We discovered that RNAi against hoe-1, a gene encoding a 3’-tRNA phosphodiesterase, attenuates hsp-6p::GFP induction—a fluorescence based transcriptional reporter of UPR^mt activation (Yoneda et al., 2004). Knockdown of hoe-1 by RNAi is sufficient to attenuate UPR^mt reporter activation induced by a loss-of-function mutation in the mitochondrial electron transport chain (ETC) complex I subunit NUO-6 (nuo-6(qm200)) (Figure 1A and B).

To further interrogate the potential role of hoe-1 in UPR^mt regulation, we used CRISPR/Cas9 to generate a hoe-1 null mutant (hoe-1(-/-)) by deleting the open reading frame of hoe-1 (Dokshin et al., 2018). The hoe-1 null mutants do not develop past late larval stage 3, thus the allele is maintained over a balancer chromosome, tmC25 (Dejima et al., 2018). UPR^mt induced by the knockdown of both the mitochondrial protease, spg-7, and ETC complex IV subunit, cco-1, is robustly attenuated in hoe-1 null animals (Figure 1C–E). Furthermore, UPR^mt induced by nuo-6(qm200) is attenuated in hoe-1 null animals similarly to what is seen in nuo-6(qm200) animals on hoe-1 RNAi (Figure 1F and G). Taken together, these findings suggest that HOE-1 is generally required for maximal UPR^mt activation.

To better understand the role of HOE-1 in UPR^mt regulation, we sought to identify where HOE-1 functions in the cell. HOE-1 is predicted to localize to both nuclei and mitochondria and this dual-localization has been shown for HOE-1 orthologs in Drosophila, mice, and human cell lines (Dubrovsky et al., 2004; Brzezniak et al., 2011; Rossmanith, 2011; Siira et al., 2018). To determine the subcellular localization of HOE-1 in C. elegans, we C-terminally tagged HOE-1 with GFP at its endogenous locus (HOE-1::GFP). Both hoe-1::GFP homozygous and hoe-1::GFP/hoe-1(-/-) trans-heterozygous animals grow and develop indistinguishably from wildtype animals suggesting that GFP-tagging HOE-1 does not compromise its essential functions (Figure 2—figure supplement 1A). We found that HOE-1 localizes to both mitochondria and nuclei (Figure 2A).

Given the dual-localization of HOE-1, we questioned whether it is mitochondrial or nuclear HOE-1 that is required for UPR^mt activation. To address this question, we created mitochondrial and nuclear compartment-specific loss-of-function mutants of HOE-1 (Figure 2B). hoe-1 contains two functional start codons. Translation beginning from the first start codon (encoding methionine 1 (M1)) produces a protein containing a mitochondrial targeting sequence (MTS). Translation beginning from the second start codon (encoding methionine 74 (M74)), which is 3’ to the MTS, produces a nuclear specific protein. This feature is conserved in human ELAC2 and it has been shown that mutating M1 to an alanine produces a mitochondrial-specific knockout (Brzezniak et al., 2011). Thus, we used the same approach to create a mitochondrial-specific knockout of C. elegans HOE-1 (hoe-1(ΔMTS)). This mutation is sufficient to strongly attenuate mitochondrial targeting without impacting nuclear localization (Figure 2—figure supplement 2A).

UPR^mt reporter activation by spg-7 and cco-1 RNAi is not attenuated in hoe-1(ΔMTS) animals (Figure 2C and D and Figure 2—figure supplement 3A and B). In fact, UPR^mt reporter activation is slightly elevated in hoe-1(ΔMTS) animals relative to wildtype. These data suggest that mitochondrial HOE-1 is not required for UPR^mt activation.

HOE-1 is predicted to contain two nuclear localization signals (NLS). Given that hoe-1 null mutant animals are developmentally arrested and hoe-1(ΔMTS) animals are superficially wildtype we reasoned that completely ablating nuclear localization of HOE-1 may result in recapitulation of the null phenotype. In effort to disentangle the developmental effects from the effect on UPR^mt we ablated only one of the nuclear localization signals of HOE-1. To compromise nuclear localization, we mutated the positively charged residues of the most N-terminal NLS to alanines (hoe-1(ΔNLS)). These mutations are sufficient to strongly attenuate, but not completely ablate, HOE-1 nuclear localization whilst still allowing animals to develop to adulthood (Figure 2—figure supplement 4A–C).

In contrast to loss of mitochondrial HOE-1, loss of nuclear HOE-1 robustly attenuates UPR^mt reporter activation induced by spg-7 RNAi (Figure 2E and F) and attenuates UPR^mt reporter activation induced by nuo-6(qm200) (Figure 2G and H). Furthermore, loss of nuclear HOE-1 attenuates the transcriptional upregulation of UPR^mt target genes hsp-6 (Figure 2I) and cyp-14A1.4 (Figure 2—figure supplement 5A) under conditions of mitochondrial stress. Together these data suggest that HOE-1 is required in the nucleus to facilitate UPR^mt activation.

Like many nuclear localized proteins (la Cour et al., 2004), HOE-1 has both nuclear localization signals and a nuclear export signal (NES). Given that loss of nuclear HOE-1 results in UPR^mt attenuation we questioned if compromising HOE-1 nuclear export, by ablating the NES of HOE-1, is sufficient to activate UPR^mt. We created a HOE-1 NES knockout mutant (hoe-1(ΔNES)) by replacing the strong hydrophobic residues of the predicted NES with alanines (Figure 3—figure supplement 1A). hoe-1(ΔNES) animals are superficially wildtype in their development but are sterile. Thus, the allele is balanced with tmC25. Homozygous hoe-1(ΔNES) animals have elevated nuclear HOE-1 accumulation relative to wildtype (Figure 3—figure supplement 1B, Figure 2—figure supplement 4B and C).

Strikingly, the UPR^mt reporter hsp-6p::GFP is robustly activated in adult hoe-1(ΔNES) animals similarly to that seen in mitochondrial stressor nuo-6(qm200) and constitutive UPR^mt activation in atfs-1 gain-of-function (atfs-1(et15)) mutant animals (Figure 3A and B). hoe-1(ΔNES) also mildly induces the less sensitive UPR^mt reporter hsp-60p::GFP (Figure 3—figure supplement 2A and B).

UPR^mt activation is characterized by the transcriptional upregulation of a suite of mitochondrial stress response genes that encode chaperone proteins, proteases, and detoxification enzymes that function to restore mitochondrial homeostasis (Nargund et al., 2012). To interrogate the extent of UPR^mt induction in hoe-1(ΔNES) animals, we measured transcript levels of a diverse set of UPR^mt associated genes. We found that the UPR^mt genes encoding a chaperone protein (hsp-6), stress response involved C-type lectin (clec-47), and P450 enzyme (cyp-14A4.1) are all upregulated in hoe-1(ΔNES) animals (Figure 3C, D and E). These data support hoe-1(ΔNES) being sufficient to activate the UPR^mt transcriptional response.

UPR^mt activation is dependent upon the transcription factor ATFS-1 (Haynes et al., 2010; Nargund et al., 2012). Thus, we tested if UPR^mt reporter activation in hoe-1(ΔNES) animals is ATFS-1 dependent. Knockdown of atfs-1 is sufficient to completely attenuate UPR^mt reporter activation in hoe-1(ΔNES) animals (Figure 3F and G), showing that UPR^mt induction by hoe-1(ΔNES) is ATFS-1 dependent.

To further interrogate how UPR^mt is activated in hoe-1(ΔNES) animals, we made double localization mutants of hoe-1. If UPR^mt is activated in hoe-1(ΔNES) animals due to elevated nuclear HOE-1 levels we reasoned that introducing a hoe-1(ΔNLS) mutation in the hoe-1(ΔNES) background (hoe-1(ΔNLS+ΔNES)) should be sufficient to attenuate UPR^mt activation. Indeed, hoe-1(ΔNLS+ΔNES) animals have UPR^mt reporter activation comparable to wildtype animals (Figure 3—figure supplement 3A and B). Furthermore, we reasoned that compromising mitochondrial localization of HOE-1 in a hoe-1(ΔNES) background (hoe-1(ΔMTS+ΔNES)) may further enhance hoe-1(ΔNES)-induced UPR^mt activation as what would be the mitochondrial targeted HOE-1 pool should be diverted to the nucleus as well. Consistent with our hypothesis, hoe-1(ΔMTS+ΔNES) animals have even higher activation of UPR^mt than hoe-1(ΔNES) alone (Figure 3—figure supplement 4A and B). Taken together, these data strongly suggest that hoe-1(ΔNES) triggers UPR^mt activation due to elevated nuclear HOE-1 levels.

Contrary to UPR^mt induced by nuo-6(qm200) and atfs-1(et15), hoe-1(ΔNES) animals appear to have UPR^mt activated specifically in the intestine (Figure 3A). We questioned if this UPR^mt activation is occurring cell autonomously or non-cell autonomously as UPR^mt has been shown to be able to be signaled across tissues, particularly from neurons to intestine (Durieux et al., 2011; Berendzen et al., 2016; Zhang et al., 2018). To determine which tissue HOE-1 is required in for UPR^mt activation we took advantage of the auxin-inducible degradation (AID) system that allows for tissue-specific protein degradation (Zhang et al., 2015). Briefly, degron-tagged proteins will be degraded in the presence of the plant hormone auxin but only in tissues wherein E3 ubiquitin ligase subunit, TIR1, is expressed. We C-terminally degron-tagged hoe-1(ΔNES) (hoe-1(ΔNES)::degron) and crossed this allele into backgrounds in which TIR1 is driven under an intestinal-specific (ges-1p::TIR1) or a neuronal-specific (rgef-1p::TIR) promoter (Ashley et al., 2021). hoe-1(ΔNES)-induced UPR^mt is only attenuated when HOE-1 is selectively degraded in the intestine (Figure 3—figure supplement 5A and B). This data strongly suggests that compromised nuclear export of HOE-1 activates UPR^mt cell-autonomously in the intestine.

Changes in protein synthesis rates and associated protein folding capacity can broadly activate cellular stress response mechanisms (Wang and Kaufman, 2016; Das et al., 2017; Boos et al., 2019). Given the role of hoe-1 in tRNA maturation, we questioned if the robust upregulation of UPR^mt in hoe-1(ΔNES) animals may be the result of compromised cellular proteostasis in general rather than specific activation of UPR^mt. One stress response mechanism that is sensitive to global proteotoxic stress is the endoplasmic reticulum unfolded protein response (UPR^ER) (Preissler and Ron, 2019). We find that the UPR^ER reporter hsp-4p::GFP is not activated in hoe-1(ΔNES) animals (Figure 3H and I), suggesting that hoe-1(ΔNES) does not cause ER stress nor cellular proteotoxic stress. Additionally, a basal reporter of GFP that has been used to proxy general protein expression (Gitschlag et al., 2016), ges-1p::GFPcyto, is only mildly upregulated in hoe-1(ΔNES) animals relative to wildtype (Figure 3J and K). Together these findings support that impaired nuclear export of HOE-1 specifically activates UPR^mt.

UPR^mt is known to be triggered when mitochondrial membrane potential is compromised (Rolland et al., 2019; Shpilka et al., 2021). Thus, we assessed mitochondrial membrane potential, using TMRE staining, in adult hoe-1(ΔNES) animals where UPR^mt is robustly activated. Consistent with UPR^mt activation, we found that mitochondrial membrane potential is severely depleted in adult hoe-1(ΔNES) animals (Figure 4A and B). However, hoe-1(ΔNLS) animals also exhibit compromised mitochondrial membrane potential without UPR^mt activation suggesting that decreased membrane potential does not guarantee UPR^mt induction (Figure 4A and B). Compromised mitochondrial membrane potential can be both a cause and consequence of UPR^mt activation (Rolland et al., 2019; Shpilka et al., 2021). Thus, we assessed whether or not compromised membrane potential in hoe-1(ΔNES) animals is atfs-1-dependent. Mitochondrial membrane potential is not rescued in hoe-1(ΔNES) animals on atfs-1 RNAi (Figure 4C and D) suggesting that reduced mitochondrial membrane potential in hoe-1(ΔNES) animals is not a result of UPR^mt activation. Taken together, these data show that compromised nuclear export of HOE-1 results in depletion of mitochondrial membrane potential. Furthermore, this depletion in membrane potential correlates with UPR^mt activation, consistent with the possibility that hoe-1(ΔNES) activates UPR^mt via depletion of mitochondrial membrane potential.

UPR^mt activation is dependent upon nuclear accumulation of the transcription factor ATFS-1 (Nargund et al., 2012; Nargund et al., 2015). Thus, we tested if ATFS-1 accumulates in nuclei of hoe-1(ΔNES) animals by assessing the fluorescence intensity of ectopically expressed mCherry-tagged ATFS-1 (atfs-1p::ATFS-1::mCherry) in wildtype, hoe-1(ΔNES), and mitochondrial-stressed nuo-6(qm200) animals. Both hoe-1(ΔNES) and nuo-6(qm200) animals have elevated nuclear accumulation of ATFS-1 relative to wildtype (Figure 5A and B). However, while nuo-6(qm200) animals exhibit elevated levels of total cellular and extranuclear levels of ATFS-1::mCherry relative to wildtype, hoe-1(ΔNES) animals do not (Figure 5C and Figure 5—figure supplement 1A). We find that atfs-1 mRNA levels are also elevated in hoe-1(ΔNES) animals relative to wildtype comparable to that seen in nuo-6(qm200) animals (Figure 5D).

The transcription factor DVE-1 is required for full UPR^mt activation (Haynes et al., 2007; Tian et al., 2016). Thus, we asked if DVE-1::GFP nuclear expression is higher in hoe-1(ΔNES) than in wildtype animals. We found that accumulation of DVE-1::GFP in intestinal cell nuclei is significantly higher in hoe-1(ΔNES) than in wildtype animals (Figure 5E and F). Qualitatively, cellular DVE-1::GFP levels appear mildly elevated in hoe-1(ΔNES) animals based on actin (Figure 5G, Figure 5—source data 1), though the difference in DVE-1::GFP levels is not significant when normalized to total protein (Figure 5H). Thus, while we cannot rule out the possibility of a slight increase in the cellular levels of DVE-1, elevation in the nuclear localization of DVE-1 in hoe-1(ΔNES) animals is the more robust phenotype. Together, these data suggest that UPR^mt induction in hoe-1(ΔNES) animals is a result of increased nuclear accumulation of UPR^mt transcription factors ATFS-1 and DVE-1.

The canonical role of HOE-1 is to cleave 3’-trailer sequences from nascent tRNAs (Nashimoto et al., 1999; Mayer et al., 2000; Schiffer et al., 2002; Takaku et al., 2003; Dubrovsky et al., 2004; Brzezniak et al., 2011; Sanchez et al., 2011; Siira et al., 2018). This enzymatic function is dependent upon zinc binding (Ma et al., 2017; Bienert et al., 2017). Thus, we queried if UPR^mt activation by hoe-1(ΔNES) is dependent upon the catalytic activity of HOE-1. To test this, we generated a catalytically-dead HOE-1 mutant by changing an essential aspartate of the zinc-binding pocket of HOE-1 to alanine in both a wildtype (hoe-1(D624A)) and hoe-1(ΔNES) (hoe-1(D624A+ΔNES)) background. Animals homozygous for D624A recapitulate the growth arrest phenotype of the hoe-1 null mutant precluding us from assessing the impact of D624A on UPR^mt induction in adult hoe-1(ΔNES) animals. To overcome this constraint, we assessed UPR^mt activation in hoe-1(ΔNES) versus hoe-1(ΔNES)/hoe-1(D624A+ΔNES) trans-heterozygous animals. A single copy of catalytically dead hoe-1 is sufficient to attenuate hoe-1(ΔNES)-induced UPR^mt (Figure 6—figure supplement 1A and B). These data suggest that hoe-1(ΔNES)-induced UPR^mt requires the catalytic activity of HOE-1.

Given that HOE-1 catalytic activity is required for UPR^mt, we further interrogated the potential role of tRNA processing as a mechanism by which HOE-1 may modulate UPR^mt induction. Production of mature tRNAs begins with transcription of tRNA gene loci by RNA polymerase III followed by sequential cleavage of 5’-leader and 3’-trailer sequences from immature tRNA transcripts by RNase P and HOE-1, respectively. Following cleavage of 3’-trailer sequences, tRNAs can be transported to the cytosol by tRNA exportin (Hopper and Nostramo, 2019).

Given that HOE-1 nuclear levels are elevated in hoe-1(ΔNES) animals, we reasoned that 3’-tRNA processing should be elevated due to increased nuclear activity of HOE-1. Thus, we questioned if UPR^mt induction in hoe-1(ΔNES) animals is a result of elevated 3’-tRNA processing. First, we knocked-down RNA pol III subunit rpc-1 to attenuate the production of total RNA pol III-dependent transcripts in hoe-1(ΔNES) animals. If hoe-1(ΔNES)-induced UPR^mt is due to elevated processing of tRNAs we hypothesized that restriction of nascent tRNA production should attenuate UPR^mt activation. Indeed, we found that rpc-1 RNAi robustly attenuates hoe-1(ΔNES)-induced UPR^mt (Figure 6—figure supplement 2A and B). Interestingly, rpc-1 RNAi has little impact on mitochondrial stress-induced UPR^mt (nuo-6(qm200)) (Figure 6—figure supplement 2C and D). These data show that rpc-1 is required for hoe-1(ΔNES)-induced UPR^mt and support our hypothesis that increased 3’-tRNA processing by HOE-1 activates UPR^mt.

For the majority of tRNAs 5’-end processing by the RNase P complex is a prerequisite for 3’-end processing by HOE-1 (Frendewey et al., 1985; Yoo and Wolin, 1997). Thus, if increased 3’-tRNA end processing is responsible for UPR^mt activation, compromising 5’-end processing by RNAi against RNAse P should attenuate hoe-1(ΔNES)-induced UPR^mt. RNAi against a subunit of the RNase P complex, popl-1, attenuates UPR^mt induction in hoe-1(ΔNES) animals (Figure 6A and B). popl-1 RNAi also attenuates both UPR^mt induced by nuo-6(qm200) (Figure 6C and D) as well as basal induction of ges-1p::GFPcyto (Figure 6E and F), albeit to a lesser extent than the attenuation seen in hoe-1(ΔNES) animals. These data suggest that popl-1 RNAi may have a broad impact on protein expression but supports that elevated 3’-tRNA processing in hoe-1(ΔNES) animals is responsible for UPR^mt activation given that popl-1 RNAi strongly attenuates hoe-1(ΔNES)-induced UPR^mt.

Following 3’-end processing in the nuclei, tRNAs can be exported to the cytosol by tRNA exportin (Hopper and Nostramo, 2019). To test if elevated levels of 3’-processed tRNAs are required in the cytosol to activate UPR^mt, we asked if restricting tRNA nuclear export via RNAi against tRNA exportin, xpo-3, attenuates hoe-1(ΔNES)-induced UPR^mt. Strikingly, xpo-3 RNAi robustly attenuates hoe-1(ΔNES)-induced UPR^mt(Figure 6G and H). However, xpo-3 RNAi does not attenuate nuo-6(qm200) induced UPR^mt (Figure 6I and J) nor basal ges-1p::GFP levels (Figure 6K and L). These data suggest that in hoe-1(ΔNES) animals 3’-processed tRNAs are required in the cytosol to activate UPR^mt.

While 5’- and 3’-tRNA processing are the only steps known to be required for tRNA export from the nucleus, there are other downstream tRNA maturation processes that occur (Hopper and Nostramo, 2019). Some nascent tRNAs include introns that need to be removed and then ligated by a tRNA ligase (Englert and Beier, 2005; Popow et al., 2012). For tRNAs to be charged with corresponding amino acids, nascent tRNAs must contain a CCA sequence as part of the 3’ acceptor stem. This can be achieved by a CCA-adding tRNA nucleotidyl transferase (Hou, 2010). Knockdown of both tRNA ligase, rtcb-1, and tRNA nucleotidyl transferase, hpo-31 mildly attenuate hoe-1(ΔNES)-induced UPR^mt (Figure 6—figure supplement 3A–C). However, rtcb-1 RNAi also mildly attenuates nuo-6(qm200)-induced UPR^mt (Figure 6—figure supplement 3D and E). Knockdown of hpo-31 severely compromised growth of nuo-6(qm200) animals and thus the impact on UPR^mt could not accurately be assessed. These data suggest that tRNA ligation and CCA addition have limited involvement in hoe-1(ΔNES)-induced UPR^mt.

Taken together, these data suggest that UPR^mt induction by nuclear export deficient HOE-1 is the result of increased 3’-tRNA processing and that these tRNA species are required in the cytosol to trigger UPR^mt.

Alteration to tRNA processing can activate cellular signaling pathways (Raina and Ibba, 2014). One such pathway is the integrated stress response (ISR) in which uncharged tRNAs activate the kinase GCN2 which, in turn, phosphorylates the eukaryotic translation initiation factor, eIF2α. This inhibitory phosphorylation of eIF2α leads to upregulation of a select number of proteins including the transcription factor ATF4 (Pakos-Zebrucka et al., 2016; Costa-Mattioli and Walter, 2020). Interestingly, ATF4 and one of its targets, ATF5, are orthologs of ATFS-1 (Fiorese et al., 2016). Moreover, GCN2 and ISR in general have been shown to be responsive to mitochondrial stress (Baker et al., 2012; Fessler et al., 2020; Guo et al., 2020; Koncha et al., 2021). Thus, we questioned if UPR^mt activation by hoe-1(ΔNES) is mediated via GCN2 and eIF2α phosphorylation. We found that hoe-1(ΔNES)-induced UPR^mt is only slightly reduced in both a gcn-2 null (gcn-2(ok871)) and an eIF2α non-phosphorylatable mutant (eIF2α(S46A,S49A)) background (Figure 7A and B). These data suggest that a mechanism independent of ISR must largely be responsible for UPR^mt activation by hoe-1(ΔNES) animals.

To better understand the potential physiological implications of HOE-1 in UPR^mt, we assessed hoe-1 expression and subcellular dynamics of HOE-1 during mitochondrial stress. It is predicted that two major transcripts are produced from the hoe-1 gene locus: one that contains an MTS and one that does not, which are translated into mitochondrial- and nuclear-targeted HOE-1, respectively. However, it has been shown in other systems that hoe-1 orthologs produce a single transcript that encodes both a mitochondrial targeted and nuclear targeted HOE-1 isoform via alternative translation initiation (Rossmanith, 2011). Thus, we first sought to determine which mechanism is used for hoe-1 expression. To do so, we designed two sets of primers complementary to hoe-1 mRNA one of which amplifies only mRNA that includes the MTS and the other which amplifies all hoe-1 mRNA (spans a sequence that is included in all predicted HOE-1 isoforms). If there are two independent hoe-1 transcripts, we expected there to be higher levels of hoe-1 mRNA measured by the primer pair for total transcripts than for the mitochondrial specific pair. However, we found that both primer pairs measured similar levels of hoe-1 mRNA (Figure 8—figure supplement 1A) suggesting that, like in other systems, there is a single hoe-1 transcript. Next, we assessed hoe-1 mRNA levels in non-stress versus mitochondrial stress conditions. We found, using both primer pairs, that hoe-1 mRNA levels are modestly elevated in nuo-6(qm200) animals relative to wildtype (Figure 8—figure supplement 1B and C) suggesting that hoe-1 may be transcriptionally upregulated under conditions of mitochondrial stress.

Next, we assessed the subcellular dynamics of HOE-1 in response to mitochondrial stress. We found that HOE-1::GFP nuclear levels are markedly diminished under mitochondrial stress induced by nuo-6(qm200), cco-1 RNAi, and spg-7 RNAi (Figure 8A and B and Figure 8—figure supplement 2A and B). This observation was unexpected given that hoe-1 transcript levels are elevated during mitochondrial stress and it runs contrary to the fact that compromising HOE-1 nuclear export is sufficient to induce UPR^mt (Figure 3A and B). A common feature of signaling pathways is negative regulation. Thus, we questioned if reduced nuclear HOE-1 is a result of negative feedback rather than a direct result of mitochondrial stress. Given that mitochondrial stress activates UPR^mt, we assessed HOE-1::GFP status in a mitochondrial stress background wherein atfs-1 is knocked down by RNAi. HOE-1 levels are significantly upregulated in nuclei of nuo-6(qm200) animals on atfs-1 RNAi relative to nuo-6(qm200) animals on control RNAi, as well as both wildtype animals on control and atfs-1 RNAi (Figure 8A and B and Figure 8—figure supplement 3A–C). Moreover, total cellular HOE-1 levels are elevated under mitochondrial stress in an atfs-1 RNAi background (Figure 8C and D and Figure 8—source data 1A–E and Figure 8—source data 2A–E). Additionally, mitochondrial HOE-1 levels are elevated under mitochondrial stress conditions irrespective of RNAi treatment (Figure 8—figure supplement 3D). Together these data suggest that HOE-1 is upregulated and accumulates in nuclei upon mitochondrial stress. Then, nuclear HOE-1 is negatively regulated by ATFS-1 once UPR^mt is activated.

To further test if nuclear HOE-1 is negatively regulated by UPR^mt activation rather than by mitochondrial stress, we assessed HOE-1 localization in ATFS-1 gain-of-function animals (atfs-1(et15)). atfs-1(et15) constitutively activates UPR^mt in the absence of mitochondrial stress (Rauthan et al., 2013). Thus, we asked if atfs-1(et15) is sufficient to reduce nuclear HOE-1 levels. Indeed, nuclear HOE-1 levels are markedly reduced in atfs-1(et15) animals relative to wildtype (Figure 8E and F and Figure 8—figure supplement 4A–C) while total and mitochondrial HOE-1 protein levels are largely unperturbed (Figure 8G and H, Figure 8—figure supplement 4D, Figure 8—source data 3A–E and Figure 8—source data 4A–E). These data further support that UPR^mt activation negatively regulates nuclear HOE-1.

Worms were grown on nematode growth media (NGM) seeded with OP50 E. coli bacteria and maintained at 20 °C.

A complete list of C. elegans strains used can be found in Supplementary file 1. All new mutant and transgenic strains generated via CRISPR/Cas9 for this study were confirmed by Sanger sequencing.

CRISPR was conducted as previously described (Dokshin et al. Genetics 2018; Paix et al. Genetics 2015) using Alt-R S.p. Cas9 Nuclease V3 (IDT #1081058) and tracrRNA (IDT #1072532). A complete list of crRNA and repair template sequences purchased from IDT can be found in Supplementary file 2.

Strains resulting from genetic crosses were generated by crossing ~20 heterozygous males of a given strain to 5–8 L4 hermaphrodites of another strain (heterozygous males were generated by first crossing L4 hermaphrodites of that strain to N2 males). F1, L4 hermaphrodites were then cloned out and allowed to have self-progeny. F2 progeny were cloned out and once they had progeny were genotyped or screened (if fluorescent marker) for presence of alleles of interest. All genotyping primers were purchased from IDT and can be found in Supplementary file 2.

All whole animal imaging was done using Zeiss Axio Zoom V16 stereo zoom microscope. For all whole animal imaging, worms were immobilized on 2% agar pads on microscope slides in ~1 μl of 100 mM levamisole (ThermoFisher #AC187870100) and then coverslip applied.

For whole animal fluorescence intensity quantification, total pixels (determined by tracing individual animals and summing the total number of pixels within the bounds of the trace) and pixel fluorescence intensity (pixel fluorescence intensity on 1–255 scale) were quantified using imageJ and mean fluorescence intensity for each worm was calculated (sum total of fluorescence intensity divided by total number of pixels within bounds of the trace). For DVE-1::GFP image analysis (Figure 5E&F), brightness threshold was set to 25 in imageJ and then the number of gut cell nuclei that were saturated at this threshold were counted. For Figure 8A&B and Figure 8E&F, and Figure 8—figure supplement 2A&B, mean fluorescence intensity was calculated within the bounds of gut cell nuclei and outside of the bounds of gut cell nuclei and then graphed as the ratio fluorescence intensity of nuclear to extranuclear signal.

RNAi by feeding was conducted as previously described (Gitschlag et al. Cell Met. 2016). Briefly, RNAi clones were grown overnight from single colony in 2 ml liquid culture of LB supplemented with 50 μg/ml ampicillin. To make 16 RNAi plates, 50 ml of LB supplemented with 50 μg/ml ampicillin was inoculated with 500 μl of overnight culture and then incubated while shaking at 37 °C for 4–5 hours (to an OD_550-600 of about 0.8). Cultures were then induced by adding 50 ml additional LB supplemented with 50 μg/ml ampicillin and 4 mM IPTG and then continued incubating while shaking at 37 °C for 4 hours. Following incubation, bacteria were pelleted by centrifugation at 3900 rpm for 6 min. Supernatant was decanted and pellets were gently resuspended in 4 ml of LB supplemented with 8 mM IPTG. 250 μl of resuspension was seeded onto standard NGM plates containing 1 mM IPTG. Plates were left to dry overnight and then used within 1 week. Bacterial RNAi feeder strains were all from Ahringer RNAi Feeding Library, grown from single colony and identity confirmed by Sanger sequencing. atfs-1 (ZC376.7), cco-1 (F26E4.9), hoe-1 (E04A4.4), hpo-31 (F55B12.4), popl-1 (C05D11.9), rpc-1 (C42D4.8), rtcb-1 (F16A11.2), spg-7 (Y47G6A.10), xpo-3 (C49H3.10).

cDNA was synthesized using Maxima H Minus First Strand cDNA Synthesis Kit, with dsDNase (ThermoFisher #K1682) according to manufacturer’s directions. Lysates for cDNA synthesis were made by transferring 10, day 2 adult worms to 10 μl of lysis buffer supplemented with 20 mg/ml proteinase K and incubating at 65 °C for 10 min, 85 °C for 1 min and 4 °C for 2 min. Quantification of gene expression was performed using droplet digital PCR (ddPCR) with Bio-Rad QX200 ddPCR EvaGreen Supermix (Bio-Rad #1864034). Primers used for ddPCR can be found in Supplementary file 2.

A total of 500 μl of 1 mM TMRE (ThermoFisher #T669) solution in M9 buffer (prepared from a stock TMRE solution of 0.5 M in DMSO) was supplemented on top of standard NGM plates pre-seeded with 200 ul lawn of OP50 and allowed to dry overnight in the dark. The following day young L4 animals were transferred to TMRE plates and incubated on TMRE for 16 hr. After 16 hr, animals were transferred from TMRE plates to seeded standard NGM plates for 1 hr to remove any non-specific TMRE signal from cuticle and intestinal lumen. Animals were then imaged via confocal microscopy as described below.

Worms were grown at 20 °C and age-synchronized by timed egg-lays on NGM plates seeded with OP50 or HT115 bacteria for RNAi experiments. Before imaging, worms were immobilized with 3 μl 0.05 µm Polybead microsphere suspension (Polysciences) on a 10% agarose pad with a coverslip (1). Images were taken in the mid- or posterior intestine using a Nikon Ti2 with CSU-W1 spinning disk and Plan-Apochromat 100 X/1.49 NA objective. HOE-1::GFP was imaged by 488 nm laser excitation and ET525/36 m emission filter. 2 X integration was applied (Nikon Elements) to increase signal strength. TMRE and ATFS-1::mCherry were imaged with 561 nm laser excitation and ET605/52 M emission filter.

Image processing and analysis was performed with Nikon Elements software. Raw images were subjected to deconvolution and rolling ball background subtraction. Mitochondrial networks were segmented using the TMRE signal after excluding dye aggregates via Bright Spot Detection. To objectively set threshold parameters across groups with different TMRE intensity levels, the low threshold for segmentation was calculated based on a linear correlation with mean TMRE intensity within each group, y = 0.6411*x + 89.71 (x = mean TMRE intensity and constants derived from an initial manual validation). Regions of interest (ROIs) were manually drawn to encompass a single intestinal cell, and nuclei were identified and segmented manually using brightfield images. Mean intensities were measured within the resulting masks.

To detect localization of HOE-1::GFP in mitochondria, images of TMRE-stained intestinal cells of control and ΔMTS worms were collected and blinded. Mitochondria were segmented by TMRE signals as above. For each cell, one representative line scan was drawn manually across the mitochondrial short axis.

Fifty adult worms were transferred into a tube containing 20 μl of M9 Buffer. Then, 20 μl of 2 x Laemmli Buffer (BioRad #161–0737) supplemented with 2-mercaptoethanol (i.e. βME) was added to worm suspension and gently pipetted up and down 5 times to mix. Worms were lysed at 95 °C for 10 min in thermocycler followed by ramp down to room temperature (25 °C). Lysates were then pipetted up and down 10 times to complete disrupt and homogenize suspension. Samples were briefly centrifuged to pellet any worm debris. 20 μl of lysate supernatant was loaded onto precast Mini-PROTEAN TGX Stain-Free Gel (BioRad #4568045). Gel was run for 30 min at 100 V and then an additional 40–45 min at 130 V in 1 x Tris/Glycine/SDS Running Buffer (BioRad #1610732). Following electrophoresis gel was activated and imaged for total protein. Gel was equilibrated in Trans-Blot Turbo Transfer Buffer (BioRad #10026938) and transferred to activated and equilibrated Trans-Blot Turbo LF PVDF Membrane (BioRad #10026934) for 7 min at 2.5 A/25 V on Trans-Blot Turbo Transfer System. Following transfer, stain-free membrane was imaged for total protein. Membrane was then blocked in 5% milk in TBST for 2 hr rocking at room temperature. Following blocking, membrane was incubated in primary antibody overnight rocking at 4 °C. Mouse monoclonal anti-β-actin (Santa Cruz Biotechnology #sc-47778) or mouse monoclonal anti-GFP (#sc-9996) were used at a dilution of 1:2,500 in 5% milk in TBST. The following day the membrane was washed three times for 5 min each with TBST and then incubated with HRP-conjugated goat anti-mouse antibody (sc-2005) at 1:2000 in 5% milk in TBST for 2 hours at room temperature. Membrane was again washed three times for 5 min each with TBST. Membranes were then incubated for 5 min in Clarity Western ECL Substrate (BioRad #1705060) and immediately imaged on a BioRad ChemiDoc MP imager. Band intensity was quantified using imageJ.

Experiment-specific details regarding sample size and statistical test used can be found in the corresponding Figure Legends. Significant p-values under 0.05 are denoted on all graphs and p-values above 0.05 are considered non-significant (ns). All statistical analysis was performed in GraphPad Prism 9. All data points for each experiment are included (no outlier exclusion was performed). For all whole animal fluorescence analysis, a sample size of 24 animals was generally used, each animal considered a biological replicate. Statistical analysis of high resolution fluorescence confocal imaging (HOE-1::GFP, ATFS-1::mCherry, and TMRE) was conducted on sample sizes between 60 and 80 animals of which animals were collected and imaged on three independent days, each animal considered a biological replicate. For western blot analysis, four independent samples were used for each condition, each sample (containing 50 worms each) is considered a biological replicate. For ddPCR analysis, a sample size of 4 was used for each condition, each sample (containing 10 worms each) is considered a biological replicate, each biological replicate was run in technical duplicate of which the average value was used for analysis.