The mitochondrial electron transport chain (ETC) supports myriad, mechanistically linked functions. It provides the driving force for ATP synthesis by oxidative phosphorylation (OXPHOS) (Mitchell, 1961; Mitchell, 2011), maintains redox balance of cofactor pairs (NADH:NAD⁺, CoQH₂:CoQ) that are coupled to hundreds of cellular reactions (Williamson et al., 1967; Ying, 2008; Santidrian et al., 2013; Titov et al., 2016; Xiao et al., 2018; Goodman et al., 2018; Mitchell, 1975; Turunen et al., 2004; Alcázar-Fabra et al., 2016), contributes to the electrochemical gradient that drives transport of proteins and metabolites across the mitochondrial inner-membrane (LaNoue and Schoolwerth, 1979; Martin et al., 1991), and is a major determinant of cellular oxygen levels and tolerance (Taivassalo et al., 2002; Campian et al., 2004; O'Hagan et al., 2009; Brand, 2016; Jain et al., 2019). Defects in the ETC and OXPHOS system are associated with a spectrum of human pathology ranging from individually rare genetic disorders to common conditions, such as neurodegeneration (Schapira, 2008; Hu and Wang, 2016; Lin and Beal, 2006; Area-Gomez et al., 2019), diabetes (Kelley et al., 2002; Mootha et al., 2004; Ritov et al., 2005; Ritov et al., 2010), select forms of cancer (Wallace, 2012; Reznik et al., 2017; Gaude and Frezza, 2014) and the aging process itself (Sun et al., 2016; Bratic and Larsson, 2013).

Mutations in nearly 300 nuclear or mitochondrial (mtDNA) genes have been implicated in mitochondrial disorders that affect at least 1 in 4300 live births and lack effective treatments (Schapira, 2012; Vafai and Mootha, 2012; Gorman et al., 2015; Gorman et al., 2016; Frazier et al., 2019). Management of these conditions is complicated by their striking heterogeneity. They can develop acutely or progressively and can impact multiple organ systems or just a single cell type. Disease often manifests in post-mitotic tissues, such as skeletal muscle or the nervous system, but this pattern does not simply correlate with ATP demand or tissue expression of the causal gene. Deciphering how cells sense and respond to mitochondrial dysfunction is thus a major challenge with the potential to inform the common conditions that exhibit a decline in mitochondrial health.

Studies in patients, animals and cultured cells have consistently identified a gene expression program associated with the integrated stress response (ISR) as a signature of mitochondrial dysfunction (Martinus et al., 1996; Zhao et al., 2002; Kühl et al., 2017; Celardo et al., 2017; Quirós et al., 2017; Fujita et al., 2007; Silva et al., 2009; Tyynismaa et al., 2010; Dogan et al., 2014; Bao et al., 2016; Magarin et al., 2016; Lehtonen et al., 2016; Khan et al., 2017). The ISR is triggered by various insults, including nutrient deficiency, unfolded protein stress and pathogen infection. In mammalian cells, it is typically activated by a family of four kinases, each of which can phosphorylate the alpha subunit of translation initiation factor 2 (eIF2α) in response to distinct stimuli. eIF2α phosphorylation acutely inhibits global protein synthesis but promotes translation of transcription factors, such as activating transcription factor 4 (ATF4) and DNA damage-inducible transcript three protein (DDIT3), that then engage their downstream targets (Harding et al., 2003; Palam et al., 2011; Harding et al., 2000; Vattem and Wek, 2004; Wek, 2018; Pakos‐Zebrucka et al., 2016; Taniuchi et al., 2016).

The transcriptional program induced by the ISR varies across species and cell types and depends on the underlying trigger. Nevertheless, it frequently encompasses amino acid transport and biosynthesis genes, cytosolic tRNA synthetases and translation factors, pro-apoptotic factors, and genes involved in antioxidant defense, proteostasis and organelle quality control (Harding et al., 2003; Han et al., 2013). In tissues affected by mitochondrial disease, the ISR activates the mitochondrial 1-carbon pathway (Kühl et al., 2017; Tyynismaa et al., 2010; Bao et al., 2016; Nikkanen et al., 2016) and is thought to underlie secretion of the circulating cytokines fibroblast growth factor 21 (FGF21) and growth/differentiation factor 15 (GDF15) that are under consideration as disease biomarkers (Lehtonen et al., 2016; Fujita et al., 2015; Yatsuga et al., 2015; Chung et al., 2017; Miyake et al., 2016; Restelli et al., 2018). It remains unclear whether the ISR plays a protective role in mitochondrial disease or rather contributes to pathology (Khan et al., 2017; Suomalainen and Battersby, 2018; Lamech and Haynes, 2015).

How mitochondrial dysfunction is sensed to trigger the ISR is a major open question. Defects in mtDNA maintenance and expression are most frequently associated with ISR activation in vivo (Kühl et al., 2017; Lehtonen et al., 2016). However, such defects are expected to impose multiple biochemical consequences on the ETC/OXPHOS system, which are difficult to disentangle. To address this challenge, we profiled global gene expression, bioenergetics and metabolism in muscle cells that were proliferating (myoblasts) or differentiated (myotubes) and acutely treated with a panel of small-molecule mitochondrial inhibitors. To isolate the specific biochemical pathways that trigger the ISR, we employed a suite of ‘protein prostheses’ that facilitate NADH recycling to NAD⁺, in the cytosol or in mitochondria, independent of the ETC (Titov et al., 2016).

Our systematic analysis reveals that multiple mechanisms can link mitochondrial dysfunction to ISR activation. In proliferating myoblasts, we show that ETC inhibition elevates the mitochondrial and cytosolic NADH/NAD⁺ ratios, hindering aspartate synthesis and ultimately depleting asparagine, which activates the ISR via the eIF2α kinase GCN2. In myotubes, we find that inhibition of ATP synthase activates the ISR via a distinct mechanism related to mitochondrial inner-membrane hyperpolarization. Our results reject the notion of a universal trigger of the ISR in mitochondrial dysfunction, and rather, reveal multiple paths to its activation that depend both on the nature of the mitochondrial defect and on the metabolic state of the cell.

We sought to interrogate the genomic response to mitochondrial dysfunction and its dependence on the metabolic state of the cell. To this end, we utilized proliferating C2C12 mouse myoblasts, which can be induced to exit the cell cycle and then terminally differentiate into myotubes (Yaffe and Saxel, 1977; Blau et al., 1985; Figure 1A). Differentiation triggers mitochondrial biogenesis and promotes oxidative metabolism as the cells shift from a primary focus on nutrient uptake and biosynthesis to self-maintenance and specialized functions (Moyes et al., 1997; Leary et al., 1998; Casadei et al., 2009; Remels et al., 2010; Sin et al., 2016; Vander Heiden et al., 2009; Lemons et al., 2010).

Figure 1

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We acutely treated myoblasts and myotubes with inhibitors of complex I (piericidin), complex III (antimycin) or ATP synthase (oligomycin) (Figure 1B) in media lacking pyruvate and uridine, to avoid masking known metabolic vulnerabilities of ETC-compromised cells (Morais et al., 1980; King and Attardi, 1989) (Materials and methods). We confirmed the inhibitors exerted the expected effects on respiration (Figure 1C). Complex I or complex III inhibition abrogated mitochondrial oxygen consumption while cells treated with the ATP synthase inhibitor retained residual respiration, not coupled to ATP synthesis, often termed oligomycin-resistant respiration or leak respiration (Jastroch et al., 2010; Divakaruni and Brand, 2011). Myotube mitochondria showed less leak respiration, indicative of tighter coupling (Sin et al., 2016).

We next performed RNA-sequencing to profile global gene expression changes following 10 hr of treatment, roughly corresponding to the duration of the myoblast cell cycle. Principal component analysis (PCA) of the combined data from myoblasts and myotubes suggested marked differences in their response to mitochondrial inhibition but the strongest trend was the fundamental distinction between the cell states (Figure 1D). We therefore proceeded to analyze the results separately for myoblasts and for myotubes.

Mechanistically distinct inhibitors of mitochondria trigger the ISR in myoblasts

PCA of myoblast gene expression revealed a dominant first principal component (PC1; 57% of the variance) along which inhibited cells progressed, with piericidin and antimycin exerting a more pronounced effect than oligomycin (Figure 2A; Supplementary file 1). To gain insight into genes driving variation along PC1, we subjected the 500 genes with the most positive weights (upregulated along PC1) and, separately, the 500 genes with the most negative weights (downregulated) to cis-regulatory analysis (Janky et al., 2014) (Materials and methods). We also performed gene set enrichment analysis (GSEA) (Mootha et al., 2003; Subramanian et al., 2005) of REACTOME pathways (Croft et al., 2014) using PC1 weights as the gene ranks (Materials and methods).

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Gene set enrichment analysis.

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Both approaches converged on activation of the ISR as a major trend driving gene expression following inhibitor treatments. Cis-regulatory analysis identified binding motifs for the ISR master regulator ATF4 as the most enriched in promoters of genes upregulated along PC1 (Figure 2B; Supplementary file 2), while GSEA highlighted pathways typically upregulated as part of the ISR, including cytosolic tRNA synthetases and translation factors, amino acid transport and biosynthesis genes, and additional ISR regulators (Harding et al., 2003; Han et al., 2013; Figure 2C).

At the same time, the analyses revealed downregulation of genes linked to cell proliferation. Promoters of downregulated genes were enriched for motifs or ChIP-seq peaks of E2F-family transcription factors, master regulators of the cell cycle (Johnson et al., 1993; Helin, 1998; Ren et al., 2002; Fischer and Müller, 2017), and of SREBP-1/2, regulators of cholesterol and lipid metabolism (Yokoyama et al., 1993; Hua et al., 1993; Wang et al., 1994; Horton et al., 2002; Figure 2B; Supplementary file 2). The cell cycle, DNA replication and cholesterol biosynthesis pathways also emerged as the most sharply depressed in the GSEA results (Figure 2C). This is consistent with the proliferative defect ETC inhibition is expected to impose in our media conditions.

We generated a volcano plot relating each gene’s magnitude and significance of association with PC1 (Materials and methods). We then annotated it with verified target genes of ATF4 and DDIT3 derived from ChIP-seq data (Han et al., 2013), and with members of the REACTOME cell cycle and cholesterol pathways (Figure 2D; Supplementary file 1). The result demonstrated the striking enrichment of these categories among differentially expressed genes. Expression changes of top genes associated with PC1 from the enriched categories further illustrated that the transcriptional signatures of ISR activation (Figure 2E) and impaired proliferation (Figure 2F) mirrored one another (Hamanaka et al., 2005).

A large fraction of ISR target genes displayed the gradation of induction with the different inhibitors reflected along PC1, that is antimycin > piericidin > oligomycin (Group 1 in Figure 2—figure supplement 1A). Other targets, however, showed comparable induction across the inhibitors (Group 2) and a minority even showed the reverse gradation (Group 3). These patterns suggest complex cross-regulation of some ISR targets and underscore the non-equivalence of different modes of mitochondrial dysfunction. Nevertheless, the transcriptional signatures we observed were accompanied in all cases by eIF2α phosphorylation and ATF4 protein accumulation (Figure 2—figure supplement 1B).

Oxidizing cytosolic NADH/NAD⁺ is sufficient to ablate ISR activation by complex I inhibition in myoblasts

In myoblasts, complex I or complex III inhibition triggered the ISR more potently than ATP synthase inhibition. We therefore reasoned that a decrease in ATP levels due to breakdown of OXPHOS was less likely to play the operative role in ISR activation, whereas impaired NADH oxidation (‘reductive stress’) presented a more compelling candidate. To explore this possibility, we generated myoblasts expressing LbNOX, a bacterial water-forming NADH oxidase that directly converts NADH to NAD⁺ independent of the ETC (Titov et al., 2016; Figure 3A, Figure 3—figure supplement 1A). We also generated cells expressing mitoLbNOX, a variant of LbNOX targeted to the mitochondrial matrix (Titov et al., 2016; Figure 3A, Figure 3—figure supplement 1A). Finally, we generated cells expressing NDI1, a piericidin-resistant yeast protein that substitutes for complex I in transferring electrons from NADH to coenzyme Q (CoQ) and thus facilitates both continued NADH oxidation and ATP synthesis by OXPHOS (Titov et al., 2016; De Vries et al., 1992; Seo et al., 1998; Seo et al., 2000; Figure 3B, Figure 3—figure supplement 1A).

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We confirmed that complex I or complex III inhibition elevated whole-cell NADH/NAD⁺, which primarily reflects the mitochondrial ratio (Titov et al., 2016; Sies, 1982; Eng et al., 1989), more so than ATP synthase inhibition (Figure 3C), consistent with leak respiration in the latter case. The increase in whole-cell NADH/NAD⁺ was prevented in all cases by mitoLbNOX, as expected, whereas LbNOX provided only limited relief given the smaller cytosolic contribution to the whole-cell signal.

We also measured the secreted [lactate]/[pyruvate] ratio, a proxy for cytosolic NADH/NAD⁺ through equilibration by the lactate dehydrogenase (LDH) reaction (Williamson et al., 1967; Figure 3D). The results aligned with the predicted effects of the inhibitors on the malate-aspartate shuttle and the glycerol-3-phosphate shuttle, which facilitate oxidation of cytosolic reducing equivalents in mitochondria (Dawson, 1979; Safer et al., 1971). Thus, the increase in mitochondrial NADH/NAD⁺ in either piericidin- or antimycin-treated cells is expected to stall reactions coupled to this ratio in the tricarboxylic acid (TCA) cycle that are required for malate-aspartate shuttle activity (LaNoue and Williamson, 1971). However, only antimycin overly-reduces the CoQ pool, which blocks the TCA cycle at complex II independently of NADH/NAD⁺ and additionally inhibits the glycerol-3-phosphate shuttle (Mráček et al., 2013; Figure 3—figure supplement 1B).

LbNOX expression was able to maintain a low [lactate]/[pyruvate] ratio in all cases, as it directly oxidizes the cytosol (Figure 3D). mitoLbNOX also buffered the impact of piericidin and oligomycin but failed to effectively oxidize the cytosol during antimycin treatment. This result demonstrated that mitoLbNOX affected cytosolic NADH/NAD⁺ through the redox shuttles, both of which are inhibited by antimycin’s effect on CoQ in a manner that mitoLbNOX cannot resolve.

Strikingly, LbNOX expression was sufficient to almost completely ablate ISR activation, represented by the Ddit3 transcript, upon complex I or complex III inhibition but only partially mitigated ISR activation by ATP synthase inhibition (Figure 3E). mitoLbNOX also significantly attenuated ISR activation during complex I inhibition but proved less effective during complex III inhibition, possibly due to its failure to fully oxidize the cytosol in this case.

We performed RNA-sequencing to ascertain the global effects of maintaining compartment-specific NADH oxidation during inhibitor treatments. PCA of the combined data from control myoblasts (shown in Figure 2) and myoblasts expressing LbNOX, mitoLbNOX or NDI1 recapitulated a first principal component driven by the ISR and proliferative gene expression. The effects following inhibitor treatments were broadly in line with the results for Ddit3. During complex I inhibition (red), oxidizing the cytosol with LbNOX (square), or the matrix and the cytosol with mitoLbNOX (triangle), was almost as effective in preventing gene expression changes along PC1 as fully maintaining OXPHOS with NDI1 (diamond) (Figure 3F; Supplementary file 1). Thus, impaired NADH oxidation was the primary trigger of ISR-related gene expression in myoblasts and correcting the NADH imbalance in the cytosol was sufficient to prevent this, which was also evident following 48 hr of inhibiting mitochondrial protein synthesis with chloramphenicol (Figure 3G).

Complex I inhibition in myoblasts activates the eIF2α kinase GCN2 due to an asparagine deficiency

We focused our mechanistic investigation on the functional consequence of complex I inhibition that triggered the ISR, as the response was most effectively blunted by oxidizing cytosolic NADH in this case. One possibility was that glycolysis was inhibited by elevated cytosolic NADH/NAD⁺, limiting the cells’ ability to defend their adenylate energy charge in the absence of OXPHOS. Using lactate secretion as a proxy for glycolytic flux, we found that LbNOX did modestly stimulate glycolysis (Figure 4A, Figure 4—figure supplement 1A). However, this did not translate into an improved energy charge, and in fact, the opposite was true (Figure 4B, Figure 4—figure supplement 1B).

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Metabolite profiling data.

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Another possibility was that elevated mitochondrial and cytosolic NADH/NAD⁺ depleted a critical nutrient, which then triggered the ISR. We performed intracellular metabolite profiling on control cells or LbNOX cells acutely treated with piericidin (Figure 4C, Figure 4—figure supplement 1C). The amino acids aspartate and its derivative asparagine, alongside related TCA cycle intermediates, emerged among the top metabolites depleted by piericidin in a manner responsive to oxidizing the cytosol. This result is consistent with the contribution to aspartate synthesis of the mitochondrial and cytosolic NAD⁺-linked malate dehydrogenase (MDH) reactions that yield its precursor, oxaloacetate (Birsoy et al., 2015; Sullivan et al., 2015; Chen et al., 2016). Absolute quantification confirmed a ~ 16 fold drop in aspartate within 1 hr of piericidin treatment in control cells compared with only a 2.5-fold drop in LbNOX cells (Figure 4D, Figure 4—figure supplement 1D).

Amino acid deficiency is a canonical ISR trigger through the eIF2α kinase GCN2, which is activated due to uncharged tRNA (Hinnebusch, 1984; Dever et al., 1992; Wek et al., 1995; Berlanga et al., 1999; Zhang et al., 2002; Castilho et al., 2014; Inglis et al., 2019; Harding et al., 2019). If amino acid deficiency triggers the ISR following complex I inhibition in myoblasts, then correcting the deficiency should abrogate the response. We began by adding aspartate since the magnitude of its depletion was greater and its addition also allows the cells to partially restore asparagine (Figure 4E, Figure 4—figure supplement 1E). Indeed, 10 mM aspartate significantly attenuated ISR activation upon complex I inhibition, though not to the same extent as LbNOX (Figure 4F, Figure 4—figure supplement 1F–I). This concentration was required since most mammalian cells do not efficiently take up aspartate (Birsoy et al., 2015). Importantly, aspartate addition did not indirectly oxidize cytosolic NADH/NAD⁺ (Figure 4G) and was insufficient to appreciably restore cell proliferation (Figure 4H). Aspartate also attenuated ISR-related gene expression in C2C12 myoblasts engineered with a genetic defect in complex I (Vafai et al., 2016), where reductive stress was induced by pyruvate withdrawal (Figure 4I), as well as in piericidin-treated primary human myoblasts (Figure 4J) and primary mouse embryonic fibroblasts (Figure 4—figure supplement 1J).

To test whether ISR activation by complex I inhibition involved GCN2 activity, we used a recently described specific, small-molecule, ATP-competitive inhibitor of the kinase (GCN2iB) (Nakamura et al., 2018). Indeed, co-treatment with GCN2iB significantly suppressed ISR gene expression in response to complex I inhibition but had no effect on ISR activation in response to endoplasmic reticulum (ER) stress induced by tunicamycin (Figure 4K). The converse was true when we inhibited the ER-resident eIF2α kinase, PERK (Figure 4—figure supplement 1K; Harding et al., 1999; Axten et al., 2013).

Finally, we wished to examine how GCN2 sensed the amino acid deficiency. Using autophosphorylation at Threonine-898 as a readout of kinase activation (Romano et al., 1998), we observed that 6 hr of complex I inhibition resulted in marked GCN2 activation, which was partially attenuated by supplementing aspartate (Figure 4L). Strikingly, asparagine alone abrogated GCN2 activation with no discernible additive effect of aspartate, suggesting the drop in asparagine was the most proximal activating signal. As expected, LbNOX expression, which enables synthesis of both amino acids during complex I inhibition, effectively prevented GCN2 activation, while GCN2iB suppressed autophosphorylation both at baseline and upon piericidin treatment.

The effects of the same interventions on eIF2α phosphorylation, the event downstream of GCN2 activation, were notably less pronounced, especially given the narrow dynamic range of this signal. We observed clear attenuation of eIF2α phosphorylation upon piericidin treatment in the case of LbNOX expression, though the rescue was still incomplete (Figure 4L, Figure 4—figure supplement 1L). ATF4 protein levels largely tracked the degree of GCN2 activation (Figure 4L) and this was reflected at the level of transcriptional ISR targets at the same time-point (Figure 4M). GCN2iB completely prevented ATF4 protein accumulation despite some residual eIF2α phosphorylation (Figure 4L), suggesting GCN2 was required to attain the threshold phosphorylation level that elicits ATF4 translation. Collectively, these experiments indicate that in myoblasts, the rise in the NADH/NAD⁺ ratio is a major driver of ISR-related gene expression following complex I inhibition, as it limits biosynthesis of aspartate, depletes asparagine and activates GCN2 (Figure 4N).

ATP synthase inhibition potently triggers the ISR in myotubes while ETC inhibition neither depletes asparagine nor activates the response

We next examined the effects of inhibitor treatments in myotubes. In contrast to myoblasts, myotubes only experienced significant gene expression changes upon ATP synthase inhibition (PC1; 50% of the variance) (Figure 5A; Supplementary file 1), driven once more by ISR activation (Figure 5B), whereas ETC inhibition had little effect. The shift in the pattern of ISR activation was already evident as soon as 24 hr after the switch to low serum, when cells are largely post-mitotic but far from differentiated (Figure 5—figure supplement 1A).

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To interrogate why myotubes did not trigger the ISR upon ETC inhibition, we again measured compartment-specific NADH/NAD⁺. Myotubes did experience a significant increase in both the mitochondrial and the cytosolic ratio during ETC inhibition (Figure 5C,D). However, the ratios were not tightly linked, as in myoblasts. The severity of NADH imbalance across the inhibitors was discordant between the compartments and oxidizing mitochondrial NADH with mitoLbNOX failed to oxidize cytosolic NADH/NAD⁺. These results suggest myotubes did not rely on complex I to oxidize cytosolic reducing equivalents via the malate-aspartate shuttle. Rather, cytosolic NADH/NAD⁺ likely increased due to activation of glycolysis, evidenced by a ~ 75% rise in lactate secretion (Figure 5E), that was triggered by the drop in the energy charge regardless of the specific inhibitor used (Figure 5F). The glycerol-3-phosphate shuttle, which is uniquely sensitive to antimycin, remained operative (Figure 5D).

We next wondered how the altered metabolic state of myotubes impacted the fate of the amino acids aspartate and asparagine, whose depletion triggered the ISR in myoblasts. Despite elevated mitochondrial and cytosolic NADH/NAD⁺, complex I or complex III inhibition caused only a modest decrease (2-fold and 4-fold, respectively) in intracellular aspartate whereas ATP synthase inhibition even led to a slight accumulation (Figure 5G). None of the treatments depleted asparagine, which in fact accumulated (Figure 5H), possibly reflecting elevated cytosolic aspartate as asparagine synthesis occurs in the cytosol. The absence of asparagine deficiency likely explains why myotubes do not activate the ISR when complex I or complex III is inhibited. Myotubes also failed to trigger the ISR after mitochondrial protein synthesis was blocked for 48 hr (Figure 5—figure supplement 1B).

Several recent studies have reported activation of the integrated stress response (ISR) during mitochondrial dysfunction, however, tracing the underlying mechanisms has proved challenging. Here, we interrogated this response in two cell states, proliferating myoblasts and differentiated myotubes, by employing a battery of small-molecule inhibitors of mitochondria in combination with genetic and chemical tools for selectively buffering their effects. A chief result of this work is that there is no universal mechanism linking mitochondrial dysfunction to ISR activation but, rather, manifold paths that depend on the nature of the mitochondrial defect and on the metabolic state of the cell.

Our results in proliferating myoblasts mechanistically relate two recently recognized consequences of ETC dysfunction: a defect in aspartate biosynthesis and ISR activation. Aspartate is largely produced from TCA cycle-derived oxaloacetate (OAA) but elevated mitochondrial NADH/NAD⁺ blocks the oxidative TCA cycle when complex I is inhibited (Chen et al., 2016). OAA must then be obtained by other means, which in myoblasts depended on correcting the concomitant rise in cytosolic NADH/NAD⁺, thus tilting the MDH1 reaction toward OAA (Birsoy et al., 2015; Sullivan et al., 2015). Absent this, we found the aspartate deficiency triggered the ISR, at least initially due to depletion of its derivative asparagine that was sensed by the eIF2α kinase GCN2 (Figure 4C–M). The cytosolic aspartate and asparagine biosynthesis enzymes are both ISR targets (Fujita et al., 2007; Harding et al., 2003; Han et al., 2013), suggesting a homeostatic logic to the response.

Supplementation with asparagine alone, which cannot be converted to aspartate in most mammalian cells (Pavlova et al., 2018), was sufficient to abrogate GCN2 activation (Figure 4L). That GCN2 did not directly sense the profound aspartate depletion seems surprising. This result may be explained by the fact that while the K_M values for aspartate and asparagine of their respective cytosolic tRNA synthetases are similar (~15–30 μM) (Bour et al., 2009; Messmer et al., 2009; Andrulis et al., 1978), the cellular concentrations of these amino acids are orders of magnitude apart. Aspartate is estimated at ~5–10 mM whereas asparagine is much closer to the K_M, at ~100–200 μM (Chen et al., 2016; Park et al., 2016). Thus, even a large fold-change drop in aspartate would not compromise its cognate tRNA charging but a smaller drop in asparagine would, activating GCN2. The K_M for aspartate of asparagine synthetase is much higher (~1 mM) (Horowitz and Meister, 1972; Ciustea et al., 2005), so aspartate deficiency readily depletes asparagine (Figure 4E). We conclude asparagine tRNA charging is a sensitive sensor of NADH reductive stress in proliferating cells.

We note that oxidizing cytosolic NADH/NAD⁺ using LbNOX attenuated the ISR better than aspartate or asparagine (Figure 4F,L,M), suggesting eIF2α kinases may be attuned to additional consequences of reductive stress. Moreover, LbNOX cells still showed residual ISR activation (Figure 4L, Figure 4—figure supplement 1L), likely due to reasons beyond reductive stress. However, our results suggest threshold and non-linear effects along the ISR pathway so that NADH reductive stress was the primary driver of ISR-related gene expression.

Surprisingly, we also found that NADH reductive stress was a key modulator of an interaction between the ISR and p53 during complex III inhibition in myoblasts. The pyrimidine deficiency resulting from an overly-reduced CoQ triggered p53 to shut down ISR gene expression only when cytosolic reductive stress was resolved (Figure 3G). It is likely that the break on DNA replication imposed by reductive stress prevented the pyrimidine deficiency from giving rise to DNA damage, so p53 was not activated. Alternatively, cytosolic NADH/NAD⁺ may regulate p53 itself, possibly via NAD(P)H:quinone oxidoreductase 1 (NQO1) (Khutornenko et al., 2010; Asher et al., 2001; Asher et al., 2002), which could have implications for cancer biology.

The mapping between mitochondrial perturbation and ISR activation changed in non-dividing cells (Figure 1D, Figure 5B, Figure 5—figure supplement 1A). The ISR was not triggered by ETC inhibition in myotubes (Figure 5B), in line with a much smaller drop in aspartate and no drop in asparagine (Figure 5G,H). This result could be explained by reduced demand for these amino acids for protein and nucleotide biosynthesis (Clissold and Cole, 1973; Krauter et al., 1979; Bowman, 1987; Larson et al., 1993; Zahradka et al., 1991; Frangini et al., 2013; Talvas et al., 2006), and by increased capacity for amino acid turnover or scavenging (Deval et al., 2008; Sadiq et al., 2007), in myotubes relative to myoblasts. Moreover, myoblasts relied on complex I to oxidize cytosolic reducing equivalents via the malate-aspartate shuttle whereas this was absent in myotubes (Figure 3D, Figure 5D), consistent with observations in skeletal muscle in vivo (Sahlin et al., 1987; MacDonald and Marshall, 2000; Alfadda et al., 2004). Myotubes did exhibit elevated cytosolic NADH/NAD⁺ upon complex I inhibition but this was likely due to activation of glycolysis (Figure 5E). Conversion of glucose to lactate is net redox neutral at steady-state but elevated NADH/NAD⁺ can emerge when flux through glyceraldehyde 3-phosphate dehydrogenase (GAPDH) acutely rises or through siphoning of intermediates between GAPDH and LDH (Noor et al., 2013; Shestov et al., 2014).

Myotubes activated the ISR upon ATP synthase inhibition (Figure 5B), independently of a drop in ATP (Figure 5F) or impaired NADH oxidation (Figure 5C,D). The response was attenuated by eliminating residual respiration or by mild depolarization (Figure 5I, Figure 5—figure supplement 1C,E), implicating membrane potential in the signal relay. In respiring cells, oligomycin typically hyperpolarizes the inner-membrane as it blocks a major consumer of the proton gradient (Brand and Nicholls, 2011). Eliminating residual respiration relieves hyperpolarization by preventing proton pumping while depolarization involves facilitated proton re-entry into the matrix. This result is reminiscent of another case we recently described where inhibition of complex I ameliorated proliferative and metabolic defects due to ATP synthase inhibition, in part by relieving hyperpolarization (To et al., 2019). The link to hyperpolarization may explain why this route to the ISR is more prominent in myotubes, which exhibit increased and tightly coupled OXPHOS, though it likely plays a role in oligomycin-treated myoblasts as well. We note that severe depolarization also triggered the ISR in myotubes (Figure 5—figure supplement 1C).

While this paper was in revision, Guo et al. and Fessler et al. reported that the eIF2α kinase HRI potently triggers the ISR in response to either ATP synthase inhibition or severe depolarization (Fessler et al., 2020; Guo et al., 2020), in agreement with a previous study in the latter case (Taniuchi et al., 2016). HRI activation depended on processing of the inner-membrane protein DELE1 by the inner-membrane protease OMA1 (Fessler et al., 2020; Guo et al., 2020). It is tempting to speculate this mechanism is operative in our system, as OMA1 has been shown to respond to both depolarization and hyperpolarization (Ehses et al., 2009; Baker et al., 2014; Zhang et al., 2014), consistent with our observation that perturbation of membrane potential in either direction triggers the response. How such opposite effects converge in terms of signaling is unclear but our results suggest the common denominator is not, as Fessler et al. proposed, the drop in ATP (Figure 5F,I, Figure 5—figure supplement 1C,E).

Our results exclude mitochondrial processes that do not seem to be involved in ISR activation. The response was not activated simply due to the effects of OXPHOS inhibition on energy metabolism, so long as glucose was available. Similarly, we did not observe contributions to ISR activation by metabolic consequences exclusively related to elevated intra-mitochondrial NADH/NAD⁺. And blockade of mtDNA-encoded protein synthesis did not appear to be directly monitored in either cell state (Figure 3G, Figure 5—figure supplement 1B). The advantage of our cellular system is that so many parameters can be carefully controlled, however, it has important limitations: (i) chemical ETC inhibition is an imperfect model for genetic defects, which may impact multiple complexes and act over longer time scales, and (ii) cell culture conditions represent an artificial nutrient milieu, do not model interactions with other cell types and do not subject the cells to physiological workloads.

Our study demonstrates that multiple paths connect mitochondrial dysfunction to the ISR, depending on the combination of the defect and the metabolic state of the cell. This framework may prove useful in explaining variability in ISR activation across cell types and stressors and in dissecting the logic of the ISR in mitochondrial disorders. There is controversy over whether the ISR is activated in vivo directly by mtDNA expression defects or by ETC/OXPHOS defects (Kühl et al., 2017; Dogan et al., 2014; Lehtonen et al., 2016). Our results help address it by defining mechanisms through which the latter trigger the response. We note pyruvate therapy, which serves to regenerate NAD⁺, was recently shown to decrease serum GDF15, a circulating marker of the ISR, in mitochondrial disease patients (Koga et al., 2019), providing rich in vivo support for our identification of NADH imbalance as a trigger of the ISR. Further study of ISR activation and its consequences in the context of mitochondrial dysfunction may help define ways to therapeutically modulate the response.

Sequencing data have been deposited in GEO under accession code GSE132234.

1. 1. Mick E
   2. Titov DV
   3. Skinner OS
   4. Sharma R
   5. Jourdain AA
   6. Mootha VK

   (2020) NCBI Gene Expression Omnibus

   ID GSE132234. Distinct mitochondrial defects trigger the integrated stress response depending on the metabolic state of the cell.

   https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE132234

1. 1. Alcázar-Fabra M
   2. Navas P
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Author details

1. Eran Mick

   1. Howard Hughes Medical Institute and Department of Molecular Biology, Massachusetts General Hospital, Boston, United States
   2. Broad Institute, Cambridge, United States
   3. Department of Systems Biology, Harvard Medical School, Boston, United States

   Present address

   1. Division of Infectious Diseases & Division of Pulmonary and Critical Care Medicine, Department of Medicine, University of California, San Francisco, San Francisco, United States
   2. Chan Zuckerberg Biohub, San Francisco, United States

   Contribution

   Conceptualization, Data curation, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing - original draft, Project administration, Writing - review and editing

   Competing interests

   listed as an inventor on a patent filed by Massachusetts General Hospital on the therapeutic uses of GCN2 inhibitors.

   "This ORCID iD identifies the author of this article:" 0000-0002-7299-808X

2. Denis V Titov

   1. Howard Hughes Medical Institute and Department of Molecular Biology, Massachusetts General Hospital, Boston, United States
   2. Broad Institute, Cambridge, United States
   3. Department of Systems Biology, Harvard Medical School, Boston, United States

   Present address

   Department of Nutritional Science and Toxicology, Department of Molecular and Cell Biology & Center for Computational Biology, University of California, Berkeley, United States

   Contribution

   Conceptualization, Resources, Methodology

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0001-5677-0651

3. Owen S Skinner

   1. Howard Hughes Medical Institute and Department of Molecular Biology, Massachusetts General Hospital, Boston, United States
   2. Broad Institute, Cambridge, United States
   3. Department of Systems Biology, Harvard Medical School, Boston, United States

   Contribution

   Resources, Data curation, Methodology

   Competing interests

   a paid consultant for Proteinaceous Inc.

   "This ORCID iD identifies the author of this article:" 0000-0002-5023-0029

4. Rohit Sharma

   1. Howard Hughes Medical Institute and Department of Molecular Biology, Massachusetts General Hospital, Boston, United States
   2. Broad Institute, Cambridge, United States
   3. Department of Systems Biology, Harvard Medical School, Boston, United States

   Contribution

   Resources, Data curation, Methodology

   Competing interests

   No competing interests declared

5. Alexis A Jourdain

   1. Howard Hughes Medical Institute and Department of Molecular Biology, Massachusetts General Hospital, Boston, United States
   2. Broad Institute, Cambridge, United States
   3. Department of Systems Biology, Harvard Medical School, Boston, United States

   Contribution

   Resources, Data curation, Methodology

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0001-5321-6938

6. Vamsi K Mootha

   1. Howard Hughes Medical Institute and Department of Molecular Biology, Massachusetts General Hospital, Boston, United States
   2. Broad Institute, Cambridge, United States
   3. Department of Systems Biology, Harvard Medical School, Boston, United States

   Contribution

   Conceptualization, Supervision, Funding acquisition, Writing - original draft, Project administration, Writing - review and editing

   For correspondence

   vamsi@hms.harvard.edu

   Competing interests

   listed as an inventor on a patent filed by Massachusetts General Hospital on the therapeutic uses of GCN2 inhibitors, and a paid advisor to Janssen Pharmaceuticals and 5am Ventures.

   "This ORCID iD identifies the author of this article:" 0000-0001-9924-642X

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