Eukaryotic Elongation Factor 2 Kinase EFK-1/eEF2K promotes starvation resistance by preventing oxidative damage in C. elegans

Yan, Junran; Bhanshali, Forum; Shuzenji, Chiaki; Mendenhall, Tsultrim T.; Taylor, Shane K. B.; Ermakova, Glafira; Cheng, Xuanjin; Bai, Pamela; Diwan, Gahan; Seraj, Donna; Meyer, Joel N.; Sorensen, Poul H.; Hartman, Jessica H.; Taubert, Stefan

doi:10.1038/s41467-025-56766-1

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Published: 18 February 2025

Eukaryotic Elongation Factor 2 Kinase EFK-1/eEF2K promotes starvation resistance by preventing oxidative damage in C. elegans

Junran Yan ORCID: orcid.org/0009-0003-6793-123X^1,2,3,4,
Forum Bhanshali ORCID: orcid.org/0000-0002-0571-9263^1,3,5^nAff11,
Chiaki Shuzenji^1,2,3,5,
Tsultrim T. Mendenhall⁶,
Shane K. B. Taylor ORCID: orcid.org/0000-0003-4904-4314^1,2,3,5,
Glafira Ermakova^1,2,3,5,
Xuanjin Cheng^1,3,5^nAff12,
Pamela Bai^1,2,3,5,
Gahan Diwan^1,3,7,
Donna Seraj^1,3,5,
Joel N. Meyer ORCID: orcid.org/0000-0003-1219-0983⁸,
Poul H. Sorensen^9,10,
Jessica H. Hartman⁶ &
…
Stefan Taubert ORCID: orcid.org/0000-0002-2432-7257^1,2,3,4,5

Nature Communications volume 16, Article number: 1752 (2025) Cite this article

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Abstract

Cells and organisms frequently experience starvation. To survive, they mount an evolutionarily conserved stress response. A vital component in the mammalian starvation response is eukaryotic elongation factor 2 (eEF2) kinase (eEF2K), which suppresses translation in starvation by phosphorylating and inactivating the translation elongation driver eEF2. C. elegans EFK-1/eEF2K phosphorylates EEF-2/eEF2 on a conserved residue and is required for starvation survival, but how it promotes survival remains unclear. Surprisingly, we found that eEF2 phosphorylation is unchanged in starved C. elegans and EFK-1’s kinase activity is dispensable for starvation survival, suggesting that efk-1 promotes survival via a noncanonical pathway. We show that efk-1 upregulates transcription of DNA repair pathways, nucleotide excision repair (NER) and base excision repair (BER), to promote starvation survival. Furthermore, efk-1 suppresses oxygen consumption and ROS production in starvation to prevent oxidative stress. Thus, efk-1 enables starvation survival by protecting animals from starvation-induced oxidative damage through an EEF-2-independent pathway.

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Introduction

One of the most common stresses in nature is starvation. The ability to adapt to starvation is essential for the survival of cells, tissues, and organisms. When challenged by starvation, organisms mount an evolutionarily conserved molecular and physiological response that involves the rewiring of gene expression and metabolism. In free-living animals such as the nematode worm Caenorhabditis elegans, timely activation of the starvation response promotes stress survival and increases evolutionary fitness. Similarly, in poorly vascularized tumors, cancer cells hijack the conserved starvation response to survive and proliferate in nutrient-poor conditions. Therefore, understanding the mechanism and regulation of the starvation response has fundamental biological and biomedical relevance.

A major mechanism of starvation adaptation involves attenuating the energy-demanding process of mRNA translation^1,2. Protein synthesis represents 30 ~ 35% of the cellular ATP economy, of which >99% is used for translation elongation³. The rate of translation elongation is regulated by eukaryotic elongation factor 2 kinase (eEF2K), which is also a major regulator of starvation survival^4,5. In starved mammalian cells, eEF2K attenuates protein synthesis by inactivating eukaryotic elongation factor 2 (eEF2), the rate-limiting driver of translation elongation. Specifically, eEF2K inhibits eEF2 by phosphorylating a conserved T56 residue, which disrupts eEF2’s normal association with the ribosome complex^6,7. eEF2K activation thus allows starved cells to conserve energy for survival⁵. eEF2K is aberrantly upregulated and promotes tumor cell survival in nutrient-poor solid tumors, leading to poor patient prognoses^2,5,8,9,10. Conversely, eEF2K inhibition blocks cancer cell migration and tumor growth^8,11,12,13. However, it is unclear how eEF2K regulates downstream cytoprotective processes to promote starvation survival.

In addition to its role in translation regulation, evidence has recently emerged that eEF2K may act via alternative mechanisms. In cancer cells, eEF2K directly interacts with and phosphorylates the metabolic regulator pyruvate kinase isozyme M2 (PKM2) in a translation-independent manner¹⁴. Mammalian eEF2K also phosphorylates several other proteins¹⁵, suggesting a broader regulatory role. However, these interactions and their relevance have not yet been explored in detail, and eEF2 remains the only certain substrate for this enzyme at this time.

To uncover regulatory roles of EFK-1/eEF2K in stress response, we used the animal model C. elegans, where eEF2K is evolutionarily conserved and plays an important role in stress adaptation. The C. elegans eEF2K ortholog EFK-1 shares sequence homology in its catalytic domain, and phosphorylates the C. elegans eEF2 ortholog, EEF-2, on the equivalent T56 residue^16,17. Similar to eEF2K^-/- mammalian cells⁵, C. elegans efk-1(ok3609) loss-of-function mutants completely lack EEF-2 T56 phosphorylation^16,17 and exhibit impaired starvation survival⁵. This suggests that C. elegans EFK-1 acts via EEF-2 and translation modulation to promote organism survival in starvation, similar to the mechanism described in mammalian cells.

To study how efk-1 promotes starvation survival, we used the efk-1(ok3609) mutant which we confirmed was functionally null and had diminished efk-1 mRNA expression, abolished EEF-2 T56 phosphorylation, and defective starvation survival. Surprisingly, unlike in mammalian cells, EEF-2 T56 phosphorylation is constitutive in both fed and starved wild-type C. elegans and does not increase in starvation. Moreover, C. elegans strains engineered to lack EFK-1 kinase activity or EEF-2 T56 phosphorylation are indistinguishable from wild type with regards to their ability to survive and recover from starvation. These findings strongly suggest an alternative, kinase-independent mechanism of efk-1. To map this noncanonical pathway, we identified two transcription factors, bZIP transcription factor family 2 (ZIP-2) and C. elegans p53-like protein 1 (CEP-1), which are also required for starvation resistance and function downstream in the efk-1 pathway. Transcriptomic profiling of efk-1, zip-2, and cep-1 mutants revealed that the three factors are jointly required for increased expression of DNA repair pathway genes during starvation. Specifically, efk-1 is required to upregulate nucleotide excision repair (NER) and base excision repair (BER) to increase resistance to oxidative DNA damage, which is linked to starvation-induced oxidative stress. Additionally, efk-1 prevents reactive oxygen species (ROS) accumulation in the cell, maintains mitochondrial physiology and morphology, and represses oxygen consumption, all of which may ameliorate oxidative stress during starvation. As these cytoprotective effects of efk-1 are independent from translation elongation regulation and phosphorylation status of EEF-2, our studies reveal a noncanonical mechanism of efk-1-mediated starvation resistance.

Results

efk-1(ok3609) is a null allele of efk-1

The efk-1(ok3609) mutant shows a deficiency in starvation survival⁵, but its molecular lesion has not been precisely defined. Sequencing the efk-1(ok3609) loss-of-function allele revealed that ok3609 is a deletion spanning the 3’ half of exon 2 and the beginning of exon 3, resulting in a frameshift (Fig. 1A). In line with loss of kinase activity, phosphorylation of T56 of EEF-2, the evolutionarily conserved substrate of EFK-1, is abolished in efk-1 mutants (Fig. 1B; Fig. S1A, B), as previously shown^16,17. Thus, like mammalian eEF2K^5,7, C. elegans efk-1 appears to be the sole kinase that phosphorylates T56 of EEF-2. Consistent with frameshift and presumptive nonsense-mediated decay of efk-1 mRNA (Fig. 1A), RT-qPCR revealed that efk-1 mutants have approximately 10-fold decreased efk-1 transcript expression (Fig. 1C). We conclude that ok3609 is a null allele of efk-1.

Fig. 1: *ok3609* is a null allele of *efk-1.*

efk-1 is required for survival and recovery from starvation-induced arrest at the first larval stage⁵, with defects evident in assays measuring both survival and recovery (Fig. 1D, E) as well as in assays only measuring starvation survival but not recovery (Fig. S1C). efk-1 mutants exhibit a milder, but still substantial and highly significant defect compared to strong regulators of starvation survival such as daf-16/Forkhead box O (FOXO) (Fig. S1D). To ascertain that the starvation sensitivity of the efk-1(ok3609) mutant is due to loss of efk-1 function and to study the expression of EFK-1 in vivo, we generated a strain that stably expresses a translational efk-1::GFP fusion protein from its own promoter (efk-1p::EFK-1::GFP; see Methods). In L4 stage worms, efk-1::GFP is expressed mainly in the cytoplasm of the hypodermis, neurons, and intestinal cells (Fig. S1E). In the efk-1(ok3609) mutant background, this transgene restored EEF-2 T56 phosphorylation and L1 starvation survival and recovery to wild-type levels (Fig. 1B, E), indicating complete functional rescue. RT-qPCR analysis revealed approximately 10-fold higher efk-1 mRNA expression in this strain compared to wild type (Fig. 1C), as well as a significant increase of EFK-1 protein compared to an endogenously tagged efk-1::GFP strain (Fig. S1F). However, compared to wild type, the efk-1::GFP overexpressing allele resulted in only a minor increase in substrate phosphorylation (Fig. 1B; Fig. S1A, G), and conferred no gain-of-function activity in starvation resistance (Fig. 1E). Re-expression of efk-1 in the hypodermis, but not in neurons, was sufficient to rescue the starvation survival and recovery defects of the efk-1 mutant (Fig. S1H, I), indicating that EFK-1 expression in this tissue is sufficient to elicit a protective effect. In sum, we validated that efk-1(ok3609) is a null allele of efk-1, and that efk-1 is required for starvation survival and EEF-2 T56 phosphorylation.

Starvation does not increase EEF-2 phosphorylation

In mammalian cells, eEF2K phosphorylates and inactivates eEF2 in nutrient deprivation and hypoxia, which attenuates translation elongation, reduces energy expenditure, and promotes cell survival^5,18. In C. elegans, starvation also leads to global translation attenuation¹⁹, suggesting that EFK-1–EEF-2 might function in this nematode as it does in mammals. To test if C. elegans efk-1 attenuates translation elongation in starvation, we examined EEF-2 T56 phosphorylation (p-EEF-2) in starved L1 and L4 larvae with Western blots. As expected^16,17, EEF-2 T56 phosphorylation was abolished in efk-1 mutants, both in L4 (Fig. 2A; Fig. S2A–C) and L1 (Fig. 2B; Fig. S2D–F) larval stages. In fed efk-1 mutants, we also observed increased basal levels of eIF2α phosphorylation (p-eIF2α), a regulatory mark of translation initiation, possibly indicating compensation for the lack of EEF-2 phosphorylation (Fig. S2B, E).

Fig. 2: EEF-2 T56 phosphorylation is unchanged in starving *C. elegans.*

Surprisingly, however, we did not observe an increase in p-EEF-2 levels in wild-type L4 worms subjected to 8 h of starvation (Fig. 2A; Fig. S2A), or in L1 worms subjected to prolonged starvation of up to nine days (Fig. 2B; Fig. S2D). We validated that the lack of p-EEF-2 induction in 8-h L4 starvation was consistent across various amounts of extracted protein (Fig. S2G), ruling out signal saturation as a possible cause. Transient induction in a shorter time frame is also unlikely, as p-EEF-2 levels remained unchanged in L4 animals starved as briefly as 5 minutes and up to 2 h (Fig. 2C; Fig. S2H). In all the above conditions, p-EEF-2 levels were readily detectable at baseline in well-fed L1 and L4 worms (Fig. 2D; Fig. S2I, J), with no apparent increase in starvation. In fact, p-EEF-2 levels decreased with starvation (Fig. S2A, D). This high baseline level of p-EEF-2 is consistent with previous Western blot and phosphoproteomics studies in C. elegans^16,17,20. In contrast, in A549 lung cancer cells, we observed low or absent p-eEF2 signals in fed conditions and substantial induction after 3 to 6 h of starvation (Fig. 2E; Fig. S2K, L), consistent with studies in multiple mammalian cell lines^5,21. Thus, although canonical EFK-1–EEF-2 signaling is clearly present in C. elegans (Fig. 2A, B), efk-1 surprisingly does not appear to mediate starvation resistance by increasing EEF-2 phosphorylation.

EFK-1 promotes starvation resistance without altering global translation rate

To further study whether starvation arrests translation elongation in a canonical, EEF-2-dependent manner, we assessed a downstream readout of translation elongation arrest. Conditions of translation elongation arrest, including RNA interference (RNAi) knockdown of EEF-2 or amino-acyl tRNA synthetases or treatment with the pharmacological translation elongation inhibitor cycloheximide (CHX)²², induce nuclear localization of the transcription factor ZIP-2 to activate transcription of a downstream gene, infection response gene 1 (irg-1), which is induced in many pathogen exposure paradigms^23,24,25. As EFK-1 is solely responsible for EEF-2 T56 phosphorylation and is a potent inhibitor of EEF-2 activity (Fig. 2A, B), irg-1 expression should be activated if starvation were to arrest translation elongation via the EFK-1–EEF-2 axis. To test this, we starved worms expressing a transcriptional irg-1p::GFP promoter reporter²⁶. In line with the lack of increased EEF-2 phosphorylation in starvation (Fig. 2A, B), we did not observe increased irg-1 promoter activity in L4 worms starved for 6, 24, or 48 h (Fig. 3A; Fig. S3A) or in L1 worms starved for 2, 6, or 15 days (Fig. S3B, C). This lack of irg-1 induction in starvation is consistent with published RNA-seq studies²⁰. In contrast, translation arrest via CHX treatment, eef-2 RNAi, or RNAi against amino-acyl tRNA synthetases nars-1 and aars-2 strongly induced irg-1 promoter activity (Fig. 3A; Fig. S3A, B), as reported^23,27.

**Fig. 3: *efk-1* functions independently of altering global translation rate.**

To complement the irg-1 promoter activity studies, we assessed ZIP-2 localization in the same conditions using a translational ZIP-2::GFP fusion reporter²⁴. As expected, CHX treatment caused ZIP-2 nuclear translocation, but this was not observed in either L4 worms starved up to 48 h (Fig. S3D, E), or in L1 worms starved 2 to 15 days (Fig. S3F–H). The lack of ZIP-2–irg-1 signaling in starvation is consistent with the above data that EFK-1 does not appear to increase EEF-2 phosphorylation in starvation, supporting the hypothesis that EFK-1 functions in starvation in a noncanonical pathway.

Next, we asked whether efk-1 promotes starvation survival via inhibiting translation elongation and whether efk-1 loss causes starvation sensitivity by derepressing translation elongation. In cancer cells, the defective starvation survival of eEF2K-inactive cells is rescued by CHX, demonstrating that eEF2K promotes starvation survival mainly via translation elongation inhibition²¹. Using the same approach in C. elegans, we investigated whether CHX is sufficient to rescue the starvation survival defect of efk-1 mutants. However, we found that various doses of CHX did not improve the starvation survival defect of efk-1 mutants (Fig. 3B; Fig. S3I, J). This was unlikely due to poor CHX uptake or activity, as similar doses of CHX extend worm lifespan²⁸, and higher doses substantially reduce translation and induce the irg-1p::gfp transcriptional reporter (Fig. 3A; Fig. S3A), as published^23,27,29. The fact that CHX supplementation does not rescue the starvation survival defect of efk-1 mutants suggests that their starvation sensitivity is not due to derepressed translation, consistent with the above findings that efk-1 is unlikely to promote starvation survival by inhibiting translation elongation.

Finally, to directly assess if efk-1 inhibits global protein synthesis in starving worms, we assayed global protein synthesis rates using the surface sensing of translation (SUnSET) assay. SUnSET measures de novo protein synthesis rates by puromycin labeling of newly synthesized peptides^30,31,32, and has been widely used in mammalian cells and C. elegans^27,29,33,34. In cell lines, puromycin incorporation is decreased in starvation and other conditions with active eEF2K^35,36. Indeed, we detected decreased protein synthesis in A549 cells after 6 h of starvation (Fig. 3C, D; Fig. S3K), as previously shown³⁵. Next, we used SUnSET to test whether C. elegans efk-1 mutants derepress translation when subjected to 8-h starvation at L4 and 6-day starvation at L1. Interestingly, we did not detect increased total puromycin incorporation in efk-1 mutants compared to wild type in either condition (Fig. 3E–H; Fig. S3L, M). This suggests that efk-1 does not suppress global translation during starvation, consistent with the above data that p-EEF-2 remained unchanged in the same conditions. Taken together, these data suggest that EFK-1 promotes starvation resistance via a noncanonical pathway independent from EEF-2 inhibition and global translation attenuation.

EFK-1 promotes starvation resistance independently of its kinase activity and EEF-2 phosphorylation

To confirm that EFK-1 regulates starvation survival and recovery independently of EEF-2 T56 phosphorylation, we used genome editing to ablate either the kinase activity of EFK-1 or the T56 phosphorylation site of EEF-2. The kinase activity of mammalian eEF2K requires D274, which is essential for ATP binding and present in other members of the alpha-kinase family³⁷. In C. elegans, D274 is conserved as D257, with its surrounding regions mostly identical to human eEF2K (Fig. S4A). To abolish EFK-1 kinase activity, we generated two independent lines bearing the D257A mutation using homology-directed CRISPR-Cas9 genome editing. efk-1 D257A mutants displayed no change in efk-1 mRNA expression when compared to wild type (Fig. 4A), but showed a complete lack of EEF-2 phosphorylation, mimicking the efk-1 null mutant (Fig. 4B; Fig. S4B). Strikingly, efk-1 D257A mutants showed no defect in starvation survival and recovery (Fig. 4C), demonstrating that EFK-1 kinase activity is not required for starvation resistance.

**Fig. 4: *efk-1* promotes starvation survival independently of its kinase activity and of EEF-2 T56 phosphorylation.**

To directly test if EEF-2 T56 phosphorylation is required for starvation survival, we used homology-directed CRISPR-Cas9 genome editing to mutate the phosphorylation site T56 in EEF-2, which is well-conserved from human eEF2 (Fig. S4C). Homozygous eef-2 T56A mutants were inviable and showed an inability to develop a functioning germline. However, balanced eef-2 T56A heterozygotes were viable, and their F1 homozygous offspring were able to undergo larval development at a normal pace, allowing them to be isolated and assayed. We confirmed using Western blots that eef-2 T56A homozygotes lack EEF-2 phosphorylation (Fig. 4D; Fig. S4D). Importantly, eef-2 T56A homozygotes showed no defect in starvation survival (Fig. 4E), consistent with a model that EEF-2 T56 phosphorylation is dispensable for starvation survival. Together, these experiments provide genetic evidence that EFK-1 promotes starvation resistance through a noncanonical, kinase-independent mechanism.

Transcription factors ZIP-2 and CEP-1 function in the efk-1 starvation response pathway

Because efk-1 regulates starvation survival outside of its canonical role in translation regulation, we hypothesized that it may act via transcriptional regulation instead. Starvation arrest induces extensive transcriptomic rewiring to repress many genes while activating specific genes required for mounting the starvation response³⁸. Accordingly, starvation survival in C. elegans requires numerous transcription factors (TFs), including insulin signaling component daf-16, metabolic regulator hlh-30/ Transcription factor EB (TFEB), nuclear hormone receptor 49 (nhr-49), zip-2, and cep-1/TP53^{38,39,40,41,42,43,44}. To delineate how efk-1 regulates downstream gene expression, we searched for starvation-responsive transcription factors with functional links to efk-1 or mammalian eEF2K. First, we selected the transcription factor ZIP-2, which not only activates irg-1 in pathogen response with the binding partner CEBP-2/ CCAAT Enhancer Binding Protein Gamma CEBPG^{23,24,26,45,46}, but is also required for nutrient responses such as starvation survival and dietary restriction-induced longevity^41,47. Interestingly, zip-2 appears to function in starvation without strong activation of irg-1 signaling (Fig. 3A; Fig. S3A), indicating that it may be required to maintain the transcriptional activity of other stress-responsive pathways. In addition, the C. elegans p53 ortholog cep-1 is required for starvation survival^43,48,49, and human p53 interacts genetically with eEF2K in tumor cells⁵⁰. Thus, we hypothesized that zip-2, cebp-2, and/or cep-1 might function with efk-1 to promote starvation survival.

To study the roles of these TFs in the starvation response, we quantified the population survival of previously characterized zip-2, cep-1, and cebp-2 loss-of-function mutants^{23,24,43,51,52} by L1 starvation survival assays. As expected, zip-2, cebp-2, or cep-1 loss caused starvation sensitivity and thus phenocopied efk-1 loss⁵ (Fig. 5A). cebp-2 mutants also showed growth defects under standard growth conditions, whereas zip-2 and cep-1 mutants did not (Fig. S5A). To examine the genetic interactions of efk-1 with zip-2, cebp-2, and cep-1, we next constructed respective double mutants. In standard growth conditions, zip-2;efk-1 and cep-1;efk-1 mutants were viable without apparent phenotypes (Fig. S5B), whereas efk-1;cebp-2 mutants could not be obtained, suggesting synthetic lethality or sterility. Interestingly, zip-2;efk-1 and cep-1;efk-1 double mutants showed no additive defect in L1 starvation survival compared to respective single mutants (Fig. 5B, C). Because zip-2 and cep-1 both genetically interact with efk-1, we next constructed a zip-2;cep-1 double mutant, which also showed no additive starvation survival defect compared to the respective single mutants (Fig. 5D; Fig. S5C). Thus, zip-2 and cep-1, but not cebp-2, are components of the efk-1 starvation response pathway.

**Fig. 5: The TFs *zip-2* and *cep-1* are required in the *efk-1* pathway for starvation survival.**

Next, we asked if zip-2 and cep-1 function epistatically to efk-1 in the pathway. Since efk-1 overexpression did not elicit a gain-of-function effect (Fig. 1E), we instead tested whether overexpressing ZIP-2 or CEP-1 rescues the starvation survival phenotype in the efk-1 null background. For ZIP-2, we observed rescue of the starvation survival and recovery defect of the efk-1 mutants with an integrated ZIP-2::GFP construct^24,53 (Fig. 5E), which caused 20-fold overexpression of zip-2 transcript (Fig. S5D). For CEP-1, we observed rescue of the starvation survival and recovery defect of the efk-1 mutant with both a CEP-1::3xFLAG extrachromosomal array (Fig. 5F), and an integrated CEP-1::GFP construct expressed at approximately 1.5-fold level of wild-type cep-1 mRNA⁵⁴ (Fig. S5E). Together, these data show that zip-2 and cep-1 function downstream of efk-1 to promote starvation survival.

DNA repair pathways are transcriptionally upregulated by efk-1, zip-2, and cep-1 during starvation

To identify genes and pathways regulated by efk-1, zip-2, and cep-1 in starvation, we performed RNA sequencing (RNA-seq) of fed and starved wild-type worms and efk-1, zip-2, and cep-1 mutants (Fig. 6A; Fig. S6A). We found that in 8-h starved, wild-type L4 stage worms, 3302 genes were significantly altered (p < 0.005, FDR < 0.05), including 1493 upregulated and 1809 downregulated genes (Supplementary Data 1). To validate our expression analysis, we reanalyzed published RNA-seq data of 6-h starved L4 stage wild-type worms²⁰ and compared them to our dataset. Significantly regulated genes (p < 0.005 and FDR < 0.05) shared a substantial overlap and correlated well between both datasets (r² = 0.87, p < 2.2e-16; Fig. S6B, C). Our data also identified several experimentally confirmed starvation-induced genes, such as fmo-2, cpt-3, fat-3, icl-1, fat-2, acs-11, lbp-1, and efk-1, and starvation-repressed genes, such as lbp-8, ech-6, acdh-1, acdh-2, acox-1.4, cpt-4, and fat-7^5,42,55 (Fig. S6D). Additionally, consistent with published transcriptomic data²⁰, we did not observe transcript induction of irg-1 (Fig. S6E, F), confirming that starvation does not activate irg-1 (Fig. 3A). Overall, our data recapitulate known short-term starvation responses.

**Fig. 6: DNA repair pathways are upregulated in WT and attenuated in *efk-1*, *zip-2*, and *cep-1* mutants during starvation.**

Next, we analyzed the transcriptional dysregulations in starved efk-1, zip-2, and cep-1 mutants. We found that among the 3302 genes significantly regulated in wild type, 1322 failed to be regulated in efk-1 mutants, 1036 in zip-2 mutants, and 1427 in cep-1 mutants. We call these efk-1, zip-2, or cep-1 dependent genes, respectively (Supplementary Data 1). Interestingly, efk-1, zip-2, and cep-1 mutants showed similar transcriptome deregulations in starvation compared to wild type: among the 1322 efk-1-dependent, 1036 zip-2-dependent, and 1427 cep-1-dependent genes, nearly half (606 genes) are dependent on all three genes (Fig. 6B; Supplementary Data 1). To obtain an unbiased profile of starvation gene dysregulations in these mutants, we performed second-order comparisons of efk-1, zip-2, and cep-1 starved-versus-fed profiles to that of wild type. efk-1 and zip-2 mutant starvation profiles showed strong positive correlation with each other (p < 0.25, correlation r² = 0.84, p < 2.2e-16; Fig. S6G), whereas efk-1 and cep-1 profiles exhibited a moderate positive correlation (p < 0.25, correlation r² = 0.57, p < 2.2e-16; Fig. S6H). Thus, efk-1, zip-2, and cep-1 control a shared set of genes in starved C. elegans.

To identify biological processes jointly regulated by efk-1, zip-2, and cep-1 in starvation, we used two complementary approaches. First, we used gene set enrichment analysis (GSEA) to capture broad transcriptomic changes in pathways in an unbiased manner. Starving wild-type worms upregulated the expression of known starvation-inducible pathways, such as “autophagy”, “FOXO signaling pathway”, and “longevity regulating pathway”, and downregulated pathways such as “drug metabolism”, “metabolism of xenobiotics by cytochrome P450”, and “oxidative phosphorylation” (Supplementary Data 2), consistent with previous reports^20,38,56. Interestingly, we also observed an upregulation of DNA repair pathways, such as “nucleotide excision repair”, “base excision repair”, “homologous recombination” (HR), and “Fanconi anemia pathway” (FA) (Supplementary Data 2). Next, we performed GSEA on the starvation profiles of efk-1, zip-2, and cep-1 mutants. Similar to the gene-level correlation, the pathways enriched in efk-1, zip-2, and cep-1 mutants correlated strongly with each other (r² = 0.83 for efk-1 vs. zip-2 regulated pathways, r² = 0.87 for efk-1 vs. cep-1 regulated pathways, p < 2.2e-16; Fig. 6C, D). Strikingly, among the pathways that were upregulated in starved wild type (p < 0.05, padj < 0.05, ES > 0.5), DNA repair pathways such as NER, BER, HR, and FA were downregulated in all three mutants compared to wild type (p < 0.05, padj < 0.05, ES < -0.5) (Fig. 6E).

To complement the GSEA analysis, we performed overrepresentation analysis (ORA) on the 606 starvation-responsive genes that are dependent on efk-1, zip-2, and cep-1 (Supplementary Data 3). In agreement with the GSEA analysis, several DNA damage response and DNA repair pathways were amongst the top ORA categories, including NER and BER, two important pathways for oxidative damage repair (Fig. 6F). Taken together, both functional analysis approaches suggest that efk-1, zip-2, and cep-1 are jointly responsible for upregulating transcription of DNA repair genes during starvation.

NER is required for starvation survival and acts in the efk-1 pathway

NER is an evolutionarily conserved DNA repair mechanism responsible for removing bulky, DNA-distorting adducts caused by UV radiation, oxidative stress, or genotoxins^57,58,59,60. NER consists of two branches, global-genome NER (GG-NER) and transcription-coupled NER (TC-NER). In C. elegans, TC-NER is vital for DNA repair in somatic cells, whereas GG-NER is required in both germline and soma^58,61. To test if NER-mediated DNA repair is required for starvation survival, we assayed the starvation sensitivity of several NER mutants. We studied the xpa-1, xpf-1, ercc-1, and xpg-1 mutants that are generally NER deficient, the xpc-1 mutant that is deficient only in GG-NER, and the csa-1 and csb-1 mutants that are deficient only in TC-NER⁶². Most NER mutants showed no visible defects in unstressed conditions, except ercc-1, which had a slower growth rate (Fig. S7A). However, all of these mutants showed reduced survival in the L1 starvation survival assay, indicating that functional NER is required for starvation survival (Fig. 7A–C). To test if NER promotes starvation survival in the efk-1 pathway, we studied the genetic interaction between efk-1 and xpa-1 and xpf-1. efk-1;xpa-1 and efk-1;xpf-1 double mutants showed no genetic interaction in the L1 starvation survival assay (Fig. 7D, E). Thus, functional NER is required for starvation survival in the efk-1 pathway. We also studied genetic interactions of NER with zip-2 and cep-1 by assaying zip-2;xpa-1 and zip-2;xpf-1 double mutants, and cep-1;xpf-1 double mutants (we did not attempt to generate a cep-1;xpa-1 double mutant as cep-1 and xpa-1 are genetically linked). In line with the above genetic interaction data, zip-2; xpa-1, zip-2;xpf-1, and cep-1;xpf-1 double mutants showed no synthetic starvation survival defects (Fig. 7F–H). All above double mutants had no growth defect compared to wild type (Fig. S7B–F). Taken together, this supports a model where efk-1, zip-2, and cep-1 jointly regulate NER to promote starvation survival.

**Fig. 7: NER is required for *efk-1* mediated starvation resistance.**

The soma is the main site of the starvation response during L1 arrest, as the germline only consists of two primordial germ cells that are transcriptionally quiescent³⁸. Somatic cells depend on TC-NER, although GG-NER is also present⁶¹. Thus, we asked if efk-1 acts with TC-NER, GG-NER, or both during L1 arrest, using xpc-1 and csb-1 mutants to assess requirements for GG- and TC-NER loss, respectively. Neither efk-1;xpc-1 nor efk-1;csb-1 double mutants showed additive starvation survival phenotypes compared to respective single mutants (Fig. S7G–I). Thus, both GG- and TC-NER are required for efk-1-mediated starvation resistance.

efk-1 and cep-1 play a minor role in somatic and germline NER

Because they are required to induce NER genes and genetically interact with the NER pathway, we hypothesized that efk-1, zip-2, and cep-1 may be involved in NER-mediated DNA repair in fed worms as well. To test this hypothesis, we assessed somatic and germline NER capacity by measuring UV-C sensitivity in mostly somatic L1 larvae and in the germline of young adults, respectively. To test if efk-1, zip-2, and cep-1 are required for NER in somatic cells⁶¹, we exposed these mutants to UV-C at the L1 stage and quantified their ability to recover to the L4 stage after irradiation. Whereas the recovery of zip-2 mutants was similar to wild type (Fig. S7J), efk-1 and cep-1 mutants displayed a mild reduction in their ability to recover from UV-C exposure (Fig. 7I, J), indicating decreased somatic NER capacity. To test if efk-1, zip-2, and cep-1 are required for NER in the germline⁶¹, we exposed these mutants to UV-C as young adults, and quantified the embryonic lethality of the immediate offspring. The proportion of unhatched eggs was reduced in efk-1 and cep-1 mutants at 40 J/m² (Fig. 7K, L), whereas zip-2 mutants showed no defect (Fig. S7K), indicating that efk-1 and cep-1, but not zip-2, are involved in germline NER. Altogether, these data show that efk-1 and cep-1 play a minor role in both somatic and germline NER, whereas zip-2 is dispensable.

BER is required for starvation survival and acts in the efk-1 pathway

BER is a DNA repair pathway that targets oxidative base lesions, such as 8-oxoguanine (8-oxoG), 5-hydroxymethyluracil (5-hmU), and uracil⁶³. To test if BER-mediated DNA repair is required for starvation survival, we studied the sensitivity of BER mutants to prolonged L1 starvation. BER requires the apurinic/apyrimidinic (AP) endonucleases apn-1 and exo-3, and glycosylases ung-1 and nth-1, which target uracil and other oxidative lesions, respectively. We also tested the poly ADP-ribose polymerase parp-2, which is involved in mammalian BER and conserved in C. elegans⁶³. apn-1, exo-3, ung-1, and nth-1 mutants have no growth phenotypes in unstressed conditions, whereas parp-2 mutants feature a mild developmental delay (Fig. S8A). Interestingly, apn-1, exo-3, ung-1, nth-1, and parp-2 loss-of-function mutants all showed starvation survival defects (Fig. 8A). To test the interaction of efk-1 with the BER, we constructed efk-1;apn-1, efk-1;exo-3, and efk-1;parp-2 double mutants (Fig. S8B–D). Both efk-1;apn-1 and efk-1;exo-3 mutants showed no additive starvation-related defects compared to respective single mutants, indicating involvement in the same pathway (Fig. 8B, C). However, efk-1;parp-2 mutants showed a substantial additive starvation survival defect (Fig. 8D), suggesting that efk-1 and parp-2 promote starvation survival in separate pathways, with parp-2 and efk-1;parp-2 animals also showing slower growth in unstressed conditions (Fig. S8D). Taken together, our data show that BER components are required for starvation resistance in the efk-1 pathway, with the exception of parp-2.

**Fig. 8: BER is required for *efk-1* mediated starvation resistance.**

efk-1 protects against elevated ROS during starvation

Oxidative stress is increased in prolonged starvation, and oxidative stress defense is required for successful starvation survival^64,65. Consistent with high oxidative burden, starving worms also accumulate more DNA lesions and express DNA damage-like signatures^66,67,68. To test if efk-1 mutants have elevated ROS during starvation, we quantified ROS levels in efk-1 mutants using two cell-permeable dyes for intracellular superoxide, dihydroethidium (DHE) and CellROX Green^64,65. Compared to wild type, efk-1 mutants exhibited increased DHE levels from day 3 of starvation onward and increased CellROX Green levels from day 0. Importantly, both signals were rescued by the antioxidant N-acetylcysteine (NAC; Fig. 9A, B; Fig. S9A–D). Thus, efk-1 mutants accumulate abnormal levels of ROS early in starvation.

**Fig. 9: *efk-1* mutants are sensitive to starvation-induced oxidative stress.**

Next, we asked if oxidative stress is a cause of the starvation-induced death in the efk-1 mutant. To address this, we treated starving L1 worms with either of two antioxidants, NAC or Vitamin C (VitC). Supplementation with 5 mM NAC completely rescued the survival defect of efk-1 mutants (Fig. 9C), whereas 2.5 mM VitC had a less substantial, but still significant effect (Fig. 9D); 2.5 mM NAC and 5 mM VitC were also effective (Fig. S9E, F). This shows that oxidative stress is an important cause of the starvation survival defect in efk-1 mutants.

To test if oxidative damage is similarly the cause of the starvation-induced death in the other components of the pathway, we treated zip-2, cep-1, xpa-1 and xpf-1 mutants with NAC during L1 starvation. As seen with efk-1 mutants, supplementation with 5 mM NAC completely rescued the starvation survival defects of zip-2, cep-1, xpa-1, and xpf-1 mutants (Fig. 9E–H; Fig. S7G, H). These data indicate that zip-2, cep-1 and the NER promote resistance to starvation-induced oxidative stress. In the case of the NER mutants, this is likely due to promoting oxidative DNA damage repair.

Next, we asked whether starvation activates NER in an efk-1 dependent manner. As starvation upregulates the expression of NER genes via efk-1 (Fig. 6E-F), we hypothesized that starvation would increase NER capacity and render worms more resistant to exogenous DNA damage. To test this hypothesis, we treated L1 stage wild-type and efk-1 mutant worms with up to 9 days of starvation, irradiated them with UV-C, and then assessed population survival. We observed that wild-type worms that have been starved for 2 to 6 days are more resistant to UV irradiation than freshly hatched ( ~ 24 h) animals or animals starved for more than 6 days (Fig. 9I). In contrast, starved efk-1 mutants do not show such increased resistance to UV (Fig. 9I). As UV resistance is indicative of NER function⁵⁸, this increase of UV resistance post-starvation implies that starvation treatment is sufficient to activate NER in an efk-1 dependent manner. Taken together, these data suggest that efk-1 is required for activation of DNA repair to counteract starvation-induced oxidative DNA damage.

efk-1 protects against mitochondrial ROS

The mitochondria are an important site of ROS production⁶⁹, and decline of mitochondrial quality during starvation^64,66,70 correlates with oxidative stress phenotypes of starved worms. To determine if the mitochondria are a source of the increased ROS levels observed in starved efk-1 mutants, we examined whether the starvation defect of the efk-1 mutant was altered by the mitochondrially-targeted antioxidant mitoquinone (MitoQ)^71,72, which ameliorates stress-induced mitochondrial ROS generation and protects against mitochondrial dysfunction-induced phenotypes in mammalian cells and C. elegans^72,73,74. Indeed, MitoQ supplementation partially rescued the starvation survival defect of efk-1 mutants (Fig. 10A; Fig. S10A). This implies that the reduced survival of starving efk-1 mutants is caused at least in part by mitochondrial defects, such as excess mitochondrial ROS.

**Fig. 10: *efk-1* mutants exhibit mitochondrial defects in starvation.**

Prolonged L1 starvation is associated with both decreased mitochondrial content and increased mitochondrial fragmentation^64,66. To further characterize the starvation-induced mitochondrial phenotypes of efk-1 mutants, we first studied mitochondrial content by quantifying mitochondrial DNA copy number. Consistent with previous studies, we observed that mitochondrial DNA copy number decreased during L1 starvation (Fig. S10B). However, mitochondrial DNA copy number in efk-1 mutants remained similar to wild type, both in prolonged (0 ~ 12 days) and early starvation (0 ~ 3 days) (Fig. S10B, C). Thus, altered mitochondrial content is likely not the cause of excess mitochondrial ROS in efk-1 mutants.

Next, we examined the mitochondrial morphology of efk-1 mutants in prolonged L1 starvation. We introduced the ok3609 null allele into the muscle mitochondrial GFP reporter myo-3::GFP(mit) using CRISPR-Cas9 genome editing, as the reporter appeared to be genetically linked with efk-1. Interestingly, efk-1 mutants showed more mitochondrial fragmentation than wild-type worms at baseline, which is exacerbated in worms starved for 6 to 9 days (Fig. 10B, C; Fig. S10D–F), implying a role for efk-1 in maintaining mitochondrial morphology.

Finally, we continuously monitored oxygen consumption during L1 starvation in live worms. Strikingly, we observed a burst of oxygen consumption in efk-1 mutants in early starvation (16 ~ 20 h after hatch), which was rescued in animals expressing the efk-1::GFP transgene (Fig. 10D; Supplementary Data 4). This excess oxygen consumption cannot be attributed to increased motility, as efk-1 mutants are no more active than wild-type worms during the specified timeframe (Fig. S10G; Supplementary Data 4). Thus, the excess oxygen consumption in efk-1 mutants is more likely linked to the heightened ROS and other oxidative phenotypes, consistent with defective mitochondrial function during starvation. Together, these data show that efk-1 likely offers protection against excess oxygen consumption and accumulation of mitochondrial ROS during L1 starvation.

Discussion

Cells and organisms survive starvation by mounting a cellular response that involves rewiring of gene expression and metabolism. During starvation, cells reprogram gene expression from growth and development to conservation and cytoprotection. The kinase eEF2K is a starvation response regulator that canonically functions by allowing the cells to conserve cellular energy, specifically by attenuating translation elongation via phosphorylation of its principal substrate, eEF2. However, emerging evidence suggests that eEF2K may act via additional mechanisms to promote cytoprotection. Here, we show that C. elegans EFK-1/eEF2K acts in a noncanonical, kinase-activity-independent, and EEF-2/eEF2 phosphorylation-independent manner to promote starvation survival and recovery. EFK-1 cooperates with TFs CEP-1/p53 and ZIP-2 to rewire transcription during early starvation, upregulating DNA repair processes such as NER and BER. In turn, these DNA repair pathways are required to counteract oxidative damage that arises from prolonged starvation. Furthermore, EFK-1 lessens oxidative burden during starvation by maintaining mitochondrial quality, preventing hyperactive oxygen consumption, and repressing ROS accumulation. Collectively, we propose a model where efk-1 promotes starvation survival via a noncanonical pathway that involves preserving mitochondrial function, preventing the accumulation of mitochondrial ROS, and promoting the repair of oxidative DNA damage (Fig. 10E). This is to our knowledge the first characterization of a EEF-2/eEF2-independent mechanism linked to EFK-1/eEF2K mediated stress adaptation.

efk-1, cep-1, and zip-2 promote genome integrity in the same starvation resistance pathway

C. elegans efk-1, zip-2, and cep-1 are all required for starvation survival and recovery^5,41,43, but how each factor promotes starvation adaptation was unknown. Our data indicate that these three regulators function in the same signaling circuit to achieve starvation resistance. Inducing DNA damage response pathways is a key output of this signaling circuit, and the activity of these repair processes is itself critical for surviving starvation. Although induction of DNA repair pathways may have been expected from cep-1, an established guardian of genome integrity^43,75, it is a previously unreported role for efk-1 and zip-2. Additionally, we also show that efk-1 and cep-1 play a part in promoting DNA repair and genome stability in feeding, growing animals, whereas zip-2 is less important in this context.

Our data provides additional links of efk-1/eEF2K and cep-1/TP53 in stress and DNA damage response. In cancer cells, mice, and C. elegans, both cep-1/TP53 and efk-1/eEF2K play a similar, conserved role in mediating the cellular decision between apoptosis and DNA repair in response to genomic insults^17,50,76. In C. elegans, both efk-1 and cep-1 are required to maintain germline integrity by regulating germline apoptosis and repair^50,75,77. Consistent with these, we show that C. elegans efk-1 and cep-1 also act together to promote genomic integrity in starvation. Overall, our study highlights intertwined regulatory roles of efk-1/eEF2K and cep-1/TP53 in evolutionarily conserved responses to genotoxic stress.

efk-1 protects against mitochondrial ROS

Why is it important for C. elegans to induce DNA repair pathways during starvation? ROS levels increase during long-term starvation in wild-type animals⁶⁴, but our data show that this phenomenon is exacerbated in efk-1 mutants. Our data suggests that efk-1 offers oxidative protection for the starving cell in two ways. On one hand, efk-1 prevents excess ROS accumulation in the cell. A substantial portion of this ROS originates from the mitochondria, as shown by the ability of mitoQ to partially rescue the starvation defect in efk-1 mutants, although non-mitochondrial ROS also play a part. On the other hand, efk-1 protects from ROS-induced oxidative damage by activating the DNA repair machinery. In line with this model, exogenous provision of antioxidants completely rescues the starvation survival defect not only of efk-1 mutants, but also zip-2, cep-1, and DNA repair pathway mutants, pinpointing oxidative DNA damage as a key adverse event during starvation. We also show that efk-1 regulates mitochondrial physiology and function during starvation. efk-1 mutants display increased mitochondrial fragmentation and higher oxygen consumption rates, which reflect mitochondrial dysfunction. In prolonged starvation, such increased mitochondrial dysfunction in efk-1 mutants might exacerbate starvation-related mitochondrial defects and cause accumulation of ROS and oxidative damage in the cell.

EFK-1 functions independently of EEF-2 phosphorylation and global translation regulation

Mammalian eEF2K protects against starvation via inducible eEF2 phosphorylation and translation attenuation. However, we found contrary evidence suggesting that this is not a mechanism of efk-1-mediated starvation resistance in C. elegans. Although EFK-1 is the sole kinase for EEF-2 T56 phosphorylation in C. elegans^16,17, substantial T56 phosphorylation is present in feeding worms and appears unperturbed by standard starvation treatments, regardless of duration of starvation or animal developmental stage (Fig. 2A-D). These observations suggest that EFK-1 is constitutively active in C. elegans fed a standard E. coli diet. Interestingly, EFK-1 overexpression, which substantially increased both efk-1 mRNA and protein expression, did not discernably increase EEF-2 phosphorylation past the observed baseline, posing the possibility that EEF-2 phosphorylation is saturated at baseline.

Besides absence of p-EEF-2 induction, other lines of evidence strongly suggest that p-EEF-2 is dispensable for starvation resistance. We showed that both catalytically inactive EFK-1 and EEF-2 phosphosite mutations, which abolish EEF-2 phosphorylation, led to no starvation defect compared to wild type. These results confirm that EEF-2 phosphorylation is not a critical regulatory event in the C. elegans starvation stress response, which contrasts with the important role of the eEF2K–eEF2 axis in mammalian cells under nutrient deprivation⁵. Indeed, the starvation resistance of efk-1 kinase-dead mutants implies that efk-1 kinase activity is altogether dispensable in this context.

The inability to phosphorylate EEF-2 does not lead to discernable physiological defects, as observed in both efk-1 null and kinase-dead mutant animals. However, mutation of the T56 phosphorylation site to alanine caused a germline defect in adult animals. Given that EEF-2 is essential and loss of EEF-2 leads to early larval lethality⁵², this implies that the threonine-to-alanine substitution may lead to a mild perturbation of EEF-2 activity in addition to lack of T56 phosphorylation. The precise structural change that underlies this specific phenotype remains to be defined.

Our data show that efk-1 exerts influence on gene expression via rewiring transcription, specifically through the downstream transcription factors zip-2 and cep-1. However, it is unclear how efk-1 signals to these transcription factors. We did not observe an efk-1-dependent increase in zip-2 and cep-1 transcript levels in starved worms (Supplementary Data 1), suggesting that their activity may be regulated on a translational or post-translational level. EFK-1 may regulate the expression of downstream factors via selective translation without affecting the global translation rate, which has been observed in mammalian cells and C. elegans under physiological stress conditions^1,78. Alternatively, EFK-1 might directly interact with these transcription factors through an unknown mechanism. EFK-1 may employ either or both of these modes of regulation to activate the downstream stress response pathway.

Our data revealed tissue-specific expression and activity of efk-1 in starvation resistance. Re-expression of efk-1 in the hypodermis rescued the starvation defects of the efk-1 mutant; this is unsurprising given the emerging role of the hypodermis in metabolism^79,80. Intriguingly, despite the neuronal presence of efk-1, neuronal overexpression of efk-1 led to a mild baseline growth defect and was unable to rescue the starvation phenotype. This is consistent with studies in murine models identifying neuronal eEF2K overexpression as detrimental to cognitive function⁸¹, suggesting that efk-1 may play a distinct role in neurons. We did not examine tissues with lower efk-1 expression such as the intestine, although this does not rule out possible function in these sites. Further tissue-specific involvement of efk-1 in stress signaling may be explored in future studies.

Taken together, our evidence points to a noncanonical, kinase activity-independent and EEF-2 phosphorylation-independent mechanism of EFK-1 in C. elegans starvation adaptation. To date, most studies on eEF2K have been conducted in transformed mammalian cell lines, which are fundamentally different from whole organisms such as C. elegans. It is unknown why C. elegans efk-1 employs a distinct mechanism, whether due to evolutionary divergence, differences between transformed and non-transformed cells, or tissue-specific effects. Our study has uncovered hitherto unknown regulatory roles of EFK-1 that may prove valuable to understanding how eEF2K regulates stress responses in different biological and evolutionary contexts.

Methods

C. elegans strains and maintenance

We cultured C. elegans strains using standard techniques on nematode growth media-lite (NGM-l) plates⁸². Each mutant was crossed into our lab N2 strain at least six times to remove background mutations. To prevent the accumulation of germline mutations, DNA repair mutants were frozen immediately after the final backcross, and an aliquot was thawed and maintained for no more than four generations prior to each experiment. E. coli OP50 was the standard food source; HT115 was used for RNAi experiments, and streptomycin-resistant OP50-1 was used for the starvation-primed UV recovery assay. All experiments were carried out at 20 °C unless otherwise specified. Worm strains used in this study are listed in Supplementary Data 5.

For synchronized worm growths, we isolated embryos by standard sodium hypochlorite treatment, and residual hypochlorite was removed by washing 3 times in M9⁸³. Isolated embryos were kept overnight in sterile M9 or on unseeded NGM-l plates until all hatched and arrested at the L1 stage via short-term fasting (16–24 hr). Synchronized L1 stage larvae were then transferred to seeded plates and grown to the desired stage.

Standard genetic crossing techniques were used to construct double mutants; genotyping primers are indicated in Supplementary Data 5. We attempted to construct the efk-1(ok3609);cebp-2(tm5421) double mutant, but failed to find homozygous double mutant animals from the clonal progeny of 100 individual candidate F2 worms, suggesting that loss of both genes causes synthetic lethality.

Feeding RNAi was performed on NGM-l plates supplemented with 25 μg/ml carbenicillin (BioBasic CDJ469), 1 mM IPTG (Santa Cruz CAS 367-93-1), and 12.5 μg/ml tetracycline (BioBasic TB0504; NGM-l-RNAi plates), and seeded with appropriate HT115 RNAi bacteria. The RNAi clones were from the Ahringer library (Source BioScience) and were sequenced prior to use.

Generation of transgenic strains

To generate the transgenic efk-1p::EFK-1::GFP reporter strain STE168, we used Phusion Hot Start II DNA Polymerase (Thermo Scientific F549L) to amplify 2000 bp upstream of efk-1 start codon, and the efk-1 genomic sequence including introns but without the final stop codon. efk-1 promoter and genomic sequence were cloned into Zero Blunt™ TOPO™ vector (Invitrogen 450245), transformed into TOPO One Shot™ MAX Efficiency DH5α-T1^R E. coli (Invitrogen 12297016), followed by restriction digestion and insertion into pSM vector⁸⁴ between restriction sites HindIII / AscI, and AscI / KpnI, respectively. 50 ng/µl of each plasmid and 25 ng/μl of co-injection marker odr-1p::RFP were microinjected into the gonad of day-one adult worms, and F2 were screened for stable transmission. We found a random integrant at 100% transmission and validated that both GFP and marker segregated in Mendelian proportions over three generations. This was crossed into efk-1 null background to generate the strain STE169.

For tissue-specific efk-1 rescue strains STE173, STE174, STE175, and STE176, we used Phusion Hot Start II DNA Polymerase (Thermo Scientific F549L) to amplify published semo-1 and rgef-1 promoter sequences⁸⁵ with flanking SphI / SgsI and NotI / SgsI sites, respectively. Resulting DNA fragments were used to generate semo-1p::EFK-1::GFP and rgef-1p::EFK-1::GFP plasmids, which were co-injected into C. elegans alongside the odr-1p::RFP marker as described above. Stable lines with >50% transmission were kept. These were crossed into the efk-1 null background to generate strains STE177, STE178, STE179, and STE180.

To generate cep-1p::CEP-1::3xFLAG overexpression lines STE181 and STE182, we used Phusion PCR to amplify the cep-1 promoter + gene fragment (up to the first codon of the previous gene) from genomic DNA template with primers bearing SphI / KpnI restriction sites. Resulting DNA fragments were purified, digested, and ligated into pSM vector as described. 3xFLAG was cloned from pJW1354 ligated into pSM between EcoRI / KpnI restriction sites, replacing the GFP sequence. The plasmid was co-injected with the odr-1p::RFP marker. Stable lines with >50% transmission were kept. These were crossed into the efk-1 null background to generate strains STE183 and STE184.

Generation of mutants with CRISPR-Cas9 genome editing

We used CRISPR-Cas9 genome editing to generate the EEF-2 phosphosite-dead (T56A) strain STE192, kinase-dead (D257A) efk-1 lines STE193 and STE194, endogenously tagged efk-1::GFP lines STE188 and STE189, and efk-1;myo-3::GFP(mit) lines STE171 and STE172, which bear the same deletion present in the efk-1(ok3609) strain. We directly injected respective gRNA/Cas9 complexes (Cas9: IDT #1081059; tracrRNA: IDT #1072534) with the homology-directed repair (HDR) donor oligos, as described⁸⁶. myo-2p::mCherry was used as co-injection marker. eef-2 T56A mutants were homozygous lethal and were maintained as heterozygotes by crossing in the hT2 balancer. For tagging GFP onto the endogenous efk-1 locus, we replicated the design of STE168 in the endogenous efk-1 locus, using Phusion PCR to amplify the linker+GFP sequence from pSM with primers bearing 60-bp homology arms. The resulting products were gel-purified and injected as HDR repair templates. For recreating the ok3609 deletion in the myo-3::GFP(mit) strain, we amplified the surrounding region of the ok3609 mutation and injected the gel-purified product as HDR repair template. After injection, F1s bearing the co-injection marker were screened for heterozygous edits using PCR or restriction fragment length polymorphism (RFLP). Homozygous offspring of each line were confirmed by Sanger sequencing. gRNAs, HDR donor oligos, and genotyping primers are listed in Supplementary Data 5.

C. elegans starvation studies

Worms were maintained for at least two generations in a fed state prior to all starvation experiments. For L4 studies, we synchronized worms by hatching in M9 overnight (16 ~ 24 h), followed by growth on seeded 100 mm NGM-l plates to mid-L4 at a density of 2000-2200 worms/plate. The fed portion was collected at mid L4 stage as judged by vulval morphology developmental staging. The starved portion was first washed 5 times in sterile M9, and then placed either on unseeded plates or suspended in sterile M9 for the number of hours specified. For L1 studies, worms were synchronized and grown on seeded plates to gravid adulthood at a density of 1000–1200 worms/plate. Worms were washed off the plate and residual embryos were collected with a cell scraper (Falcon #353085). For Western blot and RT-qPCR studies, bulk embryos were isolated by vigorous mixing in hypochlorite solution (40 ml dH₂O, 7 ml sodium hypochlorite Sigma #425044, 3 ml 1 M KOH) for 6-7 minutes, hatched overnight without food, and subjected to starvation treatments as specified. For imaging and mitochondrial DNA content studies, worms were bleached for 2 minutes, and excess carcasses were removed by filtering through a 35-micron nylon mesh (Elko Filtering #03-35/16).

L1 starvation survival assays were adapted from a previous study⁸⁷. Synchronized embryos were extracted by hypochlorite treatment for 2 minutes and resuspended in S‐basal medium without cholesterol at a concentration of ~1 worm/μl and supplemented with an antibiotic–antimycotic mix (Gibco #15240062). Worms were starved at 20 °C while being continuously rotated on a tube rotator (ThermoFisher) at 20 rpm. To assess viability, ~150 animals were transferred to seeded NGM-l plates every 2-3 days and assessed for growth to the L4 stage after 48 h at 20 °C, with day 0 being counted as 16-24 h after hypochlorite treatment. For transgenic strains with extrachromosomal arrays, odr-1p::RFP worms and non-transgenic siblings from two independent lines were studied under an upright M205FA fluorescent microscope (Leica). Statistical significance of three to four independent biological repeats was calculated using area under the curve and one-way ANOVA with Tukey’s or Dunnett’s multiple comparisons test.

For starvation studies with chemical treatments, CHX (Sigma #1810), puromycin (Sigma #P8833), NAC (Sigma #A7250), VitC (Sigma #A92902), and mitoQ (MedChemExpress #HY-100116A) stock solutions were prepared freshly in S-basal at concentrations of 10 mM, 10 mM, 100 mM, 100 mM, and 5 mM, respectively. Each chemical was filtered through a 0.2 µm filter and added to the culture at specified concentrations. To compensate for the degradation of NAC in an aqueous solution during L1 starvation survival experiments⁸⁸, we added fresh NAC at 40% of the original amount on day 6. Vitamin C containing solutions were kept in the dark in aluminum foil to minimize photodegradation⁸⁹.

A549 starvation studies

A549 cells were obtained from Dr. Poul Sorensen. For Western blot experiments, 2 × 10⁵ cells were seeded in 6-well plates and grown overnight in Dulbecco’s modified Eagle’s medium (DMEM) (Gibco #11995073) supplemented with 10% fetal bovine serum (FBS) (Gibco #12483020) at 37 °C under 5% CO₂ in a humidified incubator. The media was aspirated, and the fed condition was incubated in 10% FBS DMEM, while the starved condition was incubated in Hanks’ Balanced Salt Solution (HBSS) (Gibco #14175095) for 3, 6, and 24 h. Total cellular protein was extracted using RIPA lysis buffer supplemented with protease inhibitors and phosphatase inhibitors.

Growth rate assays

We measured the growth rate for every strain used in the L1 starvation survival assay. Staged L4 worms were picked onto a seeded plate and grown to gravid adults overnight at RT. 10 gravid adults were picked onto a fresh plate and allowed to lay eggs for 2 h at room temperature. Parent animals were removed and synchronized F1 were incubated in 20 °C for 56 h. The portion of F1 that were able to reach L4 stage was recorded. Statistical analysis was performed using one-way ANOVA with Dunnett’s multiple comparisons test.

UV sensitivity assays

UV sensitivity assays were adapted from previous studies^61,90. For the somatic NER assay, L1 worms were synchronized overnight on unseeded NGM-l plates, then irradiated uncovered in a Stratalinker 2400 UV Crosslinker (Stratagene) with wavelength 254 nm light at specified dosages. Worms were placed at 20 °C for 48 h and scored for recovery to L4 stage. Recovery rates were normalized to no UV control of the same genotype. Statistical analysis was performed using two-way ANOVA with Dunnett’s multiple comparisons test.

For the germline NER assay, embryos were extracted by hypochlorite treatment and placed on seeded NGM-l plates at 20 °C for 72 h until young adulthood. Worms were collected, washed three times in M9, and placed on unseeded NGM-l plates. Worms were allowed to disperse for an hour and then irradiated uncovered in the Stratalinker 2400 UV Crosslinker at specified dosages. Immediately following UV exposure, 15-20 staged young adults were picked onto a seeded NGM-l plate and allowed to lay eggs for 4 h. The proportion of eggs that hatched after 24 h was recorded. Embryonic lethality was normalized to no UV control of the same genotype. For each dosage, statistical significance between mutants was computed using two-way ANOVA with Dunnett’s multiple comparisons test.

For testing NER activation after exposure to starvation, worms were starved in S-basal for the specified number of days as described above. At each timepoint, a portion of worms were placed on an unseeded 60 mm plate and allowed to disperse for an hour. Uncovered plates were exposed to UV as above. After irradiation, worms were quantified, placed on a seeded OP50-1 plate for recovery at 20 °C for 48 h, and scored for recovery to L4 stage. Statistical significance of three independent biological repeats was calculated using area under the curve and one-way ANOVA with Tukey’s multiple comparisons test.

Respiration and motility measurement

Measurement of oxygen consumption rates (OCR) in live C. elegans was conducted with the Resipher system (Lucid Scientific) as described⁹¹. Embryos from wild-type, efk-1(ok3609), efk-1::GFP, and efk-1;efk-1::GFP worms were isolated by hypochlorite treatment and placed on bacteria-free NGM plates for L1 synchronization. A 96-well microplate was seeded with synchronized L1 worms at a density of 100 worms in 100 µL of M9 buffer per well and fitted with the Resipher sensor lid. The Resipher device continuously recorded OCR data for 101 h with timepoints taken every 15 minutes. Afterwards, OCR rates in each well were normalized to the number of worms in that well as determined by visual inspection. OCR data normalized per worm were averaged across technical replicates (6-12 wells) for each time point in each of 3-4 biological replicates. Time-course data for the 3-4 biological replicates was analyzed by a global area under the curve (AUC) analysis in GraphPad Prism 10. The computed AUC values and associated standard errors were compared across the four groups using one-way ANOVA with Tukey’s multiple comparison test.

C. elegans motility was recorded continuously for 40 h with the wMicrotracker (Phylum Tech). Wild-type, efk-1(ok3609), efk-1::GFP, and efk-1;efk-1::GFP embryos were isolated and synchronized as for the Resipher analysis. A 96-well microplate was seeded with synchronized L1 worms at a density of 100 worms in 100 µL of M9 buffer in each well. Movement data was recorded every hour. Afterwards, motility rates were normalized to the number of worms present in each well as determined by visual inspection. Similar to the Resipher analysis, motility data was normalized per worm were averaged across technical replicates (6-12 wells) for each time point in each of 3 biological replicates. Statistical analysis was performed as for the Resipher data.

SDS-PAGE and western blotting

For C. elegans studies, synchronized mid-L4 worms or L1 worms were collected, rapidly washed once with M9 to remove excess bacteria, and flash frozen in ethanol and dry ice. Worm pellets were sonicated in 1 ml ice-cold Protein Lysis Buffer (150 mM KCl, 1 mM EDTA, 0.25% SDS, 1.0% NP-40, 50 mM Tris/HCl pH=7.4) supplemented with 1 mM PMSF, 1x Roche cOmplete™ Protease Inhibitor Cocktail (#4693116001, Roche), 50 mM NaF and 1 mM sodium orthovanadate. For A549 cell studies, cells were washed with cold phosphate-buffered saline (PBS) twice and lysed with 400 µl RIPA buffer (150 mM NaCl, 50 mM Tris/HCl pH=7.4, 1.0% NP-40, 0.5% DOC, 0.1% SDS) supplemented with 1.15 mM PMSF, 1 x Roche cOmplete^TM Protease Inhibitor Cocktail, 50 mM NaF, and 1 mM sodium orthovanadate. Protein concentrations were determined using the RC DC Protein Assay kit (#500-0121, Bio-Rad) against a BSA standard curve, and 5 µg of protein was loaded in each well. SDS-PAGE analysis and immunoblotting were performed as described⁴². For C. elegans studies, primary antibodies were diluted in 2.5% skimmed milk as follows: 1:1000 anti-rabbit phospho-eEF2 (T56) antibody (#2331, Cell Signaling Technologies), 1:1000 anti-rabbit eEF2 antibody (#2332, Cell Signaling Technologies), 1:500 anti-rabbit phospho-eIF2α (Ser51) antibody (#9721, Cell Signaling Technologies), 1:1000 anti-mouse GFP antibody (#11814460001, Roche), 1:1000 anti-mouse α-tubulin antibody (#T9026, Sigma), or 1:1000 anti-mouse puromycin antibody (#MABE343, Millipore). For A549 cell line studies, primary antibodies were diluted in 5% skimmed milk as above, except for 1:1000 anti-rabbit eEF2K antibody (#3692, Cell Signaling Technologies), and α-tubulin which was used at 1:5000 concentration. Anti-rabbit (#7074, NEB) and anti-mouse (#7076, Cell Signaling Technologies) horseradish peroxidase (HRP) conjugated secondary antibodies were diluted at 1:5000 in 5% skim milk. Detection was done using ECL (#32109, Pierce) in a Vilber Fusion FX chemiluminescence imager. Quantification of relative signal intensity was done using ImageJ. Mean value of each band was measured with a selection of the same area, inverted, and corrected for background. Relative signal intensity was obtained by dividing the background-corrected intensity of the protein bands with that of the tubulin loading control or EEF-2, as appropriate. Statistical significance was calculated using one-way ANOVA or two-way ANOVA with the statistical test specified in Source Data.

Puromycin incorporation assay (SUnSET)

SUnSET was adapted from previous studies^27,33. For C. elegans studies, synchronized L4 or L1 worms were starved for the time specified. CHX controls were treated with 2 mg/ml CHX (Sigma #1810) added 6 h prior to harvest. All samples were incubated with 0.5 mg/ml puromycin for 3 h before harvest. Worm pellets were washed once in M9 and flash frozen. For A549 cell line studies, 2×10⁵ cells were fed or starved 6 h as above. CHX controls were treated with 2 µg/ml CHX 1 h before harvest. All samples were supplemented with 1 µg/ml puromycin 10 minutes before harvest. Statistical significance was calculated using two-way ANOVA with uncorrected Fisher’s LSD.

DIC and fluorescence microscopy

For efk-1::GFP images, synchronized mid-L4 efk-1::GFP worms were immobilized in 12.5 mM levamisole (Sigma L9756) on 2% (w/v) agarose pads. Images were captured at 40x magnification on a Hamamatsu ORCA-Flash4.0 LT+ Digital CMOS camera attached to a Leica SP8X confocal microscope.

To analyze fluorescence in the irg-1p::GFP reporter line, synchronized L1s were placed onto NGM-l plates seeded with OP50 or RNAi plates seeded with the appropriate HT115 RNAi culture. irg-1p::GFP worms were kept at 20 °C for 48 h until they reached mid L4-stage. For imaging L1 stage larvae, synchronized animals were starved in S-basal medium, and an aliquot was taken at each timepoint and filtered through a 35-micron nylon mesh (Elko Filtering #03-35/16). Worms were mounted and immobilized as above, and images were captured at 10x or 20x magnification on a CoolSnap HQ camera (Photometrics) attached to a Zeiss AxioPlan 2 compound microscope. Images were captured at 300 ms exposure time for GFP.

ZIP-2::GFP localization was assessed using the ERT201 strain. L4 or L1 larvae were synchronized and prepared as above, and GFP images of the first two intestinal cells were captured at 100x magnification on a CoolSnap HQ camera (Photometrics) attached to a Zeiss AxioPlan 2 compound microscope at 300 ms. Images were scored for GFP localization in the first intestinal cells using the following categories: cytoplasmic (no discernable nuclear signal), intermediate (nuclear and cytoplasmic signal roughly comparable), and nuclear (strong nuclear and weak cytoplasmic signal). Statistical significance was calculated using one-way ANOVA with Dunnett’s multiple comparisons test.

ROS staining

Synchronized L1 worms were starved in S-basal as above. At each timepoint of starvation, an aliquot was treated with either DHE or CellROX Green, as described^64,65. For DHE, worms were washed 3 times in M9, incubated with 10 μM DHE (Sigma #D7008) for 30 min on a tube rotator, washed 3 times with PBS, and mounted onto 2% agarose pads in 12.5 mM levamisole for imaging. For CellROX Green, worms were washed 3 times in M9, fixed in 2% formaldehyde (Sigma #F8775) for 30 min, washed 3 times with M9 with 0.01% Tween, incubated with 5 μM CellROX Green for 1 h, washed 3 times with PBS and mounted onto 2% agarose pads for imaging. L1 worms were imaged using the 20x objective. Images were captured at 300 ms exposure time for CellROX Green and 100 ms for DHE. For each experiment, 3 independent biological repeats were performed with 90 ~ 110 live worms quantified per timepoint. Images were analyzed using ImageJ software (https://imagej.nih.gov/ij/download.html). Normalized whole-worm fluorescence was quantified by measuring the whole-body fluorescence of worms minus the average of 3 spots of background taken on the same image. For each timepoint, statistical significance between conditions was calculated using one-way ANOVA with Tukey’s multiple comparisons test.

Mitochondrial morphology imaging

Synchronized L1 worms with myo-3::GFP(mit) were starved as described above. At each timepoint of starvation, an aliquot of L1 worms was filtered through a 35-micron nylon mesh (Elko Filtering #03-35/16), mounted on 2% agarose pads and immobilized with 50 mM levamisole. Mitochondrial images were captured using a SP8X STED White Laser Confocal Microscope (Leica) with a 63x objective using the Leica Application Suite X (LAS X). For each worm, Z-stack images were taken at 0.5 μm intervals using a 488 nm laser. The laser power, intensity and gain were kept consistent between strains and conditions within each experimental replicate (N = 3 replicates with 10 worms per replicate for each strain). Images were processed using LAS X and scored for changes in the muscle mitochondrial morphology of animals using the following categories: tubular (minimal to no fragmentation), intermediate (some fragmentation), and fragmented (majority of network is fragmented), as described⁹². Statistical significance was calculated using two-way ANOVA with Dunnett’s multiple comparisons test.

RNA sequencing and GO term analysis

Synchronized L1 wild-type, efk-1(ok3609), zip-2(tm4248), and cep-1(gk138) worms were allowed to grow on OP50 plates to mid-L4 stage. Fed samples were collected immediately by flash-freezing. Starved samples were washed 5 times in sterile M9, starved for 8 h on unseeded plates, and then collected. RNA was isolated from whole worms as described⁹³. RNA integrity and quality were ascertained on a BioAnalyzer. Construction of strand-specific mRNA sequencing libraries and sequencing (75 bp PET) on an Illumina HiSeq 2500 machine was done at the Sequencing Services facility of the Genome Sciences Centre, BC Cancer Agency, Vancouver BC, Canada (https://www.bcgsc.ca/services/sequencing-services). We sequenced >20 million reads per sample. The raw FASTQ reads obtained from the facility were trimmed using Trimmomatic version 0.36⁹⁴ with parameters LEADING:3 TRAILING:3 SLIDINGWINDOW:4:15 MINLEN:36. Next, the trimmed reads were aligned to the NCBI reference genome WBcel235 WS277 (https://www.ncbi.nlm.nih.gov/assembly/GCF_000002985.6/) using Salmon version 0.9.1⁹⁵ with parameters -l A -p 8 –gcBias. Then, transcript-level read counts were imported into R and summed into gene-level read counts using tximport⁹⁶. Genes not expressed at a level greater than 1 count per million (CPM) reads in at least three of the samples were excluded from further analysis. The gene-level read counts were normalized using the trimmed mean of M‐values (TMM) in edgeR⁹⁷ to adjust samples for differences in library size. Differential expression analysis was performed using the quasi-likelihood F-test with the generalized linear model (GLM) approach in edgeR⁹⁷. RNA-seq data have been deposited at NCBI Gene Expression Omnibus (https://www.ncbi.nlm.nih.gov/geo/) under the record GSE259223. Functional enrichment analysis and visualization were performed using easyGSEA and easyVizR in the eVITTA toolbox (https://tau.cmmt.ubc.ca/eVITTA/; input September 19, 2023)⁹⁸.

For the reanalysis of published starvation RNA-seq data²⁰, raw reads were downloaded from Sequence Read Archive (SRA) accession PRJNA386884, extracted using fastq-dump, and processed as above. Differential expression analysis was performed as described above. Venn diagrams of gene set overlaps and associated p-value was generated as described⁹⁹.

RNA isolation and qRT-PCR analysis

RNA isolation was performed as previously described⁹³. 2 μg total RNA was used to generate cDNA with Superscript II reverse transcriptase (Invitrogen 18064-014), random primers (Invitrogen 48190-011), dNTPs (Fermentas R0186), and RNAseOUT (Invitrogen 10777-019). Quantitative real-time PCR was performed in 10 μl reactions using PowerTrack™ SYBR Green Master Mix (Applied Biosystems A46109), 1:10 diluted cDNA, and 5 μM primer, and analyzed with an Applied Biosystems QuantStudio 5 Real-Time PCR System. We analyzed the data with the ΔΔCt method. For each sample, we calculated normalization factors by averaging the (sample expression)/(average wild type expression) ratios of three normalization genes, act-1, tba-1, and ubc-2. Average relative transcript abundance of normalization genes was kept at a constant across repeats to correct for batch variations in transcript abundance. Statistical significance was calculated using unpaired t-test or one-way ANOVA with Dunnett’s multiple comparisons test. All data originate from three or more independent biological repeats, and each PCR reaction was conducted in technical triplicate. RT-qPCR Primers were tested on serial cDNA dilutions and analyzed for PCR efficiency prior to use. Sequences of qRT-PCR primers are listed in Supplementary Data 5.

Mitochondrial DNA content assay

Mitochondrial DNA content was measured using a qPCR-based assay as described¹⁰⁰. Synchronized L1 worms were starved as above. For each timepoint, 300 worms were lysed in 30 µl lysis buffer as described, and 1 µl of 1:10 diluted lysate was used per 10 µl qPCR reaction performed in quadruplicates. Statistical analysis was performed using two-way ANOVA with Tukey’s multiple comparisons test. Primers that amplify mtDNA and a noncoding nuclear region were from a published study¹⁰⁰ and listed in Supplementary Data 5.

Statistics and reproducibility

All experiments were performed at least three times with independent biological repeats consisting of C. elegans or cancer cells grown at different times. Data was visualized using Prism (version 10) or R, and individual biological repeats are presented whenever possible. Statistical analysis was performed using Prism (version 10). p < 0.05 was considered statistically significant. Raw data and detailed statistical results are listed in the Source Data file. No statistical method was used to predetermine sample size. No data were excluded from the analyses, except for ROS staining and mitochondrial morphology experiments, where only live worms were quantified due to autofluorescence from dead carcasses.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability

The authors declare that the data underlying the findings of this study are available within the paper and its Supplementary Information files. Source data underlying the Figs. and Supplementary Figs. are provided in a Source Data file. Source data are provided with this paper. The RNA-seq transcriptomics data generated in this study have been deposited in the Gene Expression Omnibus database under accession code GSE259223. Source data are provided with this paper.

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Acknowledgements

We thank Dr. E. R. Troemel (UCSD) and Dr. Alberto Delaidelli (BC Cancer Agency) for providing invaluable suggestions and advice, and Taubert lab members for comments on the manuscript. We thank Rachel Cheng and Yuchang Zhan for generating C. elegans strains, Dr. Kota Mizumoto and Menghao Lu (UBC) for help with generating transgenic and CRISPR-Cas9 genome-edited strains, Dr. Kelsie R. S. Doering (Kwantlen Polytechnic University) for assistance with mitochondrial imaging, Dr. E. R. Troemel (UCSD) for providing backcrossed zip-2 and cebp-2 strains, Dr. N. J. O’Neil (UBC) for providing backcrossed xpa-1, ercc-1, and xpf-1 mutant strains, and Dr. Veena Prahlad (Roswell Park Comprehensive Cancer Center) for proving an integrated cep-1 overexpression strain. Some strains were provided by the CGC, which is funded by NIH Office of Research Infrastructure Programs (P40 OD010440). Grant support was from The Canadian Institutes of Health Research (CIHR; PJT-153199 and PJT-186144 to ST; FDN-143280 to PHS), the Natural Sciences and Engineering Research Council of Canada (NSERC; RGPIN-2018-05133, RGPIN-2024-06537, and ALLRP 597082-24 to ST), the British Columbia Cancer Foundation through generous donations from Team Finn and other riders in the Ride to Conquer Cancer (to PHS), and a research grant from the National Institutes of Health (R00 ES029552 to JHH). JY was supported by BCCHR and UBC scholarships, CS by BCCHR and UBC scholarships, GE by a UBC scholarship and a CIHR CGS-D scholarship, PB by an NSERC undergraduate student research award (USRA), DS by a UBC Faculty of Medicine Summer Student Research Program (FoM SSRP) studentship, and ST by a Canada Research Chair and a BCCHR Investigator Grant Award Program (IGAP) award.

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Forum Bhanshali
Present address: Catalera BioSolutions, 199 W 6th Ave, Vancouver, BC, V5Y 1K3, Canada
Xuanjin Cheng
Present address: Canada’s Michael Smith Genome Sciences Centre, 570 W 7th Ave, Vancouver, BC, V5Z 4S6, Canada

Authors and Affiliations

Centre for Molecular Medicine and Therapeutics, The University of British Columbia, 950 W 28 th Ave, Vancouver, BC, V5Z 4H4, Canada
Junran Yan, Forum Bhanshali, Chiaki Shuzenji, Shane K. B. Taylor, Glafira Ermakova, Xuanjin Cheng, Pamela Bai, Gahan Diwan, Donna Seraj & Stefan Taubert
Edwin S.H. Leong Centre for Healthy Aging, The University of British Columbia, 117-2194 Health Sciences Mall, Vancouver, BC, V6T 1Z3, Canada
Junran Yan, Chiaki Shuzenji, Shane K. B. Taylor, Glafira Ermakova, Pamela Bai & Stefan Taubert
British Columbia Children’s Hospital Research Institute, 950 W 28th Ave, Vancouver, BC, V5Z 4H4, Canada
Junran Yan, Forum Bhanshali, Chiaki Shuzenji, Shane K. B. Taylor, Glafira Ermakova, Xuanjin Cheng, Pamela Bai, Gahan Diwan, Donna Seraj & Stefan Taubert
Graduate Program in Cell & Developmental Biology, The University of British Columbia, 950 W 28th Ave, Vancouver, BC, V5Z 4H4, Canada
Junran Yan & Stefan Taubert
Department of Medical Genetics, The University of British Columbia, 950 W 28th Ave, Vancouver, BC, V5Z 4H4, Canada
Forum Bhanshali, Chiaki Shuzenji, Shane K. B. Taylor, Glafira Ermakova, Xuanjin Cheng, Pamela Bai, Donna Seraj & Stefan Taubert
Department of Biochemistry and Molecular Biology, Medical University of South Carolina, 173 Ashley Ave, Charleston, SC, 29425, USA
Tsultrim T. Mendenhall & Jessica H. Hartman
Department of Biology, University of British Columbia, Vancouver, BC, V6T 1Z4, Canada
Gahan Diwan
Nicholas School of the Environment, Duke University, Durham, NC, 27708-0328, USA
Joel N. Meyer
Department of Pathology and Laboratory Medicine, University of British Columbia, 675 W 10th Ave, Vancouver, BC, V6T 1Z4, Canada
Poul H. Sorensen
Department of Molecular Oncology, BC Cancer Research Institute, 675 W 10th Ave, Vancouver, BC, V5Z 1L3, Canada
Poul H. Sorensen

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Junran Yan
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Forum Bhanshali
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Design of research: J.Y., F.B., C.S., J.N.M., P.H.S., J.H.H., S.T. Performed experiments: J.Y., F.B., C.S., T.T.M., S.K.B.T., G.E., P.B., G.D., D.S., J.H.H., S.T. Analysed data: J.Y., F.B., T.T.M., S.K.B.T., X.C., J.Y., F.B. Wrote the manuscript: J.Y. and S.T. Edited the manuscript: All authors.

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Correspondence to Stefan Taubert.

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JHH is a paid consultant for Surrozen, Inc. All other authors declare no competing interests.

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Yan, J., Bhanshali, F., Shuzenji, C. et al. Eukaryotic Elongation Factor 2 Kinase EFK-1/eEF2K promotes starvation resistance by preventing oxidative damage in C. elegans. Nat Commun 16, 1752 (2025). https://doi.org/10.1038/s41467-025-56766-1

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Received: 26 February 2024
Accepted: 29 January 2025
Published: 18 February 2025
DOI: https://doi.org/10.1038/s41467-025-56766-1

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Abstract

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Introduction

Results

efk-1(ok3609) is a null allele of efk-1

Starvation does not increase EEF-2 phosphorylation

EFK-1 promotes starvation resistance without altering global translation rate

EFK-1 promotes starvation resistance independently of its kinase activity and EEF-2 phosphorylation

Transcription factors ZIP-2 and CEP-1 function in the efk-1 starvation response pathway

DNA repair pathways are transcriptionally upregulated by efk-1, zip-2, and cep-1 during starvation

NER is required for starvation survival and acts in the efk-1 pathway

efk-1 and cep-1 play a minor role in somatic and germline NER

BER is required for starvation survival and acts in the efk-1 pathway

efk-1 protects against elevated ROS during starvation

efk-1 protects against mitochondrial ROS

Discussion

efk-1, cep-1, and zip-2 promote genome integrity in the same starvation resistance pathway

efk-1 protects against mitochondrial ROS

EFK-1 functions independently of EEF-2 phosphorylation and global translation regulation

Methods

C. elegans strains and maintenance

Generation of transgenic strains

Generation of mutants with CRISPR-Cas9 genome editing

C. elegans starvation studies

A549 starvation studies

Growth rate assays

UV sensitivity assays

Respiration and motility measurement

SDS-PAGE and western blotting

Puromycin incorporation assay (SUnSET)

DIC and fluorescence microscopy

ROS staining

Mitochondrial morphology imaging

RNA sequencing and GO term analysis

RNA isolation and qRT-PCR analysis

Mitochondrial DNA content assay

Statistics and reproducibility

Reporting summary

Data availability

References

Acknowledgements

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Authors and Affiliations

Contributions

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Competing interests

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Peer review information

Additional information

Supplementary information

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