Effect of the mitochondrial unfolded protein response on hypoxic death and mitochondrial protein aggregation

Mitochondria are the main oxygen consumers in cells and as such are the primary organelle affected by hypoxia. All hypoxia pathology presumably derives from the initial mitochondrial dysfunction. An early event in hypoxic pathology in C. elegans is disruption of mitochondrial proteostasis with induction of the mitochondrial unfolded protein response (UPRmt) and mitochondrial protein aggregation. Here in C. elegans, we screen through RNAis and mutants that confer either strong resistance to hypoxic cell death or strong induction of the UPRmt to determine the relationship between hypoxic cell death, UPRmt activation, and hypoxia-induced mitochondrial protein aggregation (HIMPA). We find that resistance to hypoxic cell death invariantly mitigated HIMPA. We also find that UPRmt activation invariantly mitigated HIMPA. However, UPRmt activation was neither necessary nor sufficient for resistance to hypoxic death and vice versa. We conclude that UPRmt is not necessarily hypoxia protective against cell death but does protect from mitochondrial protein aggregation, one of the early hypoxic pathologies in C. elegans.


INTRODUCTION
Hypoxia, when extreme, will kill aerobic cells. The pathways whereby cells go from healthy to dead when they encounter hypoxia are many and complex. By the time cells are dying from hypoxia, virtually every energy dependent process in the cell is dysfunctional. Pathological processes that occur early in hypoxic exposure when the pathology may still be reversible are of particular interest. This pathologic time course certainly varies by cell type and organism. In mammals, central nervous system neurons and cardiac myocytes are exquisitely sensitive to hypoxia, and during complete ischemia begin dying in only a few minutes after dissolved oxygen is depleted [1,2]. On the other hand, peripheral neurons and skeletal myocytes can endure hours of a similar reduction of oxygen and fully recover. Some organisms can withstand prolonged severe hypoxia while others cannot. Certain hibernating fish and reptiles modulate anabolic and catabolic pathways and can tolerate near anoxic conditions over the winter months, while living in normal oxygen levels in the summer [3]. The primary cellular determinants of this enormous range in hypoxia tolerance are not well defined.
Mitochondria lie at the center of hypoxic biology and are logically the first organelle affected when oxygen levels fall below a crucial threshold. The vast majority of oxygen in metazoan cells is consumed by mitochondrial cytochrome C oxidase enabling the formation of ATP by ATP synthase. When oxygen levels fall, cytochrome C oxidase function fails, ATP levels decrease, and reactive oxygen species (ROS) are produced [4]. Besides ROS, the ATP deficit generated by hypoxia creates havoc throughout the cell including in the mitochondria where ATP-dependent processes such as the proton gradient, protein transport, synthesis, and folding are disrupted. The degree to which the failure of each of these processes contributes to hypoxic cell death is unclear.
In the nematode C. elegans, we and others have identified mutants and RNAis that protect worms from hypoxic cellular and organismal injury . Not surprisingly, what is clear from these studies is that multiple pathways regulate hypoxic sensitivity. However, to what degree the pathological state of the mitochondria influences these pathways and vice versa has not been systematically examined. We have previously shown that hypoxia disrupts mitochondrial proteostasis [6,26]. This disruption is evident by hypoxic induction of the mitochondrial unfolded protein response UPR mt [26], a conserved adaptive pathway that responds to an increase in mitochondrial unfolded protein levels by increasing the synthesis of mitochondrial chaperones, proteases, and other proteins to restore proteostasis [27]. Hypoxia also promotes aggregation of mitochondrial proteins [6,26]. Both UPR mt activation and hypoxia-induced mitochondrial protein aggregation (HIMPA) occur well before there is evidence of cell death in C. elegans or permanent behavioral deficits [6,12]. Thus, disruption of mitochondrial proteostasis occurs relatively early in the hypoxic pathological cascade, but the consequence of disruption of mitochondrial proteostasis and HIMPA on downstream cellular pathology, most importantly cell death is unclear. Even the relationship of the UPR mt to HIMPA is not entirely clear.
Activation of the UPR mt by three RNAis targeting mitochondrial proteins reduced HIMPA and protected from hypoxic death [6]. However, loss of function of atfs-1, a gene which is thought to be essential for UPR mt activation, also decreased HIMPA while atfs-1 gain-of-function mutants increased HIMPA [6]. Thus, whether the UPR mt promotes HIMPA or inhibits it is unclear with the relatively small number of UPR mt activators examined. Here, we make use of the wide variety of C. elegans RNAis and mutants that have been shown to be hypoxia protective, or to induce the UPR mt to address three fundamental questions about the relationship between mitochondrial proteostasis and hypoxic injury. Is activation of the UPR mt a necessary component of hypoxia protective mechanisms? Is UPR mt activation itself hypoxia protective? Does the UPR mt promote or reduce HIMPA?

RESULTS
UPR mt activation not necessary for hypoxia resistance We have previously performed a forward genome-wide screen in Caenorhabditis elegans for genes, whose inactivation by RNAi conferred a hypoxia resistance phenotype (HypR) and identified about 200 such genes [9]. To explore the hypothesis that activation of the UPR mt might be a common or even invariant component of the mechanism underlying hypoxia resistance of these RNAis, we screened through all the HypR RNAis that we could recover from the library and confirm their identity by sequence. Hundred and sixty-one of the HypR RNAi were confirmed and tested for activation of the mitoUPR under normoxic conditions as assessed by the fluorescence levels of the mitoUPR transcriptional reporter hsp-6p::GFP (hsp-6 is the ortholog of human mitochondrial chaperone HSP70 and is a canonical component of the UPR mt ) [28][29][30][31]. RNAi knockdown of only five genes reproducibly induced the mitoUPR reporter (Fig.  1A). Thus, mitoUPR activation, at least as assessed by the hsp-6 reporter, is by no means necessary for hypoxia resistance.
The five RNAis that induced the mitoUPR were against pept-1, sucl-2, C33F10.12, pdha-1, and R04F11.2 (Fig. 1A and Supplementary Table  S1). pept-1 encodes an oligopeptide transmembrane transporter, localized to the apical plasma membrane in the intestine [32]. Reduction of function of pept-1 has been found to reduce protein synthesis rates and levels of essential amino acids [5,33]. A pept-1 loss of function mutant was recently isolated in a screen for hypoxia resistant mutant worms, and its essential role in normal rates of protein synthesis was confirmed [5]. How loss of function of pept-1 activates the UPR mt is unclear although recently the UPR mt has been shown to regulate mitochondrial protein synthesis that would rely on amino acids, whose transport depends on pept-1. On the other hand, sucl-2, C33F10.12, pdha-1, R04F11.2 all encode mitochondrial proteins that form or interact with multi-subunit proteins, where perturbations in stoichiometry have been shown to induce the UPR mt [28,30,34]. Inactivation of these five genes by RNAi led to a range of UPR mt reporter induction (Fig. 1B, C), from 2.3 to 13.7-fold that had no correlation with the previously reported levels of hypoxia resistance where pept-1(RNAi) was most resistant and sucl-2(RNAi) least resistant to hypoxia [9]. atfs-1 is a transcription factor required for UPR mt activation [35,36]. The induction of hsp-6p::GFP by the five UPR mtactivating HypR RNAis was fully suppressed by an atfs-1 loss-offunction mutation (Fig. 1D, E), indicating that the induction of hsp-6p:: GFP by the five Hyp RNAis is acting through the canonical UPR mt pathway.
Hypoxia induces mitochondrial protein aggregation (HIMPA) [6,26]. We found that inactivation of all five UPR mt -activating HypR genes strongly lowered HIMPA as assessed by the ucr-11::GFP mitochondrial protein aggregation reporter [6] (Fig. 1F, G). As shown previously, atfs-1(lf) also reduced the aggregate levels; only pept-1 (RNAi) significantly reduced aggregates further in the atfs-1(lf) background indicating that at least part of the effect of pept-1(RNAi) is independent of atfs-1 (Fig. 1H). atfs-1(lf) did not suppress the hypoxia resistance of any of the five RNAis; thus, neither atfs-1 function nor UPR mt activation is essential to their mechanisms of hypoxia resistance (Fig. 1I). These results suggest that while resistance to hypoxic cell death and HIMPA may be mechanistically associated and that a functional UPR mt may be critical for aggregate formation, the UPR mt is not required for protection from hypoxia and in general activation of the UPR mt does not appear to commonly accompany a hypoxia-resistant phenotype in C. elegans.
UPR mt activation reduces HIMPA To examine more broadly the role of the UPR mt in HIMPA, we tested 12 RNAis previously shown to activate the UPR mt (Supplementary Table S2) [37]. We first confirmed that all 12 RNAis did indeed activate the UPR mt as measured by atfs-1dependent induction of the hsp-6::GFP transcriptional reporter ( Fig. 2A, C). All 12 UPR mt -activating RNAis strongly reduced HIMPA (Fig. 2B). Unlike for the pept-1(RNAi), none of the 12 RNAis significantly reduced HIMPA further in the atfs-1(lf) background (Fig. 2D). Again, atfs-1(lf) paradoxically reduced HIMPA in the no RNAi empty vector condition but not to the same degree of reduction as the UPR mt RNAis. The mitochondrial translation inhibitors doxycycline and meclocycline induce the UPR mt in an atfs-1-dependent manner ( Supplementary Fig. S1) [26,38]. Likewise, doxycycline and meclocycline reduced HIMPA and did not further reduce the aggregate levels in atfs-1(lf) animals ( Fig. 2E). Thus both genetic and pharmacological UPR mt activators reduce HIMPA, while at the same time the UPR mt master regulator gene atfs-1 paradoxically promotes HIMPA, as previously reported [6].

TMRE stains mitochondrial aggregates
While examining the effect of hypoxia on mitochondrial membrane potential, we discovered that the voltage sensitive mitochondrial dye tetramethylrhodamine ethyl ester (TMRE) not only stained functional mitochondria, but also stained hypoxiainduced mitochondrial protein aggregates ( Fig. 4A-C). This finding in the wild type N2 strain importantly demonstrates that aggregates occur in an animal with native mitochondrial proteins. We then compared the aggregates visualized with TMRE to those visible with the ucr-11::GFP reporter. The TMRE-stained aggregates strongly colocalized with UCR-11::GFP aggregates ( Fig. 4D-F). Besides demonstrating importantly that the UCR-11::GFP aggregates are not an artifact of an overexpressed and GFP-tagged protein, TMRE staining can be used in all C. elegans strains to detect mitochondrial aggregates and potentially in other models of hypoxic injury.
HIMPA blocked by hypoxia resistance without UPR mt activation We have previously identified two distinct mechanisms, the insulin/IGF-receptor pathway and the translation machinery, that regulate cytoplasmic proteostasis and profoundly modulate hypoxic survival. The insulin/IGF-receptor pathway has been shown to strongly regulate hypoxic sensitivity, aging, and proteostasis in C. elegans [12,23,39,40]. daf-2 encodes the C. elegans insulin/IGF receptor, and daf-2 reduction-of-function mutants are highly resistant to hypoxic injury via a mechanism that requires the FOXO transcription factor DAF-16 [12,23]. To test whether the daf-2 pathway affected HIMPA, we built the daf-2 (e1370lf);gcIs46[ucr-11p::GFP] strain, confirmed that it was strongly hypoxia resistant (Fig. 5A), and then scored for HIMPA. daf-2 Fig. 1 Only a small subset of hypoxia resistant RNAis activate the UPR mt . A Fluorescence screen of RNAis for UPR mt activation. hsp-6p::GFP induction was quantified for 161 Hyp RNAi clones with the functional categories as originally assigned [9]. Fold induction (mean) of GFP fluorescence is relative to empty vector (EV) RNAi (L4440) (n = 10). Expression of GFP over 2-fold (blue dash line) of empty vector is considered to be positive. Five positive hits confirmed in 2nd screen are shown in red. B Examples of hsp-6p::GFP expression induced by knockdown of the five positive RNAi clones. zcIs13[hsp-6p::GFP] worms were placed onto RNAi bacteria from egg and imaged as young adults 3 days later. Bar, 0.1 mm. C Quantification of GFP expression by corresponding RNAis as in B. Data (mean ± SD) were obtained from four independent experiments, n = 30 animals; ***p < 0.001 with pooled data shown and p-values were calculated using unpaired t-test. D Examples of zcIs13 [hsp-6p::GFP] GFP expression induced by knockdown of positive five RNAi clones in atfs-1(tm4919) background. Worms were placed onto RNAi bacteria from egg and imaged 3 days later. E Quantification of GFP expression by corresponding RNAis in atfs-1(tm4919) background. Data (mean ± SD) were obtained from five independent experiments, n = 30 animals. F Examples of HIMPA in body wall muscles after hypoxia of indicated RNAi treated gcIs46 worms. Green, UCR-11::GFP. Arrow, UCR-11::GFP aggregate. Bar, 2 μm. G Quantification of aggregate numbers as in F. n = 15 animals from three independent experiments; ***p < 0.001. with pooled data shown. P-values were calculated using unpaired ttest. H Quantification of HIMPA by UCR-11::GFP after hypoxia in body wall muscles of indicated RNAi treated atfs-1(tm4919); gcIs46 worms. n = 15 animals from three independent experiments; ***p < 0.001. with pooled data shown. P-values were calculated using unpaired t-test. I Hypoxia resistance is maintained in atfs-1(lf ) background. Wild-type (N2) or atfs-1(tm4919) treated with indicated RNAis and hypoxic survival was scored. n = 9 plates of worms (>100 worms/plate) from three independent trials. *p < 0.05, unpaired t-test. Fig. 2 UPR mt activation Reduces HIMPA. A UPR mt -activating RNAis as assessed by hsp-6p::GFP expression. hsp-6p::GFP induction was quantified for 12 RNAis previously shown to induce the UPR mt . Fold induction of GFP fluorescence is the mean fluorescence relative to empty vector RNAi (L4440) (n = 10). Data were obtained from 10-36 individual animals for all panels; mean ± SD; **p < 0.01, ****p < 0.0001, unpaired t-test. B UCR-11:GFP HIMPA reduced by the UPR mt activating RNAis. C UPR mt activation is partially or completely blocked in the atfs-1(tm4919 lf) animals. D atfs-1(lf) reduces aggregates and suppresses a further reduction in aggregates by UPR mt -activating RNAis. E Mitochondrial translation inhibitors (doxycycline and meclocycline) reduce HIMPA in a non-additive manner along with atfs-1(lf ). **p < 0.01, ***p < 0.001, ****p < 0.0001, n or ns not significant versus EV @ Bonferroni-corrected p < 0.01, unpaired t-test. Fig. 3 Activation of UPR mt does Not Necessarily Produce Hypoxia Resistance. The twelve UPR mt -activating RNAis are tested for hypoxic sensitivity in the wild type background A-L and in the atfs-1(lf ) mutant background M-Q; significance @ p < 0.0001, F-test, simultaneous nonlinear regression; data for each point is from 3 to 12 independent trials with at least 100 animals/trial, mean +/− sem. A-E Five RNAis produced significant hypoxia resistance; F-J Five RNAis have no significant effect on hypoxic sensitivity; and K, L Two RNAis produced significant hypersensitivity to hypoxia. M-Q The hypoxia resistance of the five RNAis is suppressed by atfs-1(lf ).
The translation machinery has been repeatedly implicated in C. elegans hypoxic sensitivity [5,8,9,11]. Translation machinery mutations and RNAis that reduce protein synthesis are strongly hypoxia protective [5,11]. tars-1(gc52) and rars-1(gc47) carry reduction of function mutations in threonyl-aminoacyl and arginyl-aminoacyl tRNA synthetases, respectively, and are strongly resistant to hypoxic death (Fig. 6A), as previously shown [5,11]. Both translation machinery mutants fully blocked HIMPA as measured by both UCR-11::GFP and TMRE ( Fig. 6B-E, G, H). A combination of loss-of-function mutations in two negative regulators of the translation machinery, larp-1 and ncl-1 [5], suppresses the hypoxia resistance of tars-1(gc52) (Fig. 6A) as previously reported [5] and likewise restores HIMPA (Fig. 6F, H). As was seen for daf-2, RNAi knockdown of rars-1 did not activate UPR mt (Supplementary Fig. S2), demonstrating the effect on rars-1 reduction-of-function on hypoxic death and HIMPA was not due to an activation of the UPR mt . These results indicate that hypoxia protective mechanisms of daf-2/daf-16 signaling and reduction of translation do not require UPR mt activation yet protect from HIMPA.

DISCUSSION
Our study shows that a hypoxia resistant phenotype in C. elegans is not commonly accompanied by UPR mt activation and that UPR mt activation is not always hypoxia protective. In other words, UPR mt activation is neither necessary nor sufficient for protection from hypoxic cell death in C. elegans. However, among RNAis selected solely for their ability to induce the UPR mt we did find a strong enrichment for hypoxia resistance, where five of the twelve UPR mt -activating RNAis conferred hypoxia resistance. By comparison, in a screen of over 16,000 RNAis, only 199 were found to reproducibly produce hypoxia resistance [9]. While these twelve are a small sample and the whole genome RNAi screen deliberately ignored weak hypoxia resistance phenotypes, this approximately 30-fold enrichment for hypoxia resistance suggests that UPR mt activation may provide hypoxia protection in certain genetic backgrounds.
We also examined the effect of hypoxia resistance and UPR mt activation on HIMPA. We found that resistance to hypoxic death and resistance to HIMPA were invariantly associated. All RNAis or mutants that were resistant to hypoxic death reduced HIMPA. Importantly, the strongly hypoxia resistant daf-2, tars-1, and rars-1 mutants or RNAis block HIMPA without activating the UPR mt . Thus, UPR mt activation is neither necessary for resistance to hypoxia nor for abrogation of HIMPA. We also found that all UPR mt -activating RNAis as well as doxycycline and meclocycline, which strongly activate the UPR mt , brought hypoxia-induced mitochondrial aggregates to near background levels. To be clear, both hypoxia-resistant and non-hypoxia resistant UPR mt -activating RNAis strongly reduced HIMPA. The effect of the RNAis on activation of the UPR mt was mostly if not completely dependent on a functional atfs-1 gene. This was to be expected as hsp-6 activation is dependent on ATFS-1 [34,36].
The reduction in the aggregates by both UPR mt activation and atfs-1 loss-of-function is a puzzling paradox, albeit consistent with our previously published results [6]. We previously showed that atfs-1 RNAi and the atfs-1(tm4919) loss-of-function mutant reduced HIMPA and an atfs-1 gain-of-function mutant increased aggregates [6]. We had also previously shown that cco-1, letm-1, and F02A9.4 RNAis reduced HIMPA, but we attributed this effect to their hypoxia resistance or reduction in the levels of their corresponding proteins that might form the aggregates themselves [6]. We found here that of these three only letm-1 RNAi reproducibly conferred hypoxia resistance yet all three RNAis, indeed all of the UPR mt activating RNAis, strongly reduced HIMPA. Thus, clearly hypoxia resistance is not necessary for this effect. To reconcile that both atfs-1(lf) and atfs-1-dependent UPR mt activation reduce aggregates, we hypothesize that ATFS-1 functions both to inhibit HIMPA by activating the UPR mt and to promote HIMPA through some other unknown mechanism (Fig. 7). ATFS-1 could both function to reduce aggregates by activation of its downstream UPRmt targets, and in the absence of UPRmt activation function to promote HIMPA through an unknown mechanism. The fact that the level of activation of the hsp-6::GFP transcriptional reporter does not correlate with the ability to suppress hypoxia-induced aggregates ( Fig. 2A, B) suggests that the HSP-6 chaperone levels are not directly responsible for the effect of the UPR mt on aggregation, or that a very low threshold of hsp-6 induction suppresses aggregation fully. Which atfs-1regulated protein(s) might be responsible for promoting HIMPA is unknown; ATFS-1 has been shown to control the synthesis of a large number of proteins involved in mitochondrial proteostasis and mitochondrial function as well as some non-mitochondrial proteins [34,41,42]. Likewise, cytoplasmic and ER proteostasis pathways regulate the UPR mt and mitochondrial proteostasis [42]. Thus, ATFS-1 may function in concert with one of those non-UPR mt pathways to regulate HIMPA directly.
An important technical advance of this work is the discovery that TMRE stains HIMPA. TMRE is a lipophilic cationic dye that accumulates in the matrix of viable mitochondrial, where the membrane potential is highly negative relative to the intermembrane space [43,44]. Besides membrane potential driven accumulation, TMRE has been shown to bind to both surfaces of the inner membrane; however, whether it is binding to lipids, proteins, or both at the inner mitochondrial membrane is unclear [43]. The aggregates appear to be in the matrix of the mitochondria although the resolution is inadequate to be certain. Other than UCR-11::GFP which strongly colocalizes with the TMREstained aggregates, we have no empirical evidence for what composes the TMRE-aggregates. In principle, TMRE can bind nonspecifically to hydrophobic and negatively charged surfaces. Thus, denatured mitochondrial proteins, lipids, and even nucleic acids are reasonable candidates. Regardless of the nature of the binding target, the discovery that TMRE binds to hypoxia-induced mitochondrial aggregates rules out that HIMPA is an artifact of the GFP reporter overexpression. TMRE can be used to examine for HIMPA in any hypoxic/ischemic model without the need for expression of a tagged protein, electron microscopy, or immunofluorescence.
In summary, resistance to hypoxic organismal death and resistance to hypoxia-induced mitochondrial protein aggregation are strongly correlated. This suggests that HIMPA may be an early hypoxic pathology along the pathway to eventual hypoxic cell death. However, it is important to emphasize that we have not directly measured any aspect of mitochondrial function to determine whether HIMPA correlates with abnormalities in mitochondrial function. On the other hand, activation of the UPR mt is neither necessary nor sufficient for hypoxia resistance but may weakly promote hypoxic survival in certain contexts. Activation of the UPR mt did invariably reduce HIMPA whether or not it conferred resistance to hypoxic death. This effect of the UPR mt favors a model, where the UPR mt directly abrogates Fig. 5 Reduction-of daf-2 function Reduces HIMPA. A As previously reported [12], daf-2(e1370) animals are hypoxia resistant in a daf-16 dependent manner and in the gcIs46 background (mean ± SD, n = 4-12 independent trials). B, C Comparison of gcIs46::GFP HIMPA in the wild type B and daf-2(e1370) background C. D-F Comparison of HIMPA by TMRE in wild type (N2), daf-2(e1370), and in daf-2(e1370);daf-16(mu86). G Quantitation of GFP HIMPA in gcIs46 and daf-2(e1370);gcIs46 (mean ± SD of 18 animals). H Quantitation of TMRE HIMPA in the strains with the noted genotypes (mean ± SD of 11-24 animals). ****p < 0.0001, unpaired t-test. Scale bars = 2 µm. mitochondrial protein aggregation during hypoxia. Future investigation will continue to define the breadth of mechanisms that protect cells from hypoxic death, to define the components of the UPR mt that inhibit HIMPA, and the ATFS-1-regulated mechanism that promotes it.

Hypoxic incubations and hypoxic death
Synchronized young adult worms were subjected to hypoxia as described previously except that hypoxic incubation temperature was 26-26.5°C [12,46]. Briefly, each plate of worms was washed into a 1.5 ml tube with 1 ml of M9 buffer (22 mM KH 2 PO 4 , 22 mM Na 2 HPO 4 , 85 mM NaCl, and 1 mM MgSO 4 ), and after the worms settled by gravity, all but 100 μl of M9 was removed. The tubes were then placed in the anaerobic chamber (Forma Scientific, Waltham, MA, USA) for incubation times ranging from 10 to 34 h as indicated. Oxygen tension was always ≤ 0.3%. Following the hypoxic incubation unless otherwise noted, worms were transferred using glass Pasteur pipettes onto seeded NGM plates and recovered at 20°C for 24 h. Normoxic incubations were otherwise identical except performed in a 26.5°C room air incubator. Organismal death was scored as previously described [46]. Briefly, animals were scored as dead if pharyngeal pumping, spontaneous and evoked movement (touching with a platinum wire) were Fig. 6 Loss-of function mutations in the translation machinery reduces HIMPA. A As previously reported [5,11], rars-1 (gc47) and tars-1 (gc52) animals are resistant to hypoxia and the resistance of tars-1(gc52) is maintained in a gcIs46 background and is suppressed by larp-1(q783) ncl-1(gc53) (mean ± SD of 6-12 independent trial with >100 animals/trial). B, C Comparison of gcIs46::GFP HIMPA in the wild type (B) and tars-1 (gc52) backgrounds (C). D-F Comparison of HIMPA by TMRE in wild type (N2), tars-1(gc52), and in tars-1(gc52); larp-1(q783)ncl-1(gc53) animals G Quantification of GFP aggregates in gcIs46 and tars-1(gc52);gcIs46 (mean ± SD of 12-19 animals). H Quantification of TMRE aggregates in the strains with the noted genotypes (mean ± SD of 10-28 animals).****p < 0.0001, unpaired t-test. Scale bars = 2 µm. Fig. 7 Hypoxia induces organellar stress that can manifest in the mitochondria as protein aggregation (HIMPA). ATFS-1, the UPR mt master regulator, both reduces HIMPA by activating the UPR mt and promotes HIMPA by an unknown mechanism. HIMPA may contribute to but is not required for cellular and organellar death. not observed. For the mitochondrial aggregates assay, synchronized worms were kept on NGM plates and placed into anaerobic chamber for 12 h and then mounted for confocal imaging on glass slides [6].

RNAi experiments
The bacterial RNAi feeding strain L4440 carrying the empty RNAi vector strain was from the Ahringer C. elegans RNAi library (MRC Gene service, Swindon, UK) and was used as the negative control (Empty Vector-EV) for all RNAi experiments [47]. Bacterial strains containing RNAi plasmids were cultured and induced with 0.1% β-lactose in 100 mg/ml ampicillin for 24 h at 23°C as described previously [9]. Worms were synchronized on RNAi plates for 3.5 days (N2) or 4.5 days (rars-1(gc47) background) until reaching adulthood; worms not reaching adulthood were excluded.
Mitochondrial UPR reporter and screening SJ4100 (zcIs13[hsp-6p::GFP]) was used as a reporter for mitochondrial UPR activation [28]. Animals were synchronized from eggs by bleaching gravid adults, allowed to develop to adulthood, and then mounted on agar pads for imaging. All UPR mt fluorescent reporter determinations were performed with worms exposed only to normoxic conditions. Briefly, a minimum of ten worms were mounted onto an agar pad on a standard microscope slide and imaged on a Zeiss axioskope 2 at ×10 magnification. Using ImageJ software (https://imagej.nih.gov/ij/), each worm was outlined and the fluorescence intensity within the area was measured and background intensity was then subtracted to obtain the total intensity/worm as previously described [26].
Chemical treatment and TMRE staining. Concentrated stocks of doxycycline (225 mM) and meclocycline (294 mM) in DMSO (Dimethyl sulfoxide) (Sigma-Aldrich Corp, St. Louis, MO, USA), were diluted to a final concentration of 100 μM with nuclease-free, deionized water and 300 µl was spread evenly onto a NGM plate seeded with OP50 onto which were placed synchronized eggs allowed to develop into 1-day-old adults. Plates containing TMRE (ThermoFisher, Waltham, MA, USA) were made the same way as described above with 300 µl at a final concentration of 5 μM spread onto the plate. For TMRE staining, 1-day-old synchronized worms were transferred onto the TMRE-containing plates for 2 h before the plates were placed into the anaerobic chamber for 12 h for hypoxic exposure.

Aggregate imaging and counting
Images of UCR-11::GFP and TMRE staining were acquired with confocal microscopy as previously described [6,26]. Briefly, paralysis was produced by mounting worms in a solution of 50 mM levamisole (Sigma-Aldrich Corp., St. Louis, MO, USA) in M9 prior to imaging. Images were acquired from at least 5-10 randomly selected worms at 1024 × 1024 resolution using a 63× objective with 8 × zoom producing a 23.07 × 23.07 μm image (defined as one high power field, HPF). All images were acquired as a ten slice Z-stack with scan speed of 800-1800 Hz and flattened as a maximum intensity projection prior to analysis. The wavelength of the laser to capture the signal for GFP and TMRE signals was 500-530 nm and 620-700 nm, respectively. Aggregates were counted per HPF by an observer blinded to condition.

Statistics
Two-sided unpaired or paired, as appropriate, t-tests were used for statistical comparisons and all variances were of a similar magnitude except for death versus hypoxic exposure time curves (Fig. 3), where simultaneous curve-fitting was performed [48,49]. Simultaneous curvefitting forced the test RNAi death curve fit and the L4440 empty vector control curve fit to the same hypoxic exposure time producing 50% death (LT50) and F-tests were performed for statistical significance of the change in variance of the LT50 versus fitting each curve independently. Statistics were calculated using GraphPad Prism 6.01 (San Diego, CA, USA) for simultaneous curve-fitting, t-tests, or Excel 2007 for t-tests (Microsoft, Redmond, WA). Values are expressed as mean ± SD of at least three independent experiments. A minimum p value of ≤0.05 was considered significant and was Bonferroni-corrected for multiple comparisons where appropriate. Sample sizes for technical replicates for number of animals was based on the variability from animal to animal for fluorescence-based assays and was always ≥10 or a minimum of 30 animals, where a percent of the population was scored. Based on our previous experience with hypoxic death population assays biological replicates were at least three per condition, and if significant variability was observed among replicates more were performed until clear reproducibility was achieved. In terms of randomization, strains were not randomized as they are genetically identical and raised under environmentally identical conditions.