure 8—figure supplement 2—source data 1; Figure 8—figure supplement 2F). This is in stark contrast to Slc25a4 knockout, which displays a >30% reduction in complex II-based respiration (Graham et al., 1997). Third, the transcriptional profile in Slc25a4 p.A114P,A123D/+ skeletal muscle is entirely distinct from that in Slc25a4 knockout mice, suggesting different underlying mechanisms of muscle dysfunction (Morrow et al., 2017). Fourth, Ant1 likely functions as a monomer, making a dominant-negative mechanism of adenine nucleotide transport unlikely (Pebay-Peyroula et al., 2003; Ruprecht et al., 2019; Kunji and Ruprecht, 2020). Finally, the yeast equivalent of Ant1A114P,A123D, Aac2pA128P,A137D, is catalytically inactive yet clearly exerts toxicity even in conditions where aac2Δ cells are healthy (Figure 1C). Taken together, the evidence is clear that dominant SLC25A4 mutations cause gain-of-function phenotypes in patients, Slc25a4 p. A114P,A123D/+ mice and yeast models.

In this study, we elucidated a candidate molecular mechanism of dominant Ant1-induced disease using yeast, human cells, and a novel mouse model. We found that pathogenic ANT1 mutations modeled in yeast’s Aac2p cause the protein to clog global mitochondrial protein import leading to respiratory defects, cytosolic proteostatic stress (Wang and Chen, 2015), and loss of cell viability. Combining two mutations into a single Ant1 protein drastically enhanced clogging activity, which tightly correlated with toxicity. We extensively characterized one such ‘super-clogger’ variant, Aac2pA128P,A137D, and found that the main clogging site shifted to the TOM complex, providing an explanation for the increased toxicity. Protein import clogging was strikingly consistent in transfected human cells, as ANT1A114P and ANT1A114P,A123D (ortholog of yeast Aac2pA128P,A137D) physically associated with the TOM and TIM22 complexes, was exposed on the outside of intact mitochondria, and caused mitochondrial protein retention in the cytosol. Even in mice, Ant1A114P,A123D expression correlated with retention of mitochondrial proteins in the cytosol despite minute levels of the mutant protein, presumably due to degradation. Finally, we found that Slc25a4 p.A114P,A123D/+ mice phenocopy ANT1-induced autosomal dominant progressive external opthalmoplegia (adPEO) patients, as Slc25a4 p.A114P,A123D/+ mice exhibit mild OXPHOS defects, partially COX-deficient myofibers, moderate myofiber atrophy, and muscle weakness (Kaukonen et al., 1999; Komaki et al., 2002). The low-penetrant neurodegeneration in Slc25a4 p.A114P,A123D/+ mice may also reflect human disease process, as a neurodegenerative phenotype has been reported in an adPEO patient carrying the SLC25A4 p.A114P allele (Simoncini et al., 2017). Taken together, these data strongly argue that protein import clogging contributes to adPEO in humans, despite Slc25a4 p.A114P,A123D not directly genocopying a particular clinical condition. We do note, however, that additional contributory mechanisms and even SLC25A4 deficiency cannot be completely excluded without in vivo rescue experiments.

In summary, we demonstrate that pathogenic mutations in a mitochondrial carrier protein can cause clogging of the protein import pathway, which induces multifactorial cellular stress and correlates with cell toxicity and disease in mice. These findings uncovered a vulnerability of the mitochondrial protein import machinery to single amino acid substitutions in mitochondrial inner membrane preproteins. There are 53 carrier proteins in human mitochondria that have highly conserved domain organization both within and across species (Palmieri et al., 2011). Mutations altering conformational elements involved in protein import, such as hairpin loops, are likely strongly selected against during evolution. Those persisting in the human population would affect protein import and cause disease, as exemplified by the dominant pathogenic mutations in SLC25A4.

Our work provides just a first glimpse into the amino acid requirements of mitochondrial carrier proteins to maintain compatibility with the mitochondrial protein import machinery. Future work will be crucial to systematically evaluate the amino acid requirements first within ANT1/Aac2p, and then across entire carrier protein family. It will also be important to determine which specific interactions with protein import components are affected by different mutations. Overall, these efforts may enable the prediction of pathogenic mechanisms of future variants of uncertain significance as they arise with more exome sequencing of human patients.

Materials and methods

Key resources table - see Appendix 1.

Yeast growth conditions and genetic manipulation

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Yeast cells were grown using standard media. Genotypes and sources of yeast strains are listed in Key resources table. To generate the AAC2 expression plasmids, we amplified the gene from total genomic DNA and inserted it into pFA6a-ScAAC2-URA3-HIS3/2. The missense mutations were introduced using QuikChange Site-Directed Mutagenesis (Stratagene) and mutations were confirmed by sequencing. Mutant genes were then cloned into pRS416 for expression (Figure 1B) and pFA6a-KanMX6 for placing next to the KAN selection marker. The aac2A128P,A137-KAN cassette was then amplified for integration into the LYS2 locus of the W303-1B strain background. All strains in Figure 1C–E were derived from this strain by standard genetic crosses. All combinations of aac2 mutations were expressed by transforming an equal number of M2915-6A yeast with freshly prepared pRS416-based vectors (URA3) expressing wild-type or mutant aac2. Equal fractions of the transformant culture were plated on selective medium lacking uracil and grown at 25°C or 30°C for 3 days before being photographed (, Figure 1B and S1A ). tom70Δ and tim18Δ strains were generated by amplification of the knockout cassette from genomic DNA of tom70Δ and tim18Δ strains from BY4741 knockout library, followed by transformation into M2915-6A by selecting for G418R. The disruption of the genes was confirmed by PCR using an independent primer pair surrounding the native genomic locus. PRE9 and UMP1 were disruption in strains of W303 background by the insertion of kan. These alleles were then introduced into strains expressing aac2A128P, A137D by genetic crosses. For galactose-induced expression, AAC2 and its mutant alleles were placed under the control of the GAL10 promoter in the integrative vector pUC-URA3-GAL. The resulting plasmids were then linearized by cutting with StuI within the URA3 gene, before being integrated into the ura3-1 locus by selecting for Ura+ transformants. Correct chromosomal integration was confirmed by examining the stability of the Ura+ phenotype and PCR amplification of the URA3 locus. The yme1Δ and pdr5Δ alleles were introduced into these strains either by direct gene disruption or by genetic crosses.

Cell lysis, western blotting, and signal quantitation

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Unless otherwise noted, yeast cells were lysed and prepared for SDS-PAGE as previously described (Chen, 2001). HeLa cells and mouse tissues were lysed with RIPA buffer containing 1× HALT Protease and Phosphatase Inhibitor Cocktail (Thermo Fisher) and prepared for SDS-PAGE with Laemmli buffer. Standard procedures were used for chemiluminescent western blot detection of proteins. Membranes were imaged using a LI-COR Odyssey imager and signals quantitated in the associated ImageStudio software.

Detergent extraction of potential insoluble protein

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We took two approaches to detecting potentially aggregated Aac2pA128P,A137D. First, spheroplasts were generated using zymolyase followed by lysis in sample buffer (containing 8 M urea+5% SDS) with or without 2% Sarkosyl. Second, the insoluble material from our typical lysis procedure (see above) was treated with different detergents. For formic acid, pellet was resuspended in 100% formic acid, incubated at 37°C for 70 min, followed by speed-vac drying of the sample and resuspension in sample buffer for SDS-PAGE. For guanidine dissociation, pellet was resuspended in 8 M guanidine HCl, incubated at 25°C for 70 min, then sodium deoxycholate was added to a final concentration of 1%. Protein was then precipitated with TCA, washed with acetone at –20°C, and resuspended in sample buffer for SDS-PAGE.

Affinity chromatography

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His-tagged Aac2p affinity purification was performed on 1 mg mitochondria per replicate. Yeast mitochondria were isolated as previously described (Diekert et al., 2001). Mitochondria were lysed in ‘lysis buffer’ (10% glycerol, 1.5% digitonin, 50 mM potassium acetate, 2 mM phenylmethylsulfonyl fluoride [PMSF], and a protease inhibitor cocktail) including 100 mM NaCl (‘low salt’) or 200 mM NaCl (‘high salt’) and incubated on ice for 30 min. Lysate was centrifuged at 21,000×g for 30 min. Imidazole was added to supernatant for a final concentration of 4 mM, which was applied to preequilibrated Ni-NTA agarose beads (QIAGEN) and rotated at 4°C for 2 hr. Beads were then washed three times with ‘lysis buffer’ (including the corresponding NaCl) but with 0.1% digitonin. Protein was eluted from the beads with 300 mM imidazole, 2% SDS, 10% glycerol in 20 mM HEPES-KOH (pH 7.4). For mass spectrometry, samples were briefly run into an SDS-PAGE gel. Whole lanes were excised, and protein was subject to in-gel trypsin digestion. To ensure reproducibility, the eluate used for mass spectrometry was derived from two independent mitochondrial preparations per strain, and affinity chromatography was performed on 2 separate days.

HA-tagged ANT1 affinity purification was performed on whole-cell lysate. Each replicate was from an independent transfection of a 10 cm dish of HeLa cells. Briefly, 24 hr after transfection, cells were collected, washed twice in cold PBS, and then lysed in 0.5 mL lysis buffer (50 mM HEPES-KOH pH 7.4, 10% glycerol, 100 mM NaCl, 1% digitonin, 1 mM PMSF, and 1× HALT Protease and Phosphatase Inhibitor Cocktail (Thermo Fisher)) for 30 min on ice. Lysate was cleared for 30 min at 16,000×g, and supernatant applied to preequilibrated Pierce anti-HA agarose beads (Thermo Fisher) and incubated overnight with gentle agitation. Beads were subsequently washed five times with lysis buffer, except containing 0.1% digitonin, followed by elution with 6% SDS and 10% glycerol, in 50 mM HEPES-KOH pH 7.4.

Sample processing for mass spectrometry

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The excised lanes of yeast eluate (see above) were subjected to in-gel trypsin digestion (Shevchenko et al., 2006). Briefly, gel pieces were washed with 50 mM ammonium bicarbonate (Acros) in 50% acetonitrile (ACN) (Fisher), reduced with dithiothreitol (Acros) and alkylated with iodoacetamide (Sigma), washed again, and impregnated with 75 µL of 5 ng/µL trypsin (trypsin gold; Promega) solution overnight at 37°C. The resulting peptides were extracted using solutions of 50% and 80% ACN with 0.5% formic acid (Millipore), and the recovered solution dried down in a vacuum concentrator. Dried peptides were dissolved in 60 µL of 0.1% trifluoroacetic acid (TFA, Sigma), and desalted using 2-core MCX stage tips (3 M) (Rappsilber et al., 2003). The stage tips were activated with ACN followed by 3% ACN with 0.1% TFA. Next, samples were applied, followed by two washes with 3% ACN with 0.1% TFA, and one wash with 65% ACN with 0.1% TFA. Peptides were eluted with 75 µL of 65% ACN with 5% NH4OH (Sigma), and dried.

Cytosolic fractions from HeLa cells were processed using the FASP method (Wiśniewski et al., 2009). Briefly, in-solution proteins were reduced and denatured with DTT and SDS, mixed with urea to 8 M, and concentrated on a 10 kDa MWCO membrane filter (Pall, OD010C34). Cysteine residues were alkylated using iodoacetamide (Sigma) at room temperature in a dark location for 25 min. The proteins were rinsed with urea and ammonium bicarbonate solutions and digested overnight at 37°C using trypsin gold (Promega) at a ratio of 1:100. The resulting peptides were recovered from the filtrate and a 10 µg aliquot was desalted on 2-core MCX stage tips as above.

LC-MS methods

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Samples were dissolved in 20–35 µL of water containing 2% ACN and 0.5% formic acid to 0.25 µg/µL. Two µL (0.5 µg) were injected onto a pulled tip nano-LC column with 75 µm inner diameter packed to 25 cm with 3 µm, 120 Å, C18AQ particles (Dr. Maisch). The column was maintained at 45°C with a column oven (Sonation GMBH). The peptides were separated using a 60 min gradient from 3–28% ACN over 60 min, followed by a 7 min ramp to 85% ACN. The column was connected inline with an Orbitrap Lumos via a nanoelectrospray source operating at 2.2 kV. The mass spectrometer was operated in data-dependent top speed mode with a cycle time of 2.5 s. MS1 scans were collected at 60,000 resolution with AGC target of 6.0E5 and maximum injection time of 50 ms. HCD fragmentation was used followed by MS2 scans in the Orbitrap at 15,000 resolution with AGC target 1.0E4 and 100 ms maximum injection time.

Database searching and label-free quantification

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The MS data was searched using SequestHT in Proteome Discoverer (version 2.4, Thermo Scientific) against the Saccharomyces cerevisiae proteome from Uniprot, containing 6637 sequences, concatenated with common laboratory contaminant proteins. Enzyme specificity for trypsin was set to semi-tryptic with up to two missed cleavages. Precursor and product ion mass tolerances were 10 ppm and 0.6 Da, respectively. Cysteine carbamidomethylation was set as a fixed modification. Methionine oxidation was set as a variable modification. The output was filtered using the Percolator algorithm with strict FDR set to 0.01. Label-free quantification was performed in Proteome Discoverer with normalization set to total peptide amount.

Label-free quantitation data processing

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For analysis, label-free quantitation with Proteome Discoverer software generated protein abundances for each sample. Protein abundances were then analyzed using Metaboanalyst software (Pang et al., 2020). First, we refined the protein list to include only proteins whose fold-change is >1.5 (p<0.1) in Aac2p-HIS6 eluate compared with the null control. Then, to eliminate any bias introduced by mutant bait protein Aac2pA128P-HIS6 being present at ~50% the level of Aac2-HIS6 (see Figure 4—figure supplement 1C; Figure 4—figure supplement 2A), we normalized each prey protein abundance by that sample’s bait protein level (i.e. Aac2p level). Finally, we performed multiple t testing on these values, which generated the FDR-corrected p-values and were ultimately normalized to wild-type level for presentation in Figure 4C, Figure 4—figure supplement 1I, Figure 4—figure supplement 2B and E. Gene Ontology (GO) and other enrichment analyses were performed using STRING version 11.0 (Szklarczyk et al., 2019).

For the analysis shown in Figure 5F, the protein list was manually curated to include only proteins that had a GO Cellular Component term for ‘mitochondrion’, and not that of any other organelle. Proteins with missing values were excluded. The levels of each protein were averaged within an experimental group and those average values were normalized to the average value for wild-type SLC25A4-transfected samples. It is these wild-type-normalized average values for each protein that were subject to Student’s t test to probe for a significant difference in mitochondrial protein levels present in the cytosol of mutant compared with wild-type SLC25A4-transfected HeLa cells.

Targeted quantitative proteomics

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Unused yeast peptides from the label-free quantification experiment were used for absolute quantification using parallel reaction monitoring on the instrument described above. The instrument method consisted of one MS scan at 60,000 resolution followed by eight targeted Orbitrap MS2 scans at 30,000 resolution using quadrupole isolation at 1.6 m/z and HCD at 35%.

Two heavy-labeled proteotypic peptides for Aac2p and Tim22p were purchased from New England Peptide. Their sequences were TATQEGVISFWR and SDGVAGLYR; and VYTGFGLEQISPAQK and TVQQISDLPFR, respectively, each with N-terminal 13C6 15N4 arginine or 13C6 15N2 lysine. An equal portion of the peptides contained in each gel band was combined with a mixture of proteotypic peptides resulting in an on-column load of 1 fmol of each Tim22p heavy peptide and 250 fmol of each Aac2p heavy peptide. Assays were developed for the doubly charged precursor and the four or five most intense singly charged product y-ions. The data was analyzed in Skyline (version 20.2) (MacLean et al., 2010).

qRT-PCR

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Total RNA was isolated from ~1.5×107 yeast cells using Quick-RNA Fungal/Bacterial Microprep Kit (Zymo Research). Quantitative real-time PCR (qRT-PCR) was executed using 50 ng RNA in Power SYBR Green RNA-to-Ct 1-step Kit (Applied Biosystems). A CFX384 Touch Real-Time PCR Detection System (Bio-Rad) was used with the following cycling parameters: 48°C 30 min, 95°C 10 min, followed by 44 cycles of 95°C 15 s, 60°C 1 min. TFC1 was used as reference, as its mRNA level was previously established as stable across culture conditions (Teste et al., 2009). The specificity of each primer pair was validated using RNA extracted from knockout strains. Ct values were determined using CFX Maestro software (Bio-Rad) and analyzed manually using the 2−ΔΔCT method.

In vitro mitochondrial protein import assays

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The AAC2 variants were cloned into pGEM4z plasmids. The constructs were used for coupled in vitro transcription and translation, using a cell-free system based on reticulocyte lysate. The wild-type strains YPH499 and BY4741 and yeast expressing Tom40-HA were grown in YPG (1% [wt/vol] yeast extract, 2% [wt/vol] bacto-peptone, and 3% [vol/vol] glycerol) at 30°C. Mitochondria were isolated by differential centrifugation. The import of the Aac2p variants were performed as described (Ellenrieder et al., 2019). Isolated mitochondria were incubated for different time periods with radiolabeled Aac2p variants in the presence of 4 mM ATP and 4 mM NADH in import buffer (3% [wt/vol] bovine serum albumin; 250 mM sucrose; 80 mM KCl; 5 mM MgCl2; 5 mM methionine; 2 mM KH2PO4; 10 mM MOPS/KOH, pH 7.2). The import reaction was stopped by addition of 8 µM antimycin A, 1 µM valinomycin, and 20 µM oligomycin (AVO; final concentrations). In control reactions, the membrane potential was depleted by addition of the AVO mix. Subsequently, mitochondria were washed with SEM buffer (250 mM sucrose; 1 mM EDTA; 10 mM MOPS-KOH, pH 7.2), lysed with 1% (wt/vol) digitonin in lysis buffer (0.1 mM EDTA; 50 mM NaCl; 10% [vol/vol] glycerol; 20 mM Tris-HCl, pH 7.4) for 15 min on ice analysis and protein complexes were separated on blue native gels. To remove non-imported preproteins, mitochondria were treated with 50 µg/mL proteinase K for 15 min on ice. The protease was inactivated by addition of 1 mM PMSF for 10 min on ice. To study the accumulation of Aac2p variants at the TOM translocase, the Aac2p variants were imported into Tom40-HA mitochondria followed by affinity purification via anti-HA beads (Roche) (Ellenrieder et al., 2019). After the import reaction, mitochondria were lysed with 1% (wt/vol) digitonin in lysis buffer for 15 min on ice. After removing insoluble material, the mitochondrial lysate was incubated with anti-HA matrix for 60 min at 4°C. Subsequently, beads were washed with an excess amount of 0.1% (wt/vol) digitonin in lysis buffer and bound proteins were eluted under denaturing conditions using SDS sample buffer (2% [wt/vol] SDS; 10% [vol/vol] glycerol; 0.01% [wt/vol] bromophenol blue; 0.2% [vol/vol] β-mercaptoethanol; 60 mM Tris/HCl, pH 6.8).

Pulse-chase analysis of Aac2pA128P, A137D

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Cells with disrupted endogenous AAC2 but expressing GAL10-aac2A128P, A137D, with or without the disruption of YME1 and PDR5, were grown in YPD at 30°C overnight. Cells were then subcultured in complete galactose plus raffinose medium for 4 hr at 30°C. Cycloheximide was added to a concentration of 1 mg/mL, with or without the addition of 100 μM MG132. Cycloheximide chase was pursued at 30°C for 30, 60, and 120 min before cells were collected for western blot analysis.

Human cell culture

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HeLa cells were cultured in DMEM (Gibco) with 10% fetal bovine serum (Sigma) at 37°C in a humidified atmosphere of 5% CO2. The HeLa cell line was purchased from ATCC (CCL-2). Cell line was authenticated by ATCC with short tandom repeat analysis. Cells tested free of mycoplasma by the VenorGem Mycoplasma Detection Kit (Sigma-Aldrich #MP0025).

Expression of SLC25A4 in HeLa cells

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SLC25A4 cDNA was cloned into pCDNA3.1 with an HA epitope added to the C-terminus, as previously described (Liu et al., 2019). Mutant SLC25A4 alleles were generated by in vitro mutagenesis using QuikChange Site-Directed Mutagenesis (Stratagene) and confirmed by sequencing. Cells were transfected using Lipofectamine 3000 (Invitrogen) according to the manufacturer’s protocol and harvested 24 hr after transfection for all experiments.

HeLa cell fractionation

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One 10 cm dish per sample was harvested, washed twice in cold PBS, homogenized in 1 mL isotonic buffer (250 mM sucrose, 1 mM EDTA, 10 mM Tris-HCl, pH 7.4,) including HALT Protease and Phosphatase Inhibitor Cocktail (Thermo Fisher) with 10 slow strokes in a 2 mL Dounce homogenizer (KIMBLE, pestle B, clearance = 0.0005–0.0025 inches). Homogenate was centrifuged at 600×g for 15 min. To pellet mitochondria, supernatant was spun at 10,000×g for 25 min. Supernatant was the cytosolic fraction and pellet (the mitochondrial fraction) was used subsequently for protease protection or western blotting. All steps were performed on ice.

Protease protection assay in HeLa cells

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For protease protection, isolated mitochondria from four 10 cm plates were combined to generate each replicate indicated in Figure 5E. All replicates were from independent transfections on different days. Twenty μg aliquots of mitochondria were pelleted and resuspended in either isotonic buffer (as above, without protease inhibitors) or hypotonic buffer (10 mM KCl, 2 mM HEPES, pH 7.2) for swelling on ice for 20 min. Where indicated, proteinase K was added at a final concentration of 7 μg/mL, incubated at room temperature for 20 min, and inhibited with 5 mM PMSF on ice for 15 min. Where indicated, 1% Triton X-100 treatment on ice for 10 min was used to lyse the mitochondrial membranes. Ultimately, half of each reaction was loaded onto two independent gels for western blotting. Immunoblotting of reference proteins in Figure 5D was done for each replicate. If a reference protein was not properly protected/degraded, this sample was discarded.

Apoptosis assay and relative Δψ determination

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Cells were harvested 24 hr post-transfection and processed for flow cytometry detection of apoptosis and membrane potential after staining with propidium iodide and Annexin V-FITC and JC-1, respectively (Liu et al., 2019).

Slc25a4 p.A114P,A123D/+ knock-in mice

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All procedures were approved by the Animal Care and Use Committee (IACUC) at State University of New York Upstate Medical University and were in accordance with guidelines established by the National Institutes of Health.

The Slc25a4 targeting vector was prepared by recombineering as previously described (Lee et al., 2001). Briefly, 13.8 kb of Slc25a4 genomic sequence containing all four exons plus 4.4 kb of 5’ upstream and 4.96 kb 3’ downstream untranslated sequences was retrieved from the RP24-108A1 BAC clone obtained from the BACPAC Resources Center (Children’s Hospital Oakland Research Institute, Oakland, CA, USA). The first loxP site together with the Frt-PGKnew-Frt cassette was inserted approximately 0.4 kb 3’ of exon 4, which contains the polyA signal sequence and the 3’UTR. Two unique restriction sites, AscI and AsiSI, were also introduced into the 3’ end of the second loxP site to allow insertion of the Slc25a4 mini-cDNA containing the A114P and A123D knock-in mutations as well as the His-tag at the carboxyl terminus.

The Slc25a4 mini-cDNA was prepared by fusion PCR using primers with sequences overlapping different exons. The mini-cDNA contains AscI and AsiSI unique restriction sites in the 5’ and 3’ end, respectively, together with 254 bp of intron 1 sequence together with the splice acceptor followed by exon 2 with the two mutations, exons 2 and 4, and 70 bp of 3’ downstream sequence. The mini-cDNA was cloned into pSK+ and sequenced to confirm its identity prior to insertion into the AscI and AsiSI sites in the targeting vector. The final targeting vector was then linearized by NotI digestion, purified, and resuspended in PBS at 1 mg/µL for electroporation into ES cells derived from F1 (129Sv/C57BL6j) blastocyst. Targeted ES clones were identified by long-range nested PCR using Platinum HiFi Taq (Invitrogen).

Chimeric animals were generated by aggregation of ES cells with the CD1 morula. Chimeric males were bred with ROSA26-Flpe female (Jackson Labs stock no: 009086) to remove the PGKnew cassette and generate F1 pups with Slc25a4 floxed allele. Positive pups were identified by PCR genotyping using primer Lox gtF (5’-ATCCATCTCAAAGGCAAACG-3’) and Lox gtR (5’- AAATTCCCTGCAGGCTTATG-3’) to detect a fragment of 364 bp specific to the 5’-Lox site and a fragment of 270 bp specific to the wild-type allele. A heterozygous floxed male was bred with Hprt-Cre female (Jackson Labs stock no: 004032). The same primers were used for genotyping knock-in mice, with the 270 bp band present only with the knock-in allele, and no band produced from a wild-type locus with lacking loxP sites. The mixed background heterozygous knock-in (i.e. Slc25a4 p. A114P,A123D/+) male mice were backcrossed with C57BL/6NTac females (Taconic Catalog no: B6-F) for seven generations before experiments were performed.

Mouse histology

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For spinal cord histology, mice were sacrificed by isoflurane overdose, followed by intracardial perfusion with PBS followed by PBS+4% paraformaldehyde (PFA). Perfused mice were soaked in PBS+4% PFA overnight at 4°C, and then spinal cord was dissected and cryopreserved with increasing concentrations of sucrose in PBS. Tissue was then embedded in OCT and snap-frozen in 2-methylbutane on liquid N2. Tissue was sectioned at 8 μm with a cryostat (Leica) and sections stained with Cresyl Violet Stain Solution according to the manufacturer’s protocol (Abcam) or used for indirect immunofluorescence. For the latter, tissue was blocked with 10% horse serum and then incubated with rat monoclonal anti-GFAP antibody (Invitrogen) overnight at 4°C. After rinsing the tissue was incubated with fluorescently conjugated anti-rat secondary antibody (ImmunoResearch, West Grove, PA, USA).

For muscle histology, the soleus muscles were quickly dissected and fresh-frozen in 2-methylbutane on liquid N2. Tissue was cryosectioned at 10 μm and stained with hematoxylin and eosin using standard procedures. Feret’s diameter of myofibers was determined using ImageJ software; all soleus myofibers from a single muscle section per mouse (n>340 fibers per mouse) were quantitated by a blinded observer. For SDH-only staining, sections were air-dried, incubated at 37°C for 45 min in SDH medium (0.1 M succinic acid, 0.1 M sodium phosphate buffer pH 7, 0.2 mM phenazine methosulfate, +1 mg/mL NBT added fresh), drained, fixed in 10% formalin, rinsed well with water, and mounted in water soluble mounting medium. For sequential COX/SDH staining, sections were air-dried, incubated in cytochrome c medium (0.5 mg/mL 3’3’-diaminobenzidine tetrahydrochloride, 75 mg/mL sucrose in 50 mM sodium phosphate buffer, pH 7.4, with freshly added cytochrome c and catalase at 1 and 0.1 mg/mL, respectively) at 37°C for 1 hr, followed by a quick wash in water and SDH staining as described above. Abnormal COX/SDH-stained fibers were scored manually from decoded images of whole soleus sections.

Electron microscopy

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For electron microscopy, Slc25a4 p.A114P,A123D/+ mice and littermate controls were processed as previously described (Massa et al., 2004). Briefly, mice were anesthetized with isoflurane and perfused intracardially with PBS initially, followed by fixative (1% PFA, 1% glutaraldehyde, 0.12 M sodium cacodylate buffer pH 7.1, and 1 mM CaCl2). Perfused animals were refrigerated overnight, and CNS tissues were dissected the next day and processed for TEM. The samples were examined with a JOEL JEM1400 transmission electron microscope and images were acquired with a Gaten DAT-832 Orius camera.

Bioenergetic analysis

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For mitochondrial respiration experiments, the mice were sacrificed by decapitation via guillotine without the use of CO2 asphyxiation or anesthetic. Skeletal muscle mitochondria were isolated and respiration measured as previously described (Garcia-Cazarin et al., 2011). Briefly, after mitochondria isolation by differential centrifugation, oxygen tension was measured using an Oxygraph Plus oxygen electrode (Hansatech Instruments) in 0.5 mL experimental buffer containing 150 μg mitochondria in a temperature-controlled 37°C chamber. For complex I measurements, glutamate and malate were added for a final concentration of 5 and 2.5 mM, respectively. For complex II measurements, rotenone was added before succinate, for final concentrations of 5 μM and 10 mM, respectively.

Protease protection assay of mouse skeletal muscle mitochondria

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For protease protection assay, skeletal muscle mitochondria were isolated as for bioenergetic analysis with slight modifications, including the addition of 1 mM PMSF in the homogenization buffer, and resuspending mitochondria in a modified isotonic buffer lacking BSA (75 mM sucrose, 215 mM mannitol, 1 mM EGTA in 20 mM HEPES-KOH pH 7.4). Eighty μg aliquots were treated with the indicated concentrations of proteinase K (Sigma) for 30 min at room temperature and quenched with 5 mM PMSF on ice for 10 min. Where indicated, mitochondria were lysed with 1% Triton X-100 for 30 min on ice. Laemmli buffer was ultimately added to the samples such that the final protein concentration was ~40 μg mitochondria per 15 μL, which was the amount loaded onto the SDS-PAGE gel for western blotting. Such high protein amounts were required for detection of Ant1A114P,A123D.

BN-PAGE

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Snap-frozen mouse skeletal muscle mitochondria isolated for bioenergetic analysis were used for BN-PAGE. Protein complexes were solubilized with digitonin at a 4:1 detergent:protein ratio and loaded into native PAGE gel (Invitrogen) per the manufacturer’s instructions. For western blotting, gels were washed in 2% SDS followed by standard electrophoretic transfer to a PVDF membrane. Following transfer, membrane was fixed with acetic acid and dried followed by standard western blotting techniques.

Mouse skeletal muscle sub-cellular fractionation

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Five biological replicates of wild-type and Slc25a4 p.A114P,A123D/+ muscle samples were fractionated to obtain cytosolic and mitochondrial fractions using differential centrifugation, as follows. One-hundred mg Quadriceps muscle was rapidly dissected and immediately placed in ice-cold PBS. Muscle was minced, centrifuged for 1 min at 500×g, and resuspended in 1 mL buffer STM (250 mM sucrose, 50 mM Tris-HCl pH 7.4, 5 mM MgCl2 plus HALT protease and phosphatase inhibitors added fresh). Tissue was homogenized in a 2 mL Dounce homogenizer (0.15–0.25 mm clearance) with two strokes using a bench top drill press set to 570 rotations per minute. Homogenate was centrifuged for 15 min at 800×g twice, and the pellets were discarded after both spins. To obtain the mitochondrial fraction, supernatant was centrifuged for 11,000×g for 10 min. Mitochondrial fraction was washed twice in STM and frozen for further analysis. Meanwhile, the supernatant was centrifuged for 30 min at 21,000×g twice to remove any contaminating mitochondria from the cytosolic fraction, although this may not completely eliminate mitochondrial proteins trapped in small mitochondria-derived vesicles.

Mouse skeletal muscle cytosolic and mitochondrial protein digestion, labeling, cleanup, and fractionation

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Skeletal muscle cytosolic and mitochondrial fractions were then processed separately for multiplexed quantitative mass spectrometry as follows. Samples were buffer exchanged on a 3 kDa molecular weight cutoff filter (Amicon 3k Ultracel) using four additions of 50 mM triethylammonium bicarbonate, pH 8.0 (Thermo). Following a Bradford assay, 50 µg of each cytosolic fraction was taken for digestion using an EasyPep Mini MS sample prep kit (Thermo, A40006). To each buffer-exchanged sample, 70 µL of lysis buffer was added followed by 50 µL of reduction solution and 50 µL of alkylating solution. Samples were incubated at 95°C for 10 min, then cooled to room temperature. To each sample 2.5 µg of trypsin/Lys-C protease was added and the reaction was incubated at 37°C overnight. TMT reagents were reconstituted with 40 µL ACN and the contents of each label added to a digested sample. After 60 min, 50 µL of quenching solution was added, consisting of 20% formic acid and 5% ammonium hydroxide (vol/vol) in water. The labeled digests were cleaned up by a solid-phase extraction contained in the EasyPep kit, and dried by speed-vac. The 10 cytosolic fractions were dissolved in 50 µL of 30% ACN and 0.1% formic acid (v/v) in water, combined, and dried again.

Following an LC-MS experiment to check digestion and labeling quality of the pooled samples, these were fractionated using a Pierce High pH Reversed-Phase Peptide Fractionation Kit (part # 84868), per the manufacturer’s instructions for TMT-labeled peptides. In brief, samples were dissolved in 300 µL of 0.1% TFA in water and applied to the conditioned resin. Samples were washed first with water and then with 300 µL of 5% ACN, 0.1% triethylamine (TEA) in water. The second wash was collected for analysis. Peptides were step eluted from the resin using 300 µL of solvent consisting of 5–50% ACN with 0.1% TEA in eight steps. All collected fractions were dried in a speed-vac.

LC-MS/MS for TMT-labeled samples

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Dried fractions were reconstituted in 50 µL of load solvent consisting of 3% ACN and 0.5% formic acid in water. Of these, 2 µL were injected onto a pulled tip nano-LC column (New Objective, FS360-75-10-N) with 75 µm inner diameter packed to 25 cm with 2.2 µm, 120 Å, C18AQ particles (Dr. Maisch GmbH). The column was maintained at 50°C with a column oven (Sonation GmbH, PRSO-V2). The peptides were separated using a 135 min gradient consisting of 3–12.5% ACN over 60 min, 12.5–28% over 60 min, 28–85% ACN over 7 min, a 3 min hold, and 5 min re-equilibration at 3% ACN. The column was connected inline with an Orbitrap Lumos (Thermo) via a nanoelectrospray source operating at 2.3 kV. The mass spectrometer was operated in data-dependent top speed mode with a cycle time of 3 s. MS1 scans were collected from 375 to 1500 m/z at 120,000 resolution and a maximum injection time of 50 ms. HCD fragmentation at 40% collision energy was used followed by MS2 scans in the Orbitrap at 50,000 resolution with a 105 ms maximum injection time.

Database searching and reporter ion quantification

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The MS data was searched using SequestHT in Proteome Discoverer (version 2.4, Thermo Scientific) against the Mus musculus proteome from Uniprot, containing 50,887 sequences, concatenated with common laboratory contaminant proteins. Enzyme specificity for trypsin was set to semi-tryptic with up to two missed cleavages. Precursor and product ion mass tolerances were 10 ppm and 0.6 Da, respectively. Cysteine carbamidomethylation, TMT 10-plex at any N-terminus, and TMT 10-plex at lysine were set as a fixed modifications. Methionine oxidation was set as a variable modification. The output was filtered using the Percolator algorithm with strict FDR set to 0.01. Quantification parameters included the allowance of unique and razor peptides, reporter abundance based on intensity, lot-specific isotopic purity correction factors, normalization based on total peptide amount, protein ratio based on protein abundance, and hypothesis testing (ANOVA) for individual proteins.

Grip strength measurements

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Grip strength measurements were performed using Bio-CIS software connected with the Grip Strength Test Model GT3 according to the manufacturer’s protocol for forelimb-only measurement using the grid (BIOSEB). Briefly, mice were held by the tail above the grid that’s connected to a force meter, slowly lowered to allow the forelimbs to grip the grid, and then slowly and smoothly pulled horizontally along the axis of the sensor until the grasp was released. The maximum force generated is recorded by the software and reported as the average of five consecutive trials or the maximum force generated over five trials.

RNA extraction, sequencing, and analysis

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Skeletal muscle was disputed in QIAzol lysis reagent, using the QIAGEN TissueRuptor. RNA was then extracted using the Qiagen miRNeasy Mini Kit. RNA quality and quantity were assessed with the RNA 6000 Nano kit on the Agilent 2100 Bioanalyzer. Sequencing libraries were prepared using the Illumina TruSeq Stranded mRNA Library Prep kit, using 1 µg total RNA as input. Library size was assessed with the DNA 1000 Kit on the Agilent 2100 Bioanalyzer. Libraries were quantified using the Quant-IT High Sensitivity dsDNA Assay (Invitrogen) on a Qubit 3.0 Fluorometer. Libraries were sequenced on the NextSeq 500 instrument, with paired end 2×75 bp reads.

For proper comparison with Ant1 knockout mice, thresholds for significance and fold-change were replicated exactly as previously published (Morrow et al., 2017): a significance threshold of p<0.01, >1.5-fold increase for upregulated genes, >0.3-fold decrease for downregulated genes, and enrichment analysis was done using the Database for Annotation, Visualization and Integrity Discovery (DAVID) software (Dennis et al., 2003). Databases analyzed were also kept consistent with (Dennis et al., 2003), and included GO, Sequence Features (Seq), InterPro, Kyoto Encyclopedia of Genes and Genomes (KEGG), and Protein Information Resource (PIR). We were unable to perform a direct comparison of significantly changed genes in Slc25a4 p.A114P,A123D/+ vs Ant1 knockout mice because the data are not publicly available.

Statistical analysis

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Statistical analyses were performed using GraphPad Prism. For details on statistical testing of specific data, please see figure legends.

Appendix 1

Appendix 1—key resources table
Reagent type (species) or resourceDesignationSource or referenceIdentifiersAdditional information
Cell line (Homo sapiens)HeLaATCCCCL-2
AntibodyPolyclonal rabbit ant-Aac2Chen Lab1:3000
AntibodyPolyclonal rabbit anti-Ilv5Chen Lab1:5000
AntibodyPolyclonal rabbit anti-Hsp60Chen Lab1:10,000
AntibodyPolyclonal rabbit anti-TFAMSigmaCat. #SAB1401383-100UG1:1000
AntibodyMonoclonal mouse anti-hemagglutinin (HA)CovanceCat. #MMS-101R1:2000
AntibodyPolyclonal rabbit anti-Tim22 (yeast)Nikoalus Pfanner Lab51131:10,000
AntibodyPolyclonal rabbit anti-Tim23 (yeast)Ron Butow Lab1:5000
AntibodyMonoclonal rabbit anti-Tom20 (human)Cell SignalingCat. #424061:2000
AntibodyMonoclonal rabbit anti-Tom40 (human)AbcamCat. #ab1855431:1000
AntibodyPolyclonal rabbit anti-Tim22 (human)Protein TechCat. #14927–1-AP1:2000
AntibodyMonoclonal mouse anti-Tim23 (human)BD BiosciencesCat. #6112221:5000
AntibodyPolyclonal rabbit anti-Smac (human)AbcamCat. #ab81141:2000
AntibodyPolyclonal rabbit anti-Ant1SigmaCat. #SAB2108761-100UL1:2000
AntibodyMonoclonal rabbit anti-Mdh2 (D8Q5S)Cell SignalingCat. #119081:2000
AntibodyMonoclonal rat anti-GFAP (immunostaining)InvitrogenCat. #13-030010 μg/mL
AntibodyMonoclonal mouse anti-GFAP (western blot)Chemicon InternationalCat. #MAB3601:1000
AntibodyMonoclonal mouse anti-GAPDHAbcamCat. #ab94821:2000
AntibodyMonoclonal mouse anti-Pgk1InvitrogenCat. #4592501:4000
AntibodyPolyclonal rabbit anti-Sml1Rothstein lab1:5000
AntibodyPolyclonal rabbit ant-eIF2αCell SignalingCat. #97221:2000
AntibodyMonoclonal rabbit ant-Phospho-eIF2αCell SignalingCat. #35971:1000
AntibodyMonoclonal mouse anti-NDUFA9AbcamCat. #ab147131:1000
AntibodyMonoclonal mouse MitoProfile Total OXPHOS Human WB Antibody CocktailAbcamCat. #ab1104111:500
Chemical compound, drugL-[35S]-methioninePerkin ElmerCat. #NEG009005MC
Commercial assay or kitAnti-HA affinity matrix (used for yeast)RocheCat. #11815016001
Commercial assay or kitAnti-HA beads (used for HeLa cells)Thermo ScientificCat. #26181
Commercial assay or kitNi-NTA agarose beadsQIAGENCat. #1018244
Commercial assay or kitHALT Protease and Phosphatase Inhibitor CocktailThermo ScientificCat. #1861284
Commercial assay or kitJC-1Life TechnologiesCat. #T3168
Peptide, recombinant proteinAac2 Peptide 1New England PeptideTATQEGVISFWR
Peptide, recombinant proteinAac2 Peptide 2New England PeptideSDGVAGLYR
Peptide, recombinant proteinTim22 Peptide 1New England PeptideVYTGFGLEQISPAQK
Peptide, recombinant proteinTim22 Peptide 2New England PeptideTVQQISDLPFR
Commercial assay or kitTNT Quick Coupled Reaction MixPromegaCat. #L2080
Commercial assay or kitFITC Annexin V Apoptosis Detection Kit with PIBioLegendCat. #640914
Commercial assay or kitQuick-RNA Fungal/Bacterial Microprep KitZymoCat. #R2010
Commercial assay or kitPower SYBR Green RNA-to-Ct 1-step KitThermo Fisher ScientificCat. #4389986
Commercial assay or kitLipofectamine 3000InvitrogenCat. #L3000-015
Commercial assay or kitQuikChange Site-Directed MutagenesisStratageneCat. #200518
Commercial assay or kitRevert 700 Total Protein StainLI-CORCat. #926–11021
Biological sample (Saccharomyces cerevisiae)W303-1BR. RothsteinMATa, ade2, trp1, his3, leu2, ura3
Biological sample (Saccharomyces cerevisiae)CS1382-4AThis studyas W303-1B, but trp1Δ::aac2A128P-URA3
Biological sample (Saccharomyces cerevisiae)CS1458/1This studyas W303-1B, but trp1Δ::aac2A137D-URA3
Biological sample (Saccharomyces cerevisiae)CS1763-5AThis studyas W303-1B, but lys2Δ::aac2A128P, A137D-kan
Biological sample (Saccharomyces cerevisiae)CS341/1Chen labas W303-1B, but aac2Δ::kan
Biological sample (Saccharomyces cerevisiae)CY4193This studyasW303-1B, but aac2Δ::LEU2, trp1Δ::aac2A128P-URA3
Biological sample (Saccharomyces cerevisiae)CS1762/2-8AThis studyas W303-1B, but aac2Δ:kan, lys2Δ::aac2A137D-kan
Biological sample (Saccharomyces cerevisiae)CS1763-7DThis studyas W303-1B, but aac2Δ:kan, lys2Δ::aac2A128P,A137D-kan
Biological sample (Saccharomyces cerevisiae)CY6518This studyas W303-1B, but ura3::pUC-URA-GAL10-AAC2
Biological sample (Saccharomyces cerevisiae)CY6519This studyas W303-1B, but ura3::pUC-URA-GAL10-aac2A128P
Biological sample (Saccharomyces cerevisiae)CY6520This studyas W303-1B, but ura3::pUC-URA-GAL10- aac2A137D
Biological sample (Saccharomyces cerevisiae)CY6521This studyas W303-1B, but ura3::pUC-URA-GAL10- aac2A128P, A137D
Biological sample (Saccharomyces cerevisiae)CY6513This studyas W303-1B, but aac2Δ::Kan, lysΔ::aac2A128P, A137D-kan, pdr5Δ::Kan
Biological sample (Saccharomyces cerevisiae)CY6503This studyas W303-1B, but aac2Δ::kan, lys2Δ::aac2A128P, A137D-kan.
Biological sample (Saccharomyces cerevisiae)CY6510This studyas W303-1B, but lys2Δ::aac2A128P, A137D-kan, ump1Δ::kan
Biological sample (Saccharomyces cerevisiae)CY6511This studyas W303-1B, but aac2Δ::kan, lys2Δ::aac2A128P, A137D-kan, pre9Δ::kan
Biological sample (Saccharomyces cerevisiae)CY6540This studyas W303-1B, but aac2Δ::kan, ura3::pUC-URA-GAL10-AAC2
Biological sample (Saccharomyces cerevisiae)CY6542This studyas W303-1B, but aac2Δ::kan, ura3::pUC-URA-GAL10-aac2A128P
Biological sample (Saccharomyces cerevisiae)CY6544This studyas W303-1B, but aac2Δ::kan, ura3::pUC-URA-GAL10-aac2A137D
Biological sample (Saccharomyces cerevisiae)CY6546This studyas W303-1B, but aac2Δ::kan, ura3::pUC-URA-GAL10-aac2A128P, A137D
Biological sample (Saccharomyces cerevisiae)CY6558This studyas W303-1B, but aac2Δ::kan, yme1Δ::Kan, ura3::pUC-URA-GAL10-aac2A128P, A137D
Biological sample (Saccharomyces cerevisiae)CY6562This studyas W303-1B, but aac2Δ::kan, pdr5Δ::Kan, ura3::pUC-URA-GAL10-aac2A128P, A137D
Biological sample (Saccharomyces cerevisiae)CY6569This studyas W303-1B, but aac2Δ::kan, pdr5Δ::Kan, yme1Δ::LEU2, ura3::pUC-URA-GAL10-aac2A128P, A137D
Biological sample (Saccharomyces cerevisiae)CY6440This studyas W303-1B, but pre9Δ::kan
Biological sample (Saccharomyces cerevisiae)CY6504This studyas W303-1B, but ump1Δ::kan
Biological sample (Saccharomyces cerevisiae)CY6581This studyas W303-1B, but ura3::pUC-URA-GAL10-aac2R96H
Biological sample (Saccharomyces cerevisiae)CY6583This studyas W303-1B, but ura3::pUC-URA-GAL10-aac2R252G
Biological sample (Saccharomyces cerevisiae)CY6775This studyMATa/a, ade2/ade2, trp1/trp1, his3/his3, leu2/leu2, ura3/ura3, psd1∆::kan/+, lys2Δ::aac2A128P, A137D-kan/+
Biological sample (Saccharomyces cerevisiae)CY6777This studyMATa/a, ade2/ade2, trp1/trp1, his3/his3, leu2/leu2, ura3/ura3, pel1∆::LEU2/+, lys2Δ::aac2A128P, A137D-kan/+
Biological sample (Saccharomyces cerevisiae)CY3326Chen labMATa, his3Δ1, leu2Δ0, lys2Δ0, ura3Δ0, ura3Δ::aac2A128P-URA3
Biological sample (Saccharomyces cerevisiae)BY4742/AG3Chen labMATa, his3Δ1, leu2Δ0, lys2Δ0, ura3Δ0, trp1 Δ::GAL10-AAC2-HIS3
Biological sample (Saccharomyces cerevisiae)CY3322Chen labMATa, his3Δ1, leu2Δ0, lys2Δ0, ura3Δ0, trp1 Δ::GAL10-AAC2A128P-HIS3
Biological sample (Saccharomyces cerevisiae)BY4741EUROSCARFMATa, his3∆1, leu2∆0, met15∆0, ura3∆0
Biological sample (Saccharomyces cerevisiae)BY4741/ssa4DOpen Biosystemsas BY4741, but ssa4Δ::kan
Biological sample (Saccharomyces cerevisiae)BY4741/rpn4DOpen Biosystemsas BY4741, but rpn4Δ::kan
Biological sample (Saccharomyces cerevisiae)BY4741/hsp82DOpen Biosystemsas BY4741, but hsp82Δ::kan
Biological sample (Saccharomyces cerevisiae)BY4741/ssa3DOpen Biosystemsas BY4741, but ssa3Δ::kan
Biological sample (Saccharomyces cerevisiae)BY4741/cis1DOpen Biosystemsas BY4741, but cis1Δ::kan
Biological sample (Saccharomyces cerevisiae)BY4741/TOM40-HAEllenrieder et al., 2019as BY4741, but tom40::TOM40HA-HIS3MX6
Biological sample (Saccharomyces cerevisiae)M2915-6AChen labMATa, ade2, leu2, ura3
Biological sample (Saccharomyces cerevisiae)CY3323Chen labMATa, ade2, ura3, leu2, ura3Δ::aac2A128P-URA3
biological sample (Saccharomyces cerevisiae)YKSL210This studyas M2916-6A, but aac2Δ::LEU2, lys2Δ::AAC2-HIS6-kan
Biological sample (Saccharomyces cerevisiae)YKSL211This studyas M2916-6A, but aac2Δ::LEU2, lys2Δ::aac2A128P-HIS6-kan
Biological sample (Saccharomyces cerevisiae)CY3904This studyas M2915-6A, but tom70Δ::kan
biological sample (Saccharomyces cerevisiae)CY6316This studyas M2915-6A, but tim18Δ::Kan
Biological sample (Saccharomyces cerevisiae)YPH499Sikorski and Hieter, 1989MATa ura3-52, lys2-801_amber, ade2-101_ochre, trp1-Δ63, his3-Δ200, leu2-Δ1
Biological sample (Mus musculus)C57BL6/NTacTaconicCat. #: B6-F
Biological sample (Mus musculus)Hprt-Cre femaleJackson LabsStock no: 004032
Biological sample (Mus musculus)Ant1A114P,A123D knock-in miceThis studySee Materials and methods
Sequence-based reagentTFC1 fwd (B)Teste et al., 2009PCR primersGCTGGCACTCATATCTTATCGTTTCACAATGG
Sequence-based reagentTFC1 rev (B)Teste et al., 2009PCR primersGAACCTGCTGTCAATACCGCCTGGAG
Sequence-based reagentHSP82 fwdBoos et al., 2019PCR primersGCTGCTTTGGCTAAGTTGTTACGTTAC
Sequence-based reagentHSP82 revBoos et al., 2019PCR primersGAGATTCACCAGTGATGTAGTAGATGTTC
Sequence-based reagentRPN4 fwdBoos et al., 2019PCR primersGCAACAAGAGCAACACCAAGAGGAG
Sequence-based reagentRPN4 revBoos et al., 2019PCR primersCTGTCCATGTTAGAGTCAACGTAACTG
Sequence-based reagentCIS1 fwdBoos et al., 2019PCR primersATCAGTAATTGTCCCATCGGGTTAGTTTC
Sequence-based reagentCIS1 revBoos et al., 2019PCR primersCCTGGGCAGCCTTGAGTAAATCATATC
Sequence-based reagentSSA3 fwd (A)This studyPCR primersGGATAAGAAAGGCAGGGCTGA
Sequence-based reagentSSA3 rev (A)This studyPCR primersCTGCGGTAGCCTTAACCTCAA
Sequence-based reagentSSA4 fwd (B)This studyPCR primersAGGCAAGCAACAAAAGATGCC
Sequence-based reagentSSA4 rev (B)This studyPCR primersTTGTCCAGCCCATACGCAATA
Sequence-based reagentTOM70P1This studyPCR primersGAAAGAGTTTCATTGCCATTAG
Sequence-based reagentTOM70P2This studyPCR primersTTGTGGTTTATACGCACTGC
Sequence-based reagentTOM70P3This studyPCR primersAACACTGTGCAGGCAACTTC
Sequence-based reagentTOM70P4This studyPCR primersCTCCGCAAATTGGCGAGG
Sequence-based reagentTIM18KOFPThis studyPCR primersCCATTCTCGCAAAAGATCGG
Sequence-based reagentTIM18KORPThis studyPCR primersTCTGGATTTCGAGAAGAAGG
Sequence-based reagentTIM18GTFPThis studyPCR primersGTCAGTGCCCTCGAGAGC
Sequence-based reagentTIM18GTRPThis studyPCR primerscccaagcttCGCAGATAGTGCGATAGTTG
Sequence-based reagentLox gtFThis studyPCR primersATCCATCTCAAAGGCAAACG
Sequence-based reagentLox gtRThis studyPCR primersAAATTCCCTGCAGGCTTATG
Recombinant DNA reagentpRS416Chen Lab
Recombinant DNA reagentpRS416-AAC2This studySee Materials and methods
Recombinant DNA reagentpRS416-aac2(A106P)This studySee Materials and methods
Recombinant DNA reagentpRS416-aac2(M114P)This studySee Materials and methods
Recombinant DNA reagentpRS416-aac2(A128P)This studySee Materials and methods
Recombinant DNA reagentpRS416-aac2(A137D)This studySee Materials and methods
Recombinant DNA reagentpRS416-aac2(A106D,M114P)This studySee Materials and methods
Recombinant DNA reagentpRS416-aac2(A106D,A128P)This studySee Materials and methods
Recombinant DNA reagentpRS416-aac2(A106D,A137D)This studySee Materials and methods
Recombinant DNA reagentpRS416-aac2(M114P,A128P)This studySee Materials and methods
Recombinant DNA reagentpRS416-aac2(M114P,A137D)This studySee Materials and methods
Recombinant DNA reagentpRS416-aac2(A128P,A137D)This studySee Materials and methods
Recombinant DNA reagentpCDNA3.1This studySee Materials and methods
Recombinant DNA reagentpCDNA3.1-Ant1This studySee Materials and methods
Recombinant DNA reagentpCDNA3.1-Ant1(A90D)This studySee Materials and methods
Recombinant DNA reagentpCDNA3.1-Ant1(L98P)This studySee Materials and methods
Recombinant DNA reagentpCDNA3.1-Ant1(A114P)This studySee Materials and methods
Recombinant DNA reagentpCDNA3.1-Ant1(A123D)This studySee Materials and methods
Recombinant DNA reagentpCDNA3.1-Ant1(A90D,L98P)This studySee Materials and methods
Recombinant DNA reagentpCDNA3.1-Ant1(A90D,A114P)This studySee Materials and methods
Recombinant DNA reagentpCDNA3.1-Ant1(A90D,A123D)This studySee Materials and methods
Recombinant DNA reagentpCDNA3.1-Ant1(L98P,A114P)This studySee Materials and methods
Recombinant DNA reagentpCDNA3.1-Ant1(L98P,A123D)This studySee Materials and methods
Recombinant DNA reagentpCDNA3.1-Ant1(A114P,A123D)This studySee Materials and methods
Recombinant DNA reagentpCDNA3.1-Ant1(A90D,A114P,A123D)This studySee Materials and methods
Recombinant DNA reagentpCDNA3.1-Ant1(A90D,L98P,A114P,A123D)This studySee Materials and methods
Recombinant DNA reagentpGEM-4Z-AAC2 (Saccharomyces cerevisiae)This studySee Materials and methods
Recombinant DNA reagentpGEM-4Z-aac2(A128P)This studySee Materials and methods
Recombinant DNA reagentpGEM-4Z-aac2(A137D)This studySee Materials and methods
Recombinant DNA reagentpGEM-4Z-aac2(A128P,A137D)This studySee Materials and methods
Software, algorithmImageJNIH
Software, algorithmMulti Gauge v.3.2FujiFilm
Software, algorithmImage StudioLI-COR
Software, algorithmPrism version 9GraphPad, LLC
Software, algorithmProteome Discoverer version 2.4Fisher
Software, algorithmMetaboanalystPang et al., 2020
Software, algorithmSTRING version 11.0Szklarczyk et al., 2019
Software, algorithmSkyline version 20.2MacCoss Lab Software
Software, algorithmCFX Maestro softwareBio-Rad
Software, algorithmBioCISBIOSEB
OtherGrip Strength Test Model GT3BIOSEBForce meter that measures mouse grip strength.
OtherOxygraph Plus System Version 2.1Hansatech InstrumentsApparatus with Clark-type electrode to measure oxygen tension for oxygen consumption measurements from isolated mitochondria.
Commercial assay or kitClontech Labs 3P TaKaRa LA Taq DNA PolymeraseFisher ScientificCat. #50-443-973

Data availability

The small datasets generated in the study are included as source data files. The RNA-seq data has been deposited to NCBI Gene Expression Omnibus/Sequence Read Archive with the accession number GSE227295.

The following data sets were generated
    1. Coyne LP
    2. Chen XJ
    (2023) NCBI Gene Expression Omnibus
    ID GSE227295. Mitochondrial protein import clogging as a mechanism of disease.

References

    1. Simoncini C
    2. Siciliano G
    3. Tognoni G
    4. Mancuso M
    (2017)
    Mitochondrial ANT-1 related adPEO leading to cognitive impairment: is there a link?
    Acta Myologica 36:25–27.

Decision letter

  1. R Luke Wiseman
    Reviewing Editor; Scripps Research Institute, United States
  2. Benoît Kornmann
    Senior Editor; University of Oxford, United Kingdom
  3. Hilla Weidberg
    Reviewer; MIT, United States

Our editorial process produces two outputs: (i) public reviews designed to be posted alongside the preprint for the benefit of readers; (ii) feedback on the manuscript for the authors, including requests for revisions, shown below. We also include an acceptance summary that explains what the editors found interesting or important about the work.

Decision letter after peer review:

Thank you for submitting your article "Mitochondrial protein import clogging as a mechanism of disease" for consideration by eLife. Your article has been reviewed by 3 peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation has been overseen by Benoît Kornmann as the Senior Editor. The following individual involved in review of your submission has agreed to reveal their identity: Hilla Weidberg (Reviewer #3).

The reviewers have discussed their reviews with one another, and the Reviewing Editor has drafted this to help you prepare a revised submission.

Essential revisions:

1) The differentiation of the loss-of-function and gain-of-toxicity phenotypes associated with mutant AAC2 expression are well separated in the yeast models. However, the evidence supporting the specific contributions of gain-of-toxicity phenotypes in mammalian cell culture and mice expressing mutant ANT1 is less convincing. Please provide additional evidence to support the importance of gain-of-toxicity, as opposed to loss-of-function phenotypes induced by mutant ANT1 overexpression in these models.

2) The impact of ANT1/AAC2 mutant expression on biological processes including cytosolic/mitochondrial proteostasis, integrated stress response signaling, and phospholipid regulation should be expanded in a revised manuscript.

3) Please expand on the discussion regarding the mechanistic basis for clogging of mitochondrial protein import channels, as outlined in the reviewer comments.

Reviewer #1 (Recommendations for the authors):

In this manuscript, the authors evaluate the potential for destabilizing, missense mutations in ANT1 to 'clog' the mitochondrial protein import pathway and promote pathologic mitochondrial dysfunction in the context of disease. Initially using a yeast system, they show that two independent mutations in the yeast ANT1, AAC2, obstructs protein import into the mitochondria monitoring import both in vitro and in yeast. They go on to show that mutant AAC2 is degraded by the IMM AAA+ protease YME1 in yeast, consistent with this protein being clogged in the import pathway and being partially accessible within the IMS. They then go on to demonstrate that expression of mutant ANT in HeLa cells similarly appears to similarly impair import. Finally, they generate a mouse model expressing a 'super-clogger' ANT containing two mutations linked to impaired import in yeast/cells. Using these mice, they show that expression of this superclogger impairs mitochondrial protein import, disrupts mitochondrial function, and promotes age-dependent mitochondrial myopathy phenotypes.

This is an interesting manuscript that highlights the potential for 'clogging' of import channels by mutant proteins to promote mitochondrial dysfunction in disease. One of the challenges with this study is deconvoluting potential loss-of-function phenotypes associated with reductions in ANT1/AAC2 from gain-of-toxicity phenotypes linked to import clogging. This was addressed primarily in yeast, showing that phenotypes associated with overexpression of mutants (e.g., reduced growth on glucose media). The experiment showing that yeast AAC2 clogs import was also convincing including both in vitro and in vivo characterization, although it isn't clear why the proteomic experiments were performed with acute expression of A128P instead of the 'superclogger' double mutant. The extension of this work to mammalian cells and then mice is also admirable. However, the quality of characterization does begin to decline when moving into mammalian models. For example, there is no clear evidence that observed phenotypes can be attributed to gain of toxicity instead of loss of function in mammalian cells and mice. There are similarities to yeast, but this needs to be better defined. Lastly, I have questions related to the mouse model, such as how do these phenotypes compare to KO animals and why were homozygous mice used in some situations and heterozygous mice used in others.

Ultimately, the strengths of this manuscript lie in the yeast work. While the expansion into cells and mice is admirable, those characterizations are a bit weaker. I do think that it would be useful to allow authors an opportunity to address these concerns with additional experimentation and revision of the text, and I do think with additional characterization this manuscript would be of interest to the readership of eLife.

1. Does yme1 deletion increase interactions between mutant AAC2 and TOM subunits? One would expect that stabilizing the mutant protein should show even larger increases in interactions with the import complex.

2. It isn't clear to me why the MS experiments described in Figure 4 weren't performed with the double mutant 'superclogger', especially as this one showed the most interaction with TOM40 in Figure 2. I assume this is because the double mutant has lower levels, but regardless, one would expect that the double mutant would show larger recovery of cytosolic chaperones, TOM subunits, and TIM23 subunits. Could the authors explain their choice here? They claim that the A128P mutant blocks import of HSP60 in Figure S2B, which is fair, but it doesn't appear to do that in Figure 2H. Further, A128P doesn't seem to induce the same phenotypes as the double mutant 'superclogger' (e.g., Figure 1C). If the phenotypes are observed with A128P, what is the relevance of these increased interactions with import pathway? How can this be linked to import clogging? I guess the argument is linked to the acute expression in this experiment, but it isn't clear to me.

3. The transition into human cells is intriguing, but lacks some of the same necessary characterization performed in yeast. It would be useful to confirm that ANT1 mutants are causing gain of toxicity and not loss-of function by performing similar experiments in HeLa cells deficient in ANT1 to that performed in yeast growing cells in normal media or respiratory media, etc.

4. Along the same lines, as #2 above, it isn't clear to me why the authors probed interactions between A114P and TOM/TIM22 instead of the double mutant. If issues are coming with clogging, then one could expect that recovery of TOM/TIM22 subunits would be increased in double mutant purifications independent of expression levels of this mutant.

5. I don't understand how clogging of the TOM complex could lead to substrate specificity in mammalian cell models. Could the authors expand on that? I don't think it needs to be solved in this paper, but it is important to provide some context around this point.

6. ANT1-deficient mice have been generated and induce mitochondrial defects in muscle. How do the phenotypes observed in heterozygous mice expressing the 'superclogger' compare to the knockout mice? The authors indicate that the increase in maximal respiration observed at 9 months indicates that the observed phenotypes are independent of ATP/ADP transport, but this needs to be better characterized. Chronic changes could account for this.

7. It isn't clear to me why the authors use the homozygous mice to evaluate clogging in vivo, while the hets were used for all of the phenotypes. If the goal is to demonstrate phenotypes are linked to clogging, then why wouldn't the authors evaluate clogging in the context of the same mice used for phenotyping? What happens to the homozygous mice from a phenotypic perspective? One would expect it is worse considering the larger impact on 'clogging'.

Reviewer #2 (Recommendations for the authors):

There are some caveats to this work that I summarize below. After these points are largely addressed the study is well suitable for publication in eLife.

Specific points

1. The abstract states "We propose that secondary structures of mitochondrial preproteins play an essential role in preventing clogging and disease". While it is tempting to generalize the authors' findings, this statement is not supported by presented data or discussion.

2. When describing Figure 1C, the authors state that cells co-expressing aac2 A128P, A137D and AAC2 "form petites at much higher frequency…". They should provide actual numbers and include quantification to support this statement.

3. Figure 4A. Why is the Aac2p A128P mutant toxic at 25C but not at 30C? This observation is basically left unexplained.

4. Does mutant Aac2p/ANT1 expression affect mitochondrial phospholipids? One additional scenario to consider would be an impaired membrane homeostasis.

5. The graph shown in Figure 5F is confusing and not well explained. What exactly do those multiple lines depict?

6. Do mutant Aac2/ANT1 variants affect stability of the TOM40 or TOM22 complexes?

7. Does expression of mutant ANT1 induce integrated stress response (ISR) and whether some of the cytotoxic effects observed could be mitigated by ISR inhibition?

8. Would treating mice with pharmacological inhibitors of protein import (e.g. MitoBloCK-1) phenocopy the effects of mutant ANT1 expression? This would strengthen the present study and ascertain generality of the proposed model.

My suggestions are generally summarized in the Specific points section. I feel that addressing the potential involvement of phospholipids, probing mutant ANT1 contribution to integrated stress response, and testing whether pharmacological inhibition of import in vivo could mimic the effects in question would significantly strengthen the present study and its conclusions.

Reviewer #3 (Recommendations for the authors):

1. The experiments convincingly show that Aac2/Ant1 mutants clog TOM. However, the possibility that the mutants cause other type of stress, such as aggregation in the mitochondria or toxic accumulation in the cytosol, is not excluded. Therefore, the connection between clogging and the observed phenotypes is correlative and not necessarily causative. In fact, expression of Aac2 mutant in yeast deleted of Tom70 was synthetic lethal. As Tom70 is required for the entry of Aac2 to mitochondria and the TOM channel, this result might suggest that the Aac2 toxicity is caused by a cytosolic and not mitochondrial population of the mutant. The authors conclusion that this genetic interaction supports the clogging activity is thus unclear.

It will be helpful to clarify: 1. Were all mice phenotypes excluded in Ant1+/- mice? 2. Is it possible that some of the phenotypes, such as neurodegeneration, are due to a cytosolic burden caused by the Ant1 mutant, or misfolding of this mutant in the mitochondria, rather than TOM clogging?

As the evidence are still correlative the authors should tone down their conclusion and emphasize that the other possibilities were not excluded.

2. Do Aac2/Ant1 variants clog TOM, TIM22, or both? Including hypotheses in the discussion session could be helpful. In particular in regards to stabilization of mutant aac2 in yme1 deletion and not in proteasome impaired conditions, which demonstrated that Aac2 is degraded in the IMS and raises the possibility that the degradation is coupled to association with TIM22.

3. The involvement of Yme1 in Aac2's degradation is novel and exciting but not very well developed. Does mutant Aac2 interact with Yme1? Does Yme1-medaited degradation require interaction of mutant Aac2 with small TIMs? Does it require interaction with TIM22? Does Yme1 has a role in degrading other type of mitochondrial cloggers, such as Cyb2DHFR?

The above might be beyond the scope of the manuscript but could be at least raised in the discussion. In addition, Yme1-mediated degradation was shown using overexpression of Aac2A128P,A137D mutant. It would be beneficial to show stabilization of overexpressed Aac2A128P and Aac2A137D single mutants as well.

4. Figure S5- An empty vector control is missing to confirm that the overexpression of wild type Aac2 has no effect. The time frame of induction between the RPN4, HSP82, SSA3, SSA4 and CIS1 is very different. Addition of time points between 4h and 24h could clarify this issue.

5. Figure 7I- lines 317-8- "Moreover, Smac also appears to be more sensitive to proteinase K in Ant1A114P,A123D/Ant1A114P,A123D mitochondria, while Mdh2 and Tim23 does not (Figure S7I-K)." Smac is not the only protein that seems to be sensitive to proteinase K: a. Mdh2 levels are decreased as well as its size (appears as a smaller band), b. Tim23 levels seems lower as well, although its signal is oversaturated. A quantification of all proteins +/- protease could test whether they are less sensitive to the protease than Ant mutants. Same applies to Figure 5D.

The authors added the following helpful comment: "While this could indicate instability or partial rupture of the OMM in mutant mitochondria, it could also be a result of clogging and imply that clogging does not affect all mitochondrial preproteins equally. Consistent with this concept, our experiments in yeast showed minimal effects of clogging on Ilv5, a mitochondrial matrix protein, in contrast to Hsp60 (Figure 1E; 2H-I)." I agree with this conclusion and add that Tim23 is a TIM22 substrate and might be more sensitive to clogging of TIM22.

6. Figure 7G- The cytosolic fraction was collected after a relatively low speed centrifugation and might contain more than soluble proteins. For example, if the Ant1 mutants induce mitophagy or formation of MDVs, such structures might contain small mitochondrial fractions that could be detected as "cytosol".

https://doi.org/10.7554/eLife.84330.sa1

Author response

Essential revisions:

1) The differentiation of the loss-of-function and gain-of-toxicity phenotypes associated with mutant AAC2 expression are well separated in the yeast models. However, the evidence supporting the specific contributions of gain-of-toxicity phenotypes in mammalian cell culture and mice expressing mutant ANT1 is less convincing. Please provide additional evidence to support the importance of gain-of-toxicity, as opposed to loss-of-function phenotypes induced by mutant ANT1 overexpression in these models.

We appreciate this important point regarding loss- vs gain-of-function mechanisms. To provide additional evidence to support gain-of-toxicity of Ant1A114P,A123D (encoded by Slc25a4 p.A114P,A123D), we profiled the transcriptome of Slc25a4 p.A114P,A123D/+ mouse skeletal muscle, reasoning that if loss-of-function played a major role in Ant1A114P,A123D-induced pathology in mice, then the transcriptional signature should resemble that of Slc25a4 knock-out mice (Morrow et al., 2017). Slc25a4 knockout mouse muscle show a robust increase in the myokines Fgf21 and Gdf15 by up to 50-fold and 12-fold, respectively (Morrow et al., 2017). Fgf21 was not even detected in two of four Slc25a4 p.A114P,A123D/+ samples despite a sequencing depth of ~30 million reads/sample. Gdf15 was reduced by 25% (p = 0.77). Pathway analyses of significantly upregulated genes in Slc25a4 knockout muscle showed robust enrichment for genes involved in oxidative phosphorylation (OXPHOS) (Morrow et al., 2017). Using the same statistical tests and cutoffs, we didn’t find a single OXPHOS gene significantly upregulated in Slc25a4 p.A114P,A123D/+ muscle (see Data S6). Pgc-1a, the transcriptional regulator of mitochondrial biogenesis, was upregulated by 2.46-fold in Slc25a4 knockout muscle but unchanged in Slc25a4 p.A114P,A123D/+. Again, using the same statistical cutoffs and enrichment analysis software, we compared the top 25 most enriched pathways in significantly up- and down-regulated genes, and found only a single pathway of the 50 per genotype overlapped (Figure 8—figure supplement 2G). Clearly, the transcriptional signature of Slc25a4 p.A114P,A123D/+ skeletal muscle is entirely distinct from Slc25a4 knockout, excluding the possibility that loss of Ant1 function is the major pathomechanism in Slc25a4 p.A114P,A123D/+ mice.

Other comparisons with Slc25a4 knockout mice are illuminating as well. Respirometry of mitochondria isolated from Slc25a4 p.A114P,A123D/+ skeletal muscle showed a significant increase in maximal respiration rate when complex II is stimulated (Figure 6). As we noted in our original manuscript, this indicates that one functional copy of Ant1 is sufficient to support high respiratory rates, arguing against ADP/ATP transport as a limiting factor. What we failed to note in our original manuscript is that this is not observed in Slc25a4 knockout mice; in fact, complex II-based respiration is reduced by >30% in Slc25a4 knockout mice (Graham et al., Nature Genetics, 1997). We have now added this to the manuscript along with additional observations that strongly argue against loss of Ant1 function as a primary driver of Slc25a4 p.A114P,A123D/+ muscle phenotypes.

Additional clinical context may be informative as well, and we have clarified this point in the revised manuscript. Patients heterozygous for dominant SLC25A4 mutations, such as p.A114P, exhibit clinical features that are not present in patients with homozygous loss of ANT1 function, such as neurodegenerative and neuropsychiatric phenotypes (Deschauer et al., 2005; Echaniz-Laguna et al., 2012; Kashiki et al., 2022; Kaukonen et al., 2000; Napoli et al., 2001; Palmieri et al., 2005; Siciliano et al., 2003; Simoncini et al., 2017; Thompson et al., 2016; Tosserams et al., 2018). This strongly implies gain-of-function toxicity in patients. Moreover, some Slc25a4 p.A114P,A123D/+ mice undergo frank neurodegeneration ending in paralysis and death within 2-3 weeks of symptom onset, while there are no neurological features of Slc25a4 knockout mice. This is, in and of itself, a gain-of-function phenotype.

The transition to mammalian cell culture was intended to biochemically test if mutant ANT1 clogs protein import like yeast Aac2p. From this perspective, the original manuscript clearly demonstrated biochemical gain-of-function phenotypes, i.e. mutant ANT1 accumulation at the import machinery, mutant ANT1 being exposed on the outer membrane, and mitochondrial proteins accumulating in the cytosol. These biochemical phenotypes cannot reasonably be expected to result from loss of ANT1 function. We also note that total ANT levels (including ANT1, ANT2 and ANT3) can be drastically depleted in human cell culture without severely affecting OXPHOS or cell viability (Lu et al., 2017).

However, we absolutely agree with the broader point that at this stage we cannot completely exclude some contribution of haploinsufficiency in the context of clogging. We have adjusted our language in the manuscript accordingly.

2) The impact of ANT1/AAC2 mutant expression on biological processes including cytosolic/mitochondrial proteostasis, integrated stress response signaling, and phospholipid regulation should be expanded in a revised manuscript.

– To assess the impact on cytosolic proteostasis, we performed detailed analyses on the cytosolic proteome from both mammalian cell culture and mouse skeletal muscle, which showed modest increase in proteostatic stress signaling and chaperones, respectively. These data suggest possible cytosolic proteostatic stress. The data can be found in Figure 5H, Figure 5—figure supplement 1A-B, and Figure 8—figure supplement 1A-B.

– To assess the impact on mitochondrial proteostasis, we performed BN-PAGE analysis and found that, in mice, the respiratory complexes, respiratory supercomplexes, and Tim23 complex are minimally affected in Slc25a4 p.A114P,A123D/+ mouse muscle mitochondria (Figure 8—figure supplement 1C-E). This suggests no severe effect on proteostasis on the inner mitochondrial membrane. We also performed quantitative proteomics on mitochondria isolated from mouse muscle (Figure 8—figure supplement 1F). While the mitochondrial proteome was overall minimally affected, we did observe a general increase in the levels of mitochondrial chaperones and proteases (Figure 8—figure supplement 1), suggesting there is at least a low level of mitochondrial proteostatic stress.

– To assess the impact on the integrated stress response (ISR), we performed western blot analysis of eIF2a and also performed RNA sequencing of Slc25a4 p.A114P,A123D/+ mouse skeletal muscle, as described above. We found no evidence of ISR activation (Figure 8—figure supplement 2A-C).

– To explore the impact on phospholipid homeostasis, we used yeast genetics to test whether cells defective in cardioipin (CL) (pel1D) and/or phosphatidylethanolamine (PE) synthesis (psd1D) are hypersensitive to expression a super-clogger aac2 variant. Indeed, we found that expression of aac2A128P,A137D strongly inhibited the growth of cells in both genetic backgrounds (Figure 4—figure supplement 3G-H). The data are consistent with previous observations that CL and PE facilitate mitochondrial protein import (Becker et al., 2013; Gebert et al., 2009). Alternatively, mitochondrial protein import clogging may affect phospholipid homoeostasis, which synergizes with pel1D and psd1D to cause cell lethality.

3) Please expand on the discussion regarding the mechanistic basis for clogging of mitochondrial protein import channels, as outlined in the reviewer comments.

We expanded discussion on the mechanistic basis for clogging in the revised manuscript.

Reviewer #1 (Recommendations for the authors):

In this manuscript, the authors evaluate the potential for destabilizing, missense mutations in ANT1 to 'clog' the mitochondrial protein import pathway and promote pathologic mitochondrial dysfunction in the context of disease. Initially using a yeast system, they show that two independent mutations in the yeast ANT1, AAC2, obstructs protein import into the mitochondria monitoring import both in vitro and in yeast. They go on to show that mutant AAC2 is degraded by the IMM AAA+ protease YME1 in yeast, consistent with this protein being clogged in the import pathway and being partially accessible within the IMS. They then go on to demonstrate that expression of mutant ANT in HeLa cells similarly appears to similarly impair import. Finally, they generate a mouse model expressing a 'super-clogger' ANT containing two mutations linked to impaired import in yeast/cells. Using these mice, they show that expression of this superclogger impairs mitochondrial protein import, disrupts mitochondrial function, and promotes age-dependent mitochondrial myopathy phenotypes.

This is an interesting manuscript that highlights the potential for 'clogging' of import channels by mutant proteins to promote mitochondrial dysfunction in disease. One of the challenges with this study is deconvoluting potential loss-of-function phenotypes associated with reductions in ANT1/AAC2 from gain-of-toxicity phenotypes linked to import clogging. This was addressed primarily in yeast, showing that phenotypes associated with overexpression of mutants (e.g., reduced growth on glucose media). The experiment showing that yeast AAC2 clogs import was also convincing including both in vitro and in vivo characterization, although it isn't clear why the proteomic experiments were performed with acute expression of A128P instead of the 'superclogger' double mutant. The extension of this work to mammalian cells and then mice is also admirable. However, the quality of characterization does begin to decline when moving into mammalian models. For example, there is no clear evidence that observed phenotypes can be attributed to gain of toxicity instead of loss of function in mammalian cells and mice. There are similarities to yeast, but this needs to be better defined. Lastly, I have questions related to the mouse model, such as how do these phenotypes compare to KO animals and why were homozygous mice used in some situations and heterozygous mice used in others.

Ultimately, the strengths of this manuscript lie in the yeast work. While the expansion into cells and mice is admirable, those characterizations are a bit weaker. I do think that it would be useful to allow authors an opportunity to address these concerns with additional experimentation and revision of the text, and I do think with additional characterization this manuscript would be of interest to the readership of eLife.

We thank the reviewer for the thoughtful comments. Regarding the loss vs gain-of-function phenotypes, please see the discussion above. We agree with the reviewer that the extent of characterization in the mammalian systems is less than that in yeast, but note that the original manuscript did contain substantial data from mammalian cells (11 panels of data and 1 data file) and our novel mouse model (31 panels of data, 1 data file, and 2 movies). Nevertheless, we put forth a good-faith effort to better characterize the mammalian systems, as follows:

  • More extensive analysis of the cytosolic proteome in human cells (Figure 5H) and mouse muscle (Figure 8—figure supplement 1A-B).

  • Additional isolation of mouse muscle mitochondria for native gel electrophoresis followed by Coomassie staining, which can visualize the mitochondrial respiratory complexes and supercomplexes (Figure 8—figure supplement 1C). This is coupled with native gel electrophoresis with western blot to further assess the mitochondrial complexes/supercomplexes and the TIM23 complex (Figure 8—figure supplement 1D-E)

  • To further implement reviewer’s comments, we analyzed mitochondrial proteomics in Slc25a4 p.A114P,A123D/+ mouse skeletal muscle (Figure 8—figure supplement 1F-H).

  • Immunoblot validation of Tim22 increase suggested by mitochondrial proteomics data (Figure 8—figure supplement 1I-J).

  • Immunoblot analysis of eIF2a in Slc25a4 p.A114P,A123D/+ mouse skeletal muscle to assess integrated stress response signaling (Figure 8—figure supplement 2A-B)

  • Finally, we used RNA-sequencing and determine transcriptional changes in Slc25a4 p.A114P,A123D/+ mouse skeletal muscle (Figure 8—figure supplement 2C-G)

1. Does yme1 deletion increase interactions between mutant AAC2 and TOM subunits? One would expect that stabilizing the mutant protein should show even larger increases in interactions with the import complex.

We thank the reviewer for this excellent question. The interaction between mutant Aac2p and Yme1p is complex. With acute, inducible expression of mutant Aac2p (as in Figure 3), we would expect that yme1 deletion would increase Aac2p’s association with the TOM complex as suggested by the reviewer. However, deletion of Yme1p does not increase levels of chronically expressed Aac2pA128P (Liu et al., 2015), and actually leads to the general reduction in the biogenesis of wild-type carrier proteins (Kumar et al., 2023). The mechanistic details of Yme1p-based proteolysis of mutant Aac2p and potential anti-clogging activity will be important areas for future research but may be beyond the scope of the current study.

2. It isn't clear to me why the MS experiments described in Figure 4 weren't performed with the double mutant 'superclogger', especially as this one showed the most interaction with TOM40 in Figure 2. I assume this is because the double mutant has lower levels, but regardless, one would expect that the double mutant would show larger recovery of cytosolic chaperones, TOM subunits, and TIM23 subunits. Could the authors explain their choice here?

The reviewer is correct, we elected not to pull down double mutant Aac2p for mass spectrometry of co-eluted proteins because of the extremely low protein accumulation. From a technical standpoint, this experiment is unlikely to yield any interpretable information. First, co-IP experiments with wild-type Aac2p/ANT1 is notorious for yielding a large number of proteins, many of which are likely associating nonspecifically due to the protein-dense, two-dimensional environment of the inner mitochondrial membrane (Claypool et al., 2008; Lu et al., 2017). Given this high background in wild-type Aac2p/ANT1, reducing the level of the protein >20-fold will even further reduce the signal-to-noise ratio. Second, the sensitivity of mass spectrometry compounds this problem. In our experiment, the control samples lacking a HIS tag still reliably eluted >1,500 proteins, hence our workflow to eliminate this increased noise, as shown in Figure 4—figure supplement 1B. In our view, repeating this experiment with the double mutant is unlikely to be helpful.

They claim that the A128P mutant blocks import of HSP60 in Figure S2B, which is fair, but it doesn't appear to do that in Figure 2H.

In Figure 2H, expression of the mutant is from a centromeric vector and cells were grown in minimal selective medium against plasmid loss. Growth in minimal medium is expected to globally reduce protein synthesis and alleviate cytosolic proteostatic stress (mPOS). In addition, in Figure 2—figure supplement 1B, the mutant aac2 allele is expressed from a galactose-inducible promoter and the cells were grown in complete medium. We expect that the latter has higher expression of the mutant protein, which could also contribute to increased effect on protein import. However, we note that pre-Hsp60p does accumulate at steady state when mutant aac2 is integrated into the genome and cells were grown in compete medium (Figure 2G), which may be most physiologically relevant.

Further, A128P doesn't seem to induce the same phenotypes as the double mutant 'superclogger' (e.g., Figure 1C). If the phenotypes are observed with A128P, what is the relevance of these increased interactions with import pathway? How can this be linked to import clogging? I guess the argument is linked to the acute expression in this experiment, but it isn't clear to me.

Regarding lack of severe phenotype of single mutants in Figure 1C: First, W303 is well known to be particularly tolerant to many mitochondrial insults, including single mutant aac2 mutants (Wang et al., 2008a). Second, and perhaps relatedly, aac2 mutants are rho-zero lethal in non-W303 strains, meaning these mutants cannot survive the loss of mtDNA. Thus, to (1) increase the probably that the strain is viable, and (2) allow us to observe the effect on mtDNA stability, we chose to integrate the double mutant into the W303 background.

The single mutants (including A128P) are highly toxic in M2915-6A background, which is the background in which we observed Aac2pA128P accumulation at the translocase complexes (Figure 4). A more in-depth explanation of the strain-dependent effects of mitochondrial dysfunction on cell viability can be found in our response to Reviewer 3.

3. The transition into human cells is intriguing, but lacks some of the same necessary characterization performed in yeast. It would be useful to confirm that ANT1 mutants are causing gain of toxicity and not loss-of function by performing similar experiments in HeLa cells deficient in ANT1 to that performed in yeast growing cells in normal media or respiratory media, etc.

As noted above, severe depletion of all ADP/ATP transporter isoforms is surprisingly very well-tolerated by in mammalian cells in culture, even in regards to bioenergetics (Lu et al., 2017). The implication of this finding is that a very low level of ADP/ATP carriers is sufficient to support OXPHOS, which is surprising but consistent with our bioenergetic data from Slc25a4 p.A114P,A123D/+ mice. In our view, performing CRISPR knockout of SLC25A4 in HeLa cells followed by respiratory growth assay would have a limited contribution to the overall message of the manuscript, especially considering the considerable evidence of gain-of-toxicity and substantial bioenergetic characterization already present in mice.

4. Along the same lines, as #2 above, it isn't clear to me why the authors probed interactions between A114P and TOM/TIM22 instead of the double mutant. If issues are coming with clogging, then one could expect that recovery of TOM/TIM22 subunits would be increased in double mutant purifications independent of expression levels of this mutant.

Please see the above explanation (response to #2) explaining our decision not to immunoprecipitate Ant1A114P,A123D, which accumulates to just 2.2% of the wild-type level.

5. I don't understand how clogging of the TOM complex could lead to substrate specificity in mammalian cell models. Could the authors expand on that? I don't think it needs to be solved in this paper, but it is important to provide some context around this point.

This is a very interesting question. The data in this study suggest that there is substrate specific TOM complex clogging in both yeast and mammalian models. One likely contributor is the observation that the Tom40 b-barrel pore contains several distinct protein paths (Araiso et al., 2019; Shiota et al., 2015). Thus, although the pore is narrow, it is certainly conceivable that some protein paths are more affected than others. This explanation has been added to the manuscript (Lines 469-471).

6. ANT1-deficient mice have been generated and induce mitochondrial defects in muscle. How do the phenotypes observed in heterozygous mice expressing the 'superclogger' compare to the knockout mice? The authors indicate that the increase in maximal respiration observed at 9 months indicates that the observed phenotypes are independent of ATP/ADP transport, but this needs to be better characterized. Chronic changes could account for this.

We apologize for not making the phenotype comparison clearer in the original manuscript. A more rigorous comparison can be found above (in response to Essential Revisions) and in the revised manuscript (Figure 8—figure supplement 2G as well as an expanded Discussion section).

7. It isn't clear to me why the authors use the homozygous mice to evaluate clogging in vivo, while the hets were used for all of the phenotypes. If the goal is to demonstrate phenotypes are linked to clogging, then why wouldn't the authors evaluate clogging in the context of the same mice used for phenotyping? What happens to the homozygous mice from a phenotypic perspective? One would expect it is worse considering the larger impact on 'clogging'.

We bred a small number of homozygous mice to detect and characterize the low-abundance mutant protein, which is otherwise overwhelmed by the wildtype in heterozygous mice (Figure 8A-E). We deliberately refrained from phenotypic characterization of homozygous mice because the A123D mutation in Ant1A114P,A123D renders the protein unable to transport ADP/ATP (see (Palmieri et al., 2005), which was corroborated in Figure 1C). Thus, disentangling gain-of-function vs loss of function in homozygous mice would be impossible.

Reviewer #2 (Recommendations for the authors):

There are some caveats to this work that I summarize below. After these points are largely addressed the study is well suitable for publication in eLife.

Specific points

1. The abstract states "We propose that secondary structures of mitochondrial preproteins play an essential role in preventing clogging and disease". While it is tempting to generalize the authors' findings, this statement is not supported by presented data or discussion.

We thank the reviewer for pointing this out. We can understand how this statement may have been more misleading than we intended, as there are no structural data in the manuscript. We modified this statement to be more precise about what we do show, i.e. that amino acid substitutions in a carrier protein unexpectedly cause clogging.

2. When describing Figure 1C, the authors state that cells co-expressing aac2 A128P, A137D and AAC2 "form petites at much higher frequency…". They should provide actual numbers and include quantification to support this statement.

This experiment has been repeated, quantified, and depicted graphically in the revised manuscript (Figure 1E).

3. Figure 4A. Why is the Aac2p A128P mutant toxic at 25C but not at 30C? This observation is basically left unexplained.

Cold sensitivity of mutant aac2 strains has been observed for many years (Wang et al., 2008a; Wang et al., 2008b). We previously found that, specifically at 25C, mutant aac2 strains lose membrane potential which is associated with accumulation of the precursor form of Hsp60p (Wang et al., 2008b). With subsequent studies (Wang and Chen, 2015) and now the discovery of import clogging, it is likely the cold-induced reduction in membrane potential synergizes with clogging to further reduce protein import and challenge cytosolic proteostasis with the accumulation of mitochondrial precursors.

4. Does mutant Aac2p/ANT1 expression affect mitochondrial phospholipids? One additional scenario to consider would be an impaired membrane homeostasis.

This is an interesting idea that we previously investigated in single mutant aac2, hypothesizing that Aac2p misfolding causes membrane stress (Liu et al., 2015). To assess membrane protein homeostasis in double mutant Ant1 the mammalian systems, we performed a series of BN-PAGE experiments using human and mouse mitochondria. If membrane homeostasis is severely affected, we would expect a reduction in the assembly state of membrane protein complexes. However, we did not see a drastic effect in transfected HeLa cells (see Response Figure 1 below) or Slc25a4 p.A114P,A123D/+ mice (Figure 8-fuigure supplement 1C-E). Thus, proteostatic stress on the IMM may contribute to cell stress independently of clogging or simply only occur in the yeast mutants.

Prompted by the comment on mitochondrial phospholipids, we also investigated the effect of mutant aac2 expression in the context of defective mitochondrial phospholipid production. This was described above and the data can be found in the revised manuscript (Figure 4—figure supplement 3G-H).

5. The graph shown in Figure 5F is confusing and not well explained. What exactly do those multiple lines depict?

Each line was meant to connect the same protein found in the three samples (WT, ANT1A114P, and ANT1A114P,A123D-transfected cytosolic fractions). The idea was to show that there were a subset of proteins that were increased in ANT1A114P cells compared to WT, and then further increased in ANT1A114P,A123D cells. This is not an essential point so we elected to remove the lines to avoid further confusion.

6. Do mutant Aac2/ANT1 variants affect stability of the TOM40 or TOM22 complexes?

This is an interesting question, especially in light of our previous work that showed single mutant Aac2p proteins causing TIM22 destabilization (Liu et al., 2015). However, we did not observe this in transfected human cells, where mutant ANT1-transfected cells are essentially indistinguishable from wild-type (Response Figure 1). This suggests that IMM complexes in immortalized human cells may be less susceptible to mutant ANT1 relative to yeast.

Author response image 1
Mutant ANT1 does not drastically affect the assembly state of TIM22 complex, the TOM complex, or respiratory Complex I in HeLa cells.

(A-C) BN-PAGE followed by immunoblot analysis of isolated mitochondria from transfected HeLa cells. Protein ladders deduced from Coommassie-stained gel shown on the left. B and C are depicting the same membrane with sequential blotting. Regarding methods, mitochondrial isolation, BN-PAGE, and subsequent western blotting were performed 24 hours after transfection as previously described (Timon-Gomez et al., 2020) solubilizing the complexes with a 1:2 protein:digitonin ratio.

7. Does expression of mutant ANT1 induce integrated stress response (ISR) and whether some of the cytotoxic effects observed could be mitigated by ISR inhibition?

This is another very interesting question. As shown in the revised manuscript, we rigorously evaluated whether the ISR is activated in Slc25a4 p.A114P,A123D/+ mice and found no evidence to support this (Figure 8—figure supplement 2A-F). The data suggest that clogging with Ant1A114P,A123D is a unique and/or potentially mild stressor in vivo given the highly efficient degradation of the protein, as suggested by extremely low mutant protein accumulation (Figure 8A-C).

8. Would treating mice with pharmacological inhibitors of protein import (e.g. MitoBloCK-1) phenocopy the effects of mutant ANT1 expression? This would strengthen the present study and ascertain generality of the proposed model.

Treating mice with MitoBloCK-1 would be an extremely exciting experiment. We predict that its effect in vivo would largely depend on its bioavailability and tissue distribution, and may be unlikely to replicate the effects of mutant Ant1 because Ant1 expression is largely restricted to the muscle, heart and central nervous system. Given this caveat, the likely result of a distinct phenotype in MitoBloCK-1-treated mice would not provide much valuable information on the clogging model. Nevertheless, it could provide additional hypothesis-generating observations (which tissues are most susceptible to protein import defects and why?) so we thank the reviewer for this suggestion.

Reviewer #3 (Recommendations for the authors):

1. The experiments convincingly show that Aac2/Ant1 mutants clog TOM. However, the possibility that the mutants cause other type of stress, such as aggregation in the mitochondria or toxic accumulation in the cytosol, is not excluded. Therefore, the connection between clogging and the observed phenotypes is correlative and not necessarily causative. In fact, expression of Aac2 mutant in yeast deleted of Tom70 was synthetic lethal. As Tom70 is required for the entry of Aac2 to mitochondria and the TOM channel, this result might suggest that the Aac2 toxicity is caused by a cytosolic and not mitochondrial population of the mutant. The authors conclusion that this genetic interaction supports the clogging activity is thus unclear.

It will be helpful to clarify: 1. Were all mice phenotypes excluded in Ant1+/- mice? 2. Is it possible that some of the phenotypes, such as neurodegeneration, are due to a cytosolic burden caused by the Ant1 mutant, or misfolding of this mutant in the mitochondria, rather than TOM clogging?

As the evidence are still correlative the authors should tone down their conclusion and emphasize that the other possibilities were not excluded.

We thank the reviewer for this important comment. We completely agree that we cannot exclude all possible mechanisms by which mutant Aac2p/ANT1 might be toxic, and have toned down our conclusion and emphasized our openness to other possibilities in the revised manuscript.

We also apologize for not making our working model clear in the manuscript. We expect that import clogging has far-reaching consequences, both intra- and extra-mitochondrial. Particularly in the cytosol, we absolutely expect that one of the effects of protein import clogging would be toxic mitochondrial protein accumulation and aggregation, which could include mutant Aac2p/ANT1 “waiting in the traffic jam”. This would be coherent with a model of clogging-induced toxicity.

With that said, there are some key experiments that argue against predominant cytosolic aggregation of mutant Aac2p/ANT1 independent of clogging. First, in our in vitro import experiments, the mutant proteins have increased association with the TOM complex, suggesting they are properly delivered to the mitochondria surface (Figure 2E-F). Second, we were unable to detect an increase in mutant Aac2p levels when using harsh solubilization conditions on whole cells (Figure 1—figure supplement 1B-C), suggesting the protein is not aggregated in the cytosol or mitochondria. We were similarly unable to extract any potentially aggregated quadruple mutant ANT1A90D, L98P,A114P,A123D from transfected human cells (Liu et al., 2019). Thus, we think it’s unlikely that cytosolic aggregation of mutant Aac2p/ANT1 in the absence of clogging is a major mechanism of toxicity.

We thank the reviewer for her careful thinking regarding the genetic interaction between mutant aac2 and tom70D. We would like to clarify that Tom70p is not required for the entry of Aac2 into mitochondria and, surprisingly, deletion of TOM70 and its paralog TOM71 only reduce Aac2p biogenesis by ~50% (Backes et al., 2021). Tom20p and Tom22p serve as receptors for carrier proteins in the absence of Tom70p (Steger et al., 1990; Yamano et al., 2008). In this case, the mutant Aac2 would still be able to engage in clogging. Interestingly, it was recently proposed that Tom70p’s primary function is to recruit chaperones to the mitochondrial surface to prevent proteotoxicity in this protein-dense region (Backes et al., 2021). Thus, deletion of TOM70 may affect aac2A128P cells by two mechanisms: first, by further reducing carrier protein import and second by impairing cytosolic proteostasis in the context of pre-existing mitochondrial protein overaccumulation in the cytosol (Wang and Chen, 2015).

Regarding intramitochondrial protein aggregation, this was actually our original hypothesis for the mechanism of toxicity of double mutant Aac2p based on the aggregating properties of the mutant proteins in isolated mitochondria (Liu et al., 2015). However, we found no evidence of severe proteostatic stress on the inner mitochondrial membrane in the mammalian cells or mice, as discussed above.

Unfortunately, we are unable to exclude all phenotypes in Slc25a4+/- mice as we do not possess this mouse model. However, we do note that a neurodegenerative phenotype has never been reported in Slc25a4+/- or Slc25a4-/- mice despite >25 years of characterization by Doug Wallace and colleagues. Likewise, a myopathy has never been reported in Slc25a4+/- mice, consistent with normal clinical, biochemical and pathological features in heterozygous loss of function humans (Echaniz-Laguna et al., 2012).

2. Do Aac2/Ant1 variants clog TOM, TIM22, or both? Including hypotheses in the discussion session could be helpful. In particular in regards to stabilization of mutant aac2 in yme1 deletion and not in proteasome impaired conditions, which demonstrated that Aac2 is degraded in the IMS and raises the possibility that the degradation is coupled to association with TIM22.

We thank the reviewer for this helpful suggestion and have clarified our thoughts in the discussion in the revised manuscript.

3. The involvement of Yme1 in Aac2's degradation is novel and exciting but not very well developed. Does mutant Aac2 interact with Yme1? Does Yme1-medaited degradation require interaction of mutant Aac2 with small TIMs? Does it require interaction with TIM22? Does Yme1 has a role in degrading other type of mitochondrial cloggers, such as Cyb2DHFR?

The above might be beyond the scope of the manuscript but could be at least raised in the discussion. In addition, Yme1-mediated degradation was shown using overexpression of Aac2A128P,A137D mutant. It would be beneficial to show stabilization of overexpressed Aac2A128P and Aac2A137D single mutants as well.

These are excellent questions for future investigation and we have added some of these ideas to the discussion in the revised manuscript.

4. Figure S5- An empty vector control is missing to confirm that the overexpression of wild type Aac2 has no effect. The time frame of induction between the RPN4, HSP82, SSA3, SSA4 and CIS1 is very different. Addition of time points between 4h and 24h could clarify this issue.

We apologize for not being clearer in the figure legend; this has been corrected. The strains used in Figure 4—figure supplement 3B-F (formerly Figure S5B-F) have a chromosomally integrated copy of wild-type and mutant aac2 driven by the GAL10 promoter from the trp1 locus.

5. Figure 7I- lines 317-8- "Moreover, Smac also appears to be more sensitive to proteinase K in Ant1A114P,A123D/Ant1A114P,A123D mitochondria, while Mdh2 and Tim23 does not (Figure S7I-K)." Smac is not the only protein that seems to be sensitive to proteinase K: a. Mdh2 levels are decreased as well as its size (appears as a smaller band), b. Tim23 levels seems lower as well, although its signal is oversaturated. A quantification of all proteins +/- protease could test whether they are less sensitive to the protease than Ant mutants. Same applies to Figure 5D.

We thank the reviewer for pointing this out. We agree, there does seem to be some sensitivity of TIM23 and MDH2 to proteinase K, but this is the case in wild-type mitochondria. and there is no difference between wild-type and mutant mitochondria. We have quantified all proteins +/- protease, relative to untreated, and there is no difference between wild-type and mutant samples. We moved these data from supplemental to a main figure to consolidate the data (Figure 8E-H).

Similarly, in Figure 5D, there is some level of sensitivity to proteinase K even in wild-type ANT1-transfected cells. For example, the mitochondrial matrix protein TFAM is reduced in each proteinase K-treated sample with or without swelling. This likely represents physical disruption of a fraction of mitochondria during isolation. We controlled for this by normalizing the ANT1-HA level to the level of TFAM. We appreciate the reviewer’s keen eye in this regard.

The authors added the following helpful comment: "While this could indicate instability or partial rupture of the OMM in mutant mitochondria, it could also be a result of clogging and imply that clogging does not affect all mitochondrial preproteins equally. Consistent with this concept, our experiments in yeast showed minimal effects of clogging on Ilv5, a mitochondrial matrix protein, in contrast to Hsp60 (Figure 1E; 2H-I)." I agree with this conclusion and add that Tim23 is a TIM22 substrate and might be more sensitive to clogging of TIM22.

6. Figure 7G- The cytosolic fraction was collected after a relatively low speed centrifugation and might contain more than soluble proteins. For example, if the Ant1 mutants induce mitophagy or formation of MDVs, such structures might contain small mitochondrial fractions that could be detected as "cytosol".

This caveat has been added to the manuscript.

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https://doi.org/10.7554/eLife.84330.sa2

Article and author information

Author details

  1. Liam P Coyne

    Department of Biochemistry and Molecular Biology, State University of New York Upstate Medical University, Syracuse, United States
    Contribution
    Conceptualization, Data curation, Formal analysis, Funding acquisition, Validation, Investigation, Visualization, Methodology, Writing – original draft, Project administration, Writing – review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-4480-126X
  2. Xiaowen Wang

    Department of Biochemistry and Molecular Biology, State University of New York Upstate Medical University, Syracuse, United States
    Contribution
    Data curation, Formal analysis, Validation, Investigation, Visualization, Methodology
    Competing interests
    No competing interests declared
  3. Jiyao Song

    1. Institute of Biochemistry and Molecular Biology, Faculty of Medicine, University of Freiburg, Freiburg, Germany
    2. Institute of Biochemistry and Molecular Biology, Faculty of Medicine, University of Bonn, Bonn, Germany
    Contribution
    Formal analysis, Validation, Investigation, Visualization, Methodology
    Competing interests
    No competing interests declared
  4. Ebbing de Jong

    Proteomics and Mass Spectrometry Core Facility, State University of New York Upstate Medical University, Syracuse, United States
    Contribution
    Data curation, Validation, Methodology
    Competing interests
    No competing interests declared
  5. Karin Schneider

    Department of Microbiology and Immunology, State University of New York Upstate Medical University, Syracuse, United States
    Contribution
    Data curation, Visualization
    Competing interests
    No competing interests declared
  6. Paul T Massa

    1. Department of Microbiology and Immunology, State University of New York Upstate Medical University, Syracuse, United States
    2. Department of Neurology, State University of New York Upstate Medical University, Syracuse, United States
    Contribution
    Formal analysis, Supervision, Validation, Investigation, Writing – review and editing
    Competing interests
    No competing interests declared
  7. Frank A Middleton

    1. Department of Biochemistry and Molecular Biology, State University of New York Upstate Medical University, Syracuse, United States
    2. Department of Neuroscience and Physiology, State University of New York Upstate Medical University, Syracuse, United States
    Contribution
    Formal analysis, Supervision, Writing – review and editing
    Competing interests
    No competing interests declared
  8. Thomas Becker

    Institute of Biochemistry and Molecular Biology, Faculty of Medicine, University of Bonn, Bonn, Germany
    Contribution
    Data curation, Formal analysis, Supervision, Funding acquisition, Validation, Visualization, Methodology, Writing – review and editing
    Competing interests
    No competing interests declared
  9. Xin Jie Chen

    1. Department of Biochemistry and Molecular Biology, State University of New York Upstate Medical University, Syracuse, United States
    2. Department of Neuroscience and Physiology, State University of New York Upstate Medical University, Syracuse, United States
    Contribution
    Conceptualization, Resources, Data curation, Formal analysis, Supervision, Funding acquisition, Validation, Investigation, Visualization, Methodology, Writing – original draft, Project administration, Writing – review and editing
    For correspondence
    chenx@upstate.edu
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-8488-6587

Funding

National Institute on Aging (R01AG063499)

  • Xin Jie Chen

National Institute on Aging (R01AG061204)

  • Xin Jie Chen

National Institute on Aging (F30AG060702)

  • Liam P Coyne

Deutsche Forschungsgemeinschaft (project ID 269925409)

  • Thomas Becker

Deutsche Forschungsgemeinschaft (BE 4679 2/2)

  • Thomas Becker

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

We thank Nikolaus Pfanner for support and anti-sera, Joyce Qi for help with electron microscopy, and Siu-Pok Yee (University of Connecticut) for Slc25a4 p.A114P,A123D knock-in mouse generation. We’re also grateful to Yumiko Umino and Eduardo Solessio for confirming visual acuity and contrast sensitivity in Slc25a4 p.A114P,A123D/+ mice.

Ethics

This study was performed in strict accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health. All of the animals were handled according to approved institutional animal care and use committee (IACUC) protocols (D1600318) of the State University of New York Upstate Medical University. The protocol was approved by the Committee on the Ethics of Animal Experiments of SUNY Upstate Medical University (Permit Number: #268).

Senior Editor

  1. Benoît Kornmann, University of Oxford, United Kingdom

Reviewing Editor

  1. R Luke Wiseman, Scripps Research Institute, United States

Reviewer

  1. Hilla Weidberg, MIT, United States

Version history

  1. Preprint posted: September 21, 2022 (view preprint)
  2. Received: October 20, 2022
  3. Accepted: April 17, 2023
  4. Accepted Manuscript published: May 2, 2023 (version 1)
  5. Version of Record published: May 24, 2023 (version 2)

Copyright

© 2023, Coyne et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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  1. Liam P Coyne
  2. Xiaowen Wang
  3. Jiyao Song
  4. Ebbing de Jong
  5. Karin Schneider
  6. Paul T Massa
  7. Frank A Middleton
  8. Thomas Becker
  9. Xin Jie Chen
(2023)
Mitochondrial protein import clogging as a mechanism of disease
eLife 12:e84330.
https://doi.org/10.7554/eLife.84330

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https://doi.org/10.7554/eLife.84330