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. 2023 Oct 2;42(22):e114093. doi: 10.15252/embj.2023114093

Thiol reductive stress activates the hypoxia response pathway

Ravi 1, Ajay Kumar 2, Shalmoli Bhattacharyya 2, Jogender Singh 1,✉
PMCID: PMC10646554  PMID: 37902464

Abstract

Owing to their capability to disrupt the oxidative protein folding environment in the endoplasmic reticulum (ER), thiol antioxidants, such as dithiothreitol (DTT), are used as ER‐specific stressors. We recently showed that thiol antioxidants modulate the methionine‐homocysteine cycle by upregulating an S‐adenosylmethionine‐dependent methyltransferase, rips‐1, in Caenorhabditis elegans. However, the changes in cellular physiology induced by thiol stress that modulate the methionine‐homocysteine cycle remain uncharacterized. Here, using forward genetic screens in C. elegans, we discover that thiol stress enhances rips‐1 expression via the hypoxia response pathway. We demonstrate that thiol stress activates the hypoxia response pathway. The activation of the hypoxia response pathway by thiol stress is conserved in human cells. The hypoxia response pathway enhances thiol toxicity via rips‐1 expression and confers protection against thiol toxicity via rips‐1‐independent mechanisms. Finally, we show that DTT might activate the hypoxia response pathway by producing hydrogen sulfide. Our studies reveal an intriguing interaction between thiol‐mediated reductive stress and the hypoxia response pathway and challenge the current model that thiol antioxidant DTT disrupts only the ER milieu in the cell.

Keywords: C. elegans, endoplasmic reticulum, hif‐1, hypoxia, reductive stress

Subject Categories: Metabolism, Signal Transduction

Unbiased genetic screens reveal that thiol antioxidant DTT affects not only the ER milieu but also activates hypoxia responses possibly via hydrogen sulfide production.

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Introduction

Maintenance of cellular homeostasis requires a balanced redox environment in the cell. Redox reactions result in the production of reactive oxygen species (ROS). High levels of ROS are associated with several pathological conditions (Sies & Jones, 2020). However, physiological ROS plays essential roles in cellular survival, differentiation, proliferation, repair, and aging (Trachootham et al, 2008; Sies & Jones, 2020). An increased amount of antioxidants results in the depletion of ROS, leading to reductive stress that is linked with several pathological conditions (Pérez‐Torres et al, 2017; Rajasekaran, 2020). Dietary supplementation of high amounts of antioxidants correlates with accelerated cancer progression and a higher incidence of cancer‐related mortality (Villanueva & Kross, 2012; Sayin et al, 2014). A high amount of dietary antioxidants accelerates aging in the nematode Caenorhabditis elegans (Gusarov et al, 2021). Despite the association of high amounts of antioxidants with pathological conditions, the full spectrum of physiological effects of reductive stress remains to be elucidated.

Thiols, which contain a sulfhydryl group, are a major defense mechanism in the cell against oxidative stress and undergo oxidation to form disulfides under oxidative conditions (Ulrich & Jakob, 2019). Alterations in thiol‐disulfide homeostasis play a role in several pathological conditions, including cardiovascular disease (Kundi et al, 2015), chronic renal disease (Rodrigues et al, 2013), and diabetes mellitus (Ates et al, 2016). However, the physiological effects of thiol reductive stress remain obscurely characterized. Because disulfide bonds are formed in the oxidative environment of the endoplasmic reticulum (ER), ER is expected to be the primary target of reductive stress (Cuozzo & Kaiser, 1999; Merksamer et al, 2008). Indeed, thiols, such as dithiothreitol (DTT), disrupt disulfide bonds in the ER, leading to protein misfolding (Braakman et al, 1992). The ensuing protein misfolding results in ER stress; therefore, thiol‐reducing agents are used as ER‐specific stressors (Oslowski & Urano, 2011). In addition to ER stress, thiol antioxidants can also result in oxidative stress by activating futile oxidative cycles in the ER (Maity et al, 2016). Recent studies in C. elegans showed that, besides causing ER stress, thiol antioxidants modulate the methionine‐homocysteine cycle (Gokul & Singh, 2022; Winter et al, 2022). The modulation of the methionine‐homocysteine cycle is a consequence of the upregulation of an S‐adenosylmethionine (SAM)‐dependent methyltransferase, rips‐1, by thiol antioxidants. Thus, reductive stress caused by thiols could affect cellular physiology beyond the ER. However, such physiological effects of thiol antioxidants remain to be fully characterized.

In this study, using C. elegans and human cell lines, we characterized the physiological effects of thiol reductive stress. Because thiol antioxidants result in the increased expression of rips‐1 (Gokul & Singh, 2022; Winter et al, 2022), we reasoned that understanding the regulation of rips‐1 expression might provide insights into the physiological effects of thiol reductive stress. To this end, we carried out forward genetic screens to understand the regulation of rips‐1 expression. The expression levels of rips‐1 correlated with the activity of the hypoxia response pathway. We observed that exposure to DTT resulted in the activation of a functional hypoxia response pathway that protected C. elegans against cyanide‐mediated fast killing on Pseudomonas aeruginosa PAO1. The hypoxia response pathway has rips‐1‐dependent and independent opposing roles in regulating thiol‐mediated toxicity. The activation of the hypoxia response pathway by DTT exposure is conserved in human cell lines. We showed that DTT produces hydrogen sulfide, which might be responsible for activating the hypoxia response pathway. These studies reveal an intriguing relationship between thiol reductive stress and the hypoxia response pathway.

Results

The hypoxia response pathway modulates rips‐1 expression levels

The thiol antioxidants, DTT and β‐mercaptoethanol, cause toxicity in C. elegans by upregulating a SAM‐dependent methyltransferase, rips‐1, that results in the modulation of the methionine‐homocysteine cycle (Gokul & Singh, 2022; Winter et al, 2022). Previous studies have shown that rips‐1 expression might be altered by hydrogen sulfide, mitochondrial dysfunction, and the hypoxia‐inducible factor (Miller et al, 2011; Nargund et al, 2012; Pender & Horvitz, 2018; Vora et al, 2022; Winter et al, 2022). However, the full spectrum of physiological changes induced by thiol reductive stress that results in rips‐1 upregulation remains to be discovered. Because thiol antioxidants activate the unfolded protein response (UPR) in the ER by enhancing protein misfolding, we asked whether the ER UPR pathways were involved in the upregulation of rips‐1 upon DTT exposure. To this end, we crossed a rips‐1p::GFP reporter strain with mutants of different ER UPR pathways, including ire‐1(v33), xbp‐1(tm2482), atf‐6(ok551), and pek‐1(ok275). The green fluorescent protein (GFP) levels were lower in the ire‐1/xbp‐1 and atf‐6 pathway mutants compared to the wild‐type animals in the absence of DTT (Fig EV1A and B). This could be because of compensatory protein translation inhibition by pek‐1 pathway due to increased ER stress in the ire‐1/xbp‐1 and atf‐6 pathway mutants (Harding et al, 2000). It is also possible that the ire‐1/xbp‐1 and atf‐6 pathways control the basal expression of rips‐1. Nevertheless, exposure to DTT resulted in the upregulation of GFP levels in all the ER UPR mutants (Fig EV1A and B), indicating that the ER UPR pathways are not involved in the DTT‐mediated rips‐1 upregulation. Recently, we showed that DTT also results in the activation of the mitochondrial UPR (Gokul & Singh, 2022). Therefore, we asked whether the mitochondrial UPR is required for DTT‐mediated upregulation of rips‐1. DTT exposure resulted in the upregulation of rips‐1p::GFP in atfs‐1(gk3094) animals (Fig EV1A and B), ruling out a role of mitochondrial UPR in the upregulation of rips‐1.

Figure EV1. The unfolded protein response (UPR) pathways in the endoplasmic reticulum (ER) and mitochondria do not control DTT‐mediated rips‐1 upregulation.

Figure EV1

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  1. Representative fluorescence images of rips‐1p::GFP in the background of WT, ire‐1(v33), xbp‐1(tm2482), atf‐6(ok551), pek‐1(ok275), and atfs‐1(gk3094) animals grown on 0 mM DTT until the young adult stage, followed by incubation on 0 or 10 mM DTT for 10 h. The red fluorescence in the pharynx region is from the myo‐2p::mCherry coinjection marker. Scale bar = 200 μm.
  2. Quantification of GFP levels of rips‐1p::GFP in the background of WT, ire‐1(v33), xbp‐1(tm2482), atf‐6(ok551), pek‐1(ok275), and atfs‐1(gk3094) animals grown on 0 mM DTT until the young adult stage, followed by incubation on 0 or 10 mM DTT for 10 h. ***P < 0.001 via the t‐test (n = 14 worms each). In the boxplots, the central bands represent the median value, the boxes represent the upper and lower quartile, and the whiskers represent the minimum and maximum values.
  3. Scheme for a forward genetic screen for mutants with constitutive upregulation of rips‐1p::GFP.

Source data are available online for this figure.

To characterize the physiological changes that trigger the upregulation of rips‐1, we designed a forward genetic screen to isolate mutants that had constitutive activation of rips‐1 (Fig EV1C). We obtained three independent mutants with enhanced GFP expression compared to the parental strain JSJ13 (rips‐1p::GFP) (Fig 1A and B). To identify the causative mutations, we performed whole‐genome sequencing of the mutants after backcrossing them six times with the parental strain. The sequenced genomes of mutants were aligned with the reference genome of C. elegans to identify variants. Evaluation of the protein‐coding genes having mutations in the isolated mutants revealed that all of the mutants had mutations in different negative regulators of the hypoxia response pathway (Fig 1C and Appendix Table S1). The JSJ14 mutant animals had two missense mutations in the gene rhy‐1, the JSJ15 mutant animals had a splice site acceptor mutation in the gene egl‐9, and the JSJ16 mutant animals had a premature nonsense mutation in the gene vhl‐1 (Fig 1C and Appendix Table S1). Because egl‐9(jsn15) and vhl‐1(jsn16) are likely loss‐of‐function alleles, these data suggested that inhibition of the negative regulators of the hypoxia response pathway may result in the enhanced expression of rips‐1. Indeed, the knockdown of egl‐9, rhy‐1, and vhl‐1 by RNA interference (RNAi) resulted in the upregulation of rips‐1p::GFP (Fig 1D and E).

Figure 1. The hypoxia response pathway modulates rips‐1 expression levels.

Figure 1

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  1. Representative fluorescence images of the parental strain JSJ13 (rips‐1p::GFP) and the mutants isolated from the forward genetic screen. The isolated mutants were designated JSJ14 to JSJ16. Scale bar = 200 μm.
  2. Quantification of GFP levels of rips‐1p::GFP in JSJ13‐JSJ16 animals. ***P < 0.001 via the t‐test (n = 15 worms each).
  3. Table summarizing the alleles identified by whole‐genome sequencing in JSJ14‐JSJ16 strains.
  4. Representative fluorescence images of rips‐1p::GFP animals following RNAi against egl‐9, rhy‐1, and vhl‐1. Animals grown on empty RNAi vector (EV) were used as the control. Scale bar = 200 μm.
  5. Quantification of GFP levels of rips‐1p::GFP animals following RNAi against egl‐9, rhy‐1, and vhl‐1, along with EV control. ***P < 0.001 via the t‐test (n = 10 worms each).
  6. Representative fluorescence images of rips‐1p::GFP and hif‐1(ia4);rips‐1p::GFP animals grown on 0 mM DTT until the young adult stage, followed by incubation on 0 or 10 mM DTT for 10 h. The red fluorescence in the pharynx region is from the myo‐2p::mCherry coinjection marker. Scale bar = 200 μm.
  7. Quantification of GFP levels of rips‐1p::GFP and hif‐1(ia4);rips‐1p::GFP animals grown on 0 mM DTT until the young adult stage, followed by incubation on 0 or 10 mM DTT for 10 h. ***P < 0.001 and **P < 0.01 via the t‐test (n = 14–15 worms each).

Data information: In the boxplots in panels (B, E, and G), the central bands represent the median value, the boxes represent the upper and lower quartile, and the whiskers represent the minimum and maximum values.

Source data are available online for this figure.

The genes egl‐9, rhy‐1, and vhl‐1 are involved in the degradation of the transcription factor hypoxia‐inducible factor 1 (HIF‐1) (Epstein et al, 2001; Shen et al, 2006). Inhibition of these genes results in the accumulation of HIF‐1 and activates the expression of hypoxia response genes (Shen et al, 2006). We asked whether the upregulation of rips‐1 also required HIF‐1 by using hif‐1(ia4) animals. The hif‐1(ia4) allele is a predicted null allele with deletion of exons 2–4 and lacks the induction of hypoxia response genes (Shen et al, 2005). The basal expression of rips‐1 was independent of HIF‐1 as hif‐1(ia4);rips‐1p::GFP animals had GFP in the posterior region of the intestine (Fig 1F and G). Indeed, hif‐1(ia4);rips‐1p::GFP animals showed slightly increased levels of GFP in the posterior region of the intestine compared to rips‐1p::GFP animals suggesting a compensatory induction of basal levels of rips‐1 in the absence of HIF‐1. Next, we exposed the hif‐1(ia4);rips‐1p::GFP animals to DTT and monitored changes in GFP levels. While DTT exposure resulted in high GFP levels in rips‐1p::GFP animals, GFP levels did not increase in hif‐1(ia4);rips‐1p::GFP animals (Fig 1F and G). Taken together, these data indicated that the hypoxia response pathway regulates the expression of rips‐1.

CYSL‐1 is required for DTT‐mediated rips‐1 upregulation

To better understand the interplay between rips‐1 expression and the hypoxia response pathway, we carried out another forward genetic screen to isolate mutants that failed to upregulate rips‐1 upon exposure to DTT—a phenotype similar to hif‐1 loss‐of‐function mutants (Fig EV2A). We isolated two mutants that failed to upregulate rips‐1 upon exposure to DTT (Fig 2A–D). To identify the causative mutations, the variants in the mutant animals were identified by whole‐genome sequencing. Evaluation of the protein‐coding genes having mutations in the linked regions revealed that both mutants had a missense mutation in the gene cysl‐1 (Fig 2E and Appendix Table S1). Previous studies have identified cysl‐1 as a positive regulator of the hypoxia response pathway that inhibits egl‐9 activities (Ma et al, 2012; Wang et al, 2022).

Figure EV2. CYSL‐1 is required for DTT‐mediated rips‐1 upregulation.

Figure EV2

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  1. Scheme for a forward genetic screen for mutants that lack upregulation of rips‐1p::GFP on 10 mM DTT.
  2. Representative fluorescence images of JSJ18 (cysl‐1(jsn18);rips‐1p::GFP) animals following RNAi against egl‐9, rhy‐1, and vhl‐1, along with EV control. Scale bar = 200 μm.
  3. Quantification of GFP levels of JSJ18 (cysl‐1(jsn18);rips‐1p::GFP) animals following RNAi against egl‐9, rhy‐1, and vhl‐1, along with EV control. ***P < 0.001 and *P < 0.05 via the t‐test (n = 15 worms each). In the boxplots, the central bands represent the median value, the boxes represent the upper and lower quartile, and the whiskers represent the minimum and maximum values.

Source data are available online for this figure.

Figure 2. CYSL‐1 is required for DTT‐mediated rips‐1 upregulation.

Figure 2

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  • A
    Representative fluorescence images of the parental strain JSJ13 (rips‐1p::GFP) and the isolated mutants, JSJ17 and JSJ18, grown on 0 mM DTT until the young adult stage, followed by incubation on 0 or 10 mM DTT for 10 h. The red fluorescence in the pharynx region is from the myo‐2p::mCherry coinjection marker. Scale bar = 200 μm.
  • B–D
    Quantification of GFP levels of rips‐1p::GFP in JSJ13 (B), JSJ17 (C), and JSJ18 (D) animals grown on 0 mM DTT until the young adult stage, followed by incubation on 0 or 10 mM DTT for 10 h. 0 mM DTT data in (B) is the same as 0 mM WT data in Fig 1G. ***P < 0.001 via the t‐test (n = 14–15 worms each).
  • E
    Mapping of the cysl‐1 alleles identified in the forward genetic screen.
  • F
    Representative fluorescence images of JSJ17 (cysl‐1(jsn17);rips‐1p::GFP) animals following RNAi against egl‐9, rhy‐1, and vhl‐1, along with EV control. Scale bar = 200 μm.
  • G
    Quantification of GFP levels of JSJ17 (cysl‐1(jsn17);rips‐1p::GFP) animals following RNAi against egl‐9, rhy‐1, and vhl‐1, along with EV control. ***P < 0.001 and **P < 0.01 via the t‐test (n = 15 worms each).

Data information: In the boxplots in panels (B–D and G), the central bands represent the median value, the boxes represent the upper and lower quartile, and the whiskers represent the minimum and maximum values.

Source data are available online for this figure.

Because cysl‐1 may act upstream or downstream of hif‐1 depending on the context (Budde & Roth, 2011; Ma et al, 2012), we carried out an epistasis analysis of cysl‐1 with the other components of the hypoxia response pathway in the context of rips‐1 expression. To this end, we knocked down egl‐9, rhy‐1, and vhl‐1 genes in cysl‐1(jsn17);rips‐1p::GFP and cysl‐1(jsn18);rips‐1p::GFP animals and monitored the changes in GFP expression. Knockdown of egl‐9 and vhl‐1 resulted in the upregulation of rips‐1 in both the alleles of cysl‐1 (Figs 2F and G, and EV2B and C), indicating that egl‐9 and vhl‐1 do not require cysl‐1 to regulate the expression of rips‐1 and work downstream of cysl‐1 to regulate hif‐1. On the other hand, the knockdown of rhy‐1 did not lead to the upregulation of rips‐1 in cysl‐1 mutants (Figs 2F and G, and EV2B and C), indicating rhy‐1 functions upstream of cysl‐1. Because egl‐9 and vhl‐1 are known to work upstream of hif‐1, these results indicated that cysl‐1 acts upstream of egl‐9, vhl‐1, and hif‐1 in rips‐1 expression. Our results are in line with previous observations that showed that cysl‐1 acts downstream of rhy‐1 and upstream of egl‐9, vhl‐1, and hif‐1 (Ma et al, 2012).

Thiol reductive stress activates the hypoxia response pathway

The involvement of the hypoxia response pathway in DTT‐mediated upregulation of rips‐1 suggested that thiol reductive stress may activate the hypoxia response pathway. To test this possibility, we analyzed the mRNA levels of several genes that are induced by hypoxia, including cysl‐2, rhy‐1, nhr‐57, fmo‐2, sqrd‐1, gst‐19, and oac‐54 (Shen et al, 2005). All of these genes had significantly increased levels in DTT‐exposed animals compared with the control animals (Fig 3A). The DTT‐mediated induction of most of the hypoxia‐response genes was dependent on hif‐1 (Fig 3B). The only exceptions were nhr‐57 and fmo‐2. The upregulation of nhr‐57 was partially dependent on hif‐1, while the upregulation of fmo‐2 was independent of hif‐1 (Fig 3A and B). The expression of fmo‐2 under hypoxic conditions is known to be regulated by the transcription factor NHR‐49 (Doering et al, 2022). Further, previous studies have also reported that the expression of nhr‐57 is either partially or fully independent of HIF‐1 (Shen et al, 2005; Doering et al, 2022). Taken together, these studies showed that DTT exposure results in the activation of a primarily HIF‐1‐mediated hypoxia response.

Figure 3. Thiol reductive stress activates the hypoxia response pathway.

Figure 3

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  • A, B
    Gene expression analysis of N2 (A) and hif‐1(ia4) (B) animals grown on 0 mM DTT until the young adult stage, followed by incubation on 0 or 10 mM DTT for 10 h. ***P < 0.001, **P < 0.01, and *P < 0.05 via the t‐test. n.s., nonsignificant (n = 3 biological replicates). Data represent the mean and standard deviation from three independent experiments.
  • C–E
    Survival plots of N2 (C), hif‐1(ia4) (D), and cysl‐1(jsn17) (E) animals on P. aeruginosa PAO1 under fast‐killing assay conditions. Before transferring to P. aeruginosa PAO1 lawns at the L4 stage, the animals were grown on Comamonas aquatica DA1877 diet containing 0 or 5 mM DTT. The plots of cysl‐1(jsn17) animals are identical, as all the animals (from 0 and 5 mM DTT) are dead by the first time point (4 h). Data represent the mean and standard deviation from three independent experiments.
  • F
    Percent dead animals on P. aeruginosa PAO1 under fast‐killing assay conditions after 4 h of exposure. Before transferring to P. aeruginosa PAO1 lawns at the L4 stage, the animals were grown on Comamonas aquatica DA1877 diet containing 0 or 5 mM DTT. Data represent the mean and standard deviation from three independent experiments. **P < 0.01 via the t‐test. n.s., nonsignificant.

Source data are available online for this figure.

Next, we asked whether DTT activated a functional hypoxia response pathway. Enhanced activity of the hypoxia response pathway has been shown to impart resistance to hydrogen cyanide‐mediated fast paralytic killing of C. elegans by P. aeruginosa PAO1 (Gallagher & Manoil, 2001). We observed that pre‐treatment with DTT enhanced the resistance of wild‐type N2 animals to fast paralytic killing by P. aeruginosa PAO1 (Fig 3C). The hypoxia response pathway mutants exhibited enhanced susceptibility to P. aeruginosa PAO1 (Fig 3D and E), confirming the requirement of the hypoxia response pathway to protect against paralytic killing by P. aeruginosa PAO1. Importantly, pre‐treatment with DTT did not enhance the survival of the hypoxia response pathway mutants (Fig 3D–F). These results indicated that DTT activates a functional hypoxia response pathway that protects C. elegans against hydrogen cyanide‐mediated fast paralytic killing by P. aeruginosa PAO1.

Hypoxia response pathway has opposing roles in regulating thiol‐mediated toxicity

Next, we asked whether activation of the hypoxia response pathway had any roles in regulating thiol‐mediated toxicity. Thiol antioxidants‐mediated upregulation of rips‐1 modulates the methionine‐homocysteine cycle and results in developmental toxicity (Gokul & Singh, 2022; Winter et al, 2022). Therefore, it appears that the activation of the hypoxia response pathway by thiol antioxidants, which results in the upregulation of rips‐1, might be harmful to the host. To understand the role of the hypoxia response pathway in toxicity by thiol reductive stress, we studied the response of hif‐1(ia4), cysl‐1(jsn17), and cysl‐1(jsn18) animals to varying concentrations of DTT. Compared to wild‐type N2 animals, hif‐1 and cysl‐1 loss‐of‐function mutants had improved development on 5 and 7.5 mM DTT (Figs 4A–D and EV3A–C). These results indicated that the hypoxia response pathway exacerbated thiol toxicity, likely by increasing the expression of rips‐1.

Figure 4. Hypoxia response pathway has opposing roles in regulating thiol‐mediated toxicity.

Figure 4

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  • A
    Representative images of N2, hif‐1(ia4), and cysl‐1(jsn17) animals on various concentrations of DTT on E. coli OP50 diet after 72 h of hatching at 20°C. Scale bar = 1 mm.
  • B–D
    Quantification of different developmental stages of N2 (B), hif‐1(ia4) (C), and cysl‐1(jsn17) (D) animals on various concentrations of DTT on E. coli OP50 diet after 72 h of hatching at 20°C (n = 3 biological replicates; animals per condition per replicate > 100).
  • E
    Representative images of N2, hif‐1(ia4), cysl‐1(jsn17), and cysl‐1(jsn18) animals after 72 h of hatching at 20°C on E. coli OP50 diet containing 0, 10, and 10 mM DTT supplemented with 50 nM vitamin B12. Scale bar = 1 mm.
  • F
    Quantification of different developmental stages of N2, hif‐1(ia4), cysl‐1(jsn17), and cysl‐1(jsn18) animals after 72 h of hatching at 20°C on E. coli OP50 diet containing 0, 10, and 10 mM DTT supplemented with 50 nM vitamin B12. The 10 mM DTT is the same as in (B–D) (n = 3 biological replicates; animals per condition per replicate > 100).

Data information: For (B–D and F), data represent the mean and standard deviation from three independent experiments.

Source data are available online for this figure.

Figure EV3. cysl‐1(jsn18) animals develop better than N2 animals at 5 mM DTT.

Figure EV3

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  • A
    Representative images of N2 and cysl‐1(jsn18) animals on various concentrations of DTT on E. coli OP50 diet after 72 h of hatching at 20°C. Scale bar = 1 mm.
  • B, C
    Quantification of different developmental stages of N2 (B) and cysl‐1(jsn18) (C) animals on various concentrations of DTT on E. coli OP50 diet after 72 h of hatching at 20°C. The N2 data in (B) are the same as in Fig 4B (n = 3 biological replicates; animals per condition per replicate > 100). Data represent the mean and standard deviation from three independent experiments.

Source data are available online for this figure.

Interestingly, at 10 mM DTT, the hypoxia response pathway mutants failed to develop, similar to the wild‐type animals (Figs 4A–D and EV3A–C). Because vitamin B12 restores the DTT‐perturbed methionine‐homocysteine cycle, vitamin B12 alleviates DTT toxicity in wild‐type animals (Gokul & Singh, 2022; Winter et al, 2022). Despite lacking induction of rips‐1, the hypoxia response pathway mutants failed to develop on 10 mM DTT (Fig 4A–D). Therefore, we asked whether supplementation of vitamin B12 would alleviate DTT toxicity in the hypoxia response pathway mutants. While the supplementation of 50 nM vitamin B12 alleviated DTT toxicity in wild‐type N2 animals, it failed to do so in the hypoxia response pathway mutants (Fig 4E and F). Similarly, the wild‐type animals developed on 10 mM DTT in the presence of the bacterial diet Comamonas aquatica DA1877, which has higher vitamin B12 compared to the E. coli OP50 diet (Watson et al, 2014; Gokul & Singh, 2022), while the hypoxia response pathway mutants failed to develop under these conditions (Fig EV4A and B). Vitamin B12 did not affect the expression of rips‐1 either in the absence or presence of DTT (Fig EV4C and D), indicating that vitamin B12 does not regulate thiol toxicity by modulating the hypoxia response pathway. Together, these results revealed a complicated picture of the role of the hypoxia response pathway in regulating thiol‐mediated toxicity. By enhancing the expression of rips‐1 under thiol reductive stress, the hypoxia response pathway predisposes C. elegans to thiol‐mediated toxicity. However, at higher concentrations of DTT, it emerges that the hypoxia response pathway protects against thiol reductive stress independent of rips‐1 expression as hif‐1 and cysl‐1 mutants fail to develop on 10 mM DTT even upon supplementation of vitamin B12.

Figure EV4. Hypoxia response pathway protects against thiol reductive stress.

Figure EV4

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  1. Representative images of N2, hif‐1(ia4), cysl‐1(jsn17), and cysl‐1(jsn18) animals after 72 h of hatching at 20°C on C. aquatica DA1877 diet containing 0 or 10 mM DTT. Scale bar = 1 mm.
  2. Quantification of different developmental stages of N2, hif‐1(ia4), cysl‐1(jsn17), and cysl‐1(jsn18) animals after 72 h of hatching at 20°C on C. aquatica DA1877 diet containing 0 or 10 mM DTT (n = 3 biological replicates; animals per condition per replicate > 100). Data represent the mean and standard deviation from three independent experiments.
  3. Representative fluorescence images of rips‐1p::GFP animals grown on E. coli OP50 diet without any supplements until the young adult stage, followed by incubation on E. coli OP50 diet containing no supplement (0 mM DTT), 50 nM vitamin B12, 10 mM DTT, and 10 mM DTT + 50 nM vitamin B12 for 10 h. Scale bar = 200 μm.
  4. Quantification of GFP levels of rips‐1p::GFP animals grown on E. coli OP50 diet without any supplements until the young adult stage, followed by incubation on E. coli OP50 diet containing no supplement (0 mM DTT), 50 nM vitamin B12, 10 mM DTT, and 10 mM DTT + 50 nM vitamin B12 for 10 h. n.s., nonsignificant via the t‐test (n = 14 animals each). In the boxplots, the central bands represent the median value, the boxes represent the upper and lower quartile, and the whiskers represent the minimum and maximum values.

Source data are available online for this figure.

Activation of the hypoxia response pathway by thiol reductive stress is conserved in human cell lines

Next, we asked whether the activation of the hypoxia response pathway by thiol reductive stress is a conserved response. To this end, we used three human cell lines from different tissue origins, including HeLa (cervical cancer), LN229 (glioblastoma), and Cal27 (oral squamous cell carcinoma). First, we tested whether the survival of these cell lines was affected by exposure to 10 mM DTT. Exposure of these cell lines to 10 mM DTT for 24 h did not affect their viability (Appendix Fig S1). We then evaluated the mRNA levels of five hypoxia‐response genes, including hypoxia‐inducible factor‐1α (HIF‐1α), HIF‐1β, 3‐phosphoinositide‐dependent protein kinase‐1 (PDK1), glucose transporter 1 (GLUT1), and BCL2/adenovirus E1B 19 kDa protein‐interacting protein 3 (BNIP3) upon exposure of the cells to 10 mM DTT for 24 h. All of these genes are known to be upregulated under hypoxic conditions (Guo et al, 2001; BelAiba et al, 2007; Choudhry & Harris, 2018). DTT treatment also resulted in the upregulation of these genes in HeLa, LN229, and Cal27 cell lines (Fig 5A–C). These results showed that thiol reductive stress activates the hypoxia response pathway in human cell lines.

Figure 5. Thiol reductive stress activates the hypoxia response pathway in human cell lines.

Figure 5

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  • A–C
    Gene expression analysis of HeLa (A), LN229 (B), and Cal27 (C) cell lines with and without exposure to 10 mM DTT for 24 h. ***P < 0.001, **P < 0.01, and *P < 0.05 via the t‐test. n.s., nonsignificant (n = 3 biological replicates with 2–3 technical repeats each). Data represent the mean and standard deviation from three independent experiments.
  • D
    Representative fluorescence images of HeLa, LN229, and Cal27 cell lines upon treatment with Image‐iT™ green hypoxia reagent. The cells were pretreated with 0 or 10 mM DTT for 24 h before treatment with Image‐iT™ green hypoxia reagent. Scale bar = 100 μm.
  • E–G
    Quantification of fluorescence levels of Image‐iT™ green hypoxia reagent‐treated HeLa (E), LN229 (F), and Cal27 (G) cell lines. The cells were pretreated with 0 or 10 mM DTT for 24 h before treatment with Image‐iT™ green hypoxia reagent. ***P < 0.001 via the t‐test (n = 9 wells each). In the boxplots, the central bands represent the median value, the boxes represent the upper and lower quartile, and the whiskers represent the minimum and maximum values.

Source data are available online for this figure.

Next, we tested whether DTT resulted in a hypoxia‐like cellular milieu. To this end, we stained human cell lines with Image‐iT green hypoxia reagent after treatment with 10 mM DTT. Image‐iT green hypoxia reagent is a fluorogenic compound that is live cell permeable and senses hypoxic environments. While it is unclear whether the Image‐iT green hypoxia reagent senses oxygen levels directly or reports other hypoxia‐related physiological changes, it fluoresces only under hypoxic conditions. DTT treatment resulted in a drastic increase in fluorescence in HeLa, LN229, and Cal27 cell lines (Fig 5D–G), indicating a hypoxia‐like cellular milieu upon DTT exposure. To study whether the increase in the fluorescence was because of a hypoxic milieu or a change in the redox environment because of DTT, we tested the fluorescence changes in cells upon exposure to 10 mM N‐acetylcysteine (NAC). Similar to DTT, NAC results in a reduced cellular environment, but it does not lead to the upregulation of rips‐1 (Gokul & Singh, 2022). The Image‐iT hypoxia reagent fluorescence remained unaltered in cells exposed to 10 mM NAC for 24 h (Appendix Fig S2), indicating that the increased fluorescence upon DTT exposure was unlikely because of a change in the redox environment.

DTT leads to hydrogen sulfide production

Finally, we asked how thiol reductive stress led to the activation of the hypoxia response pathway. Thiol antioxidants result in the breakage of disulfide bonds in the ER (Maity et al, 2016). This is followed by the activation of futile oxidative cycles in the ER, enhancing disulfide bond turnover (Maity et al, 2016). Disulfide bond formation consumes oxygen and produces hydrogen peroxide (Zito, 2015). Therefore, thiol reductive stress generates hydrogen peroxide and causes oxidative stress (Maity et al, 2016). We hypothesized that thiol reductive stress might activate the hypoxia response pathway by creating oxidative stress or directly enhancing oxygen consumption. To test these possibilities, we probed the role of oxidative stress in the upregulation of rips‐1. While oxidative stress by tert‐butyl hydroperoxide, ferric chloride, and paraquat resulted in the induction of oxidative stress reporter gst‐4p::GFP, it did not affect the expression of rips‐1p::GFP (Fig EV5A–D). These results suggested that oxidative stress is not responsible for the DTT‐triggered hypoxia response pathway activation.

Figure EV5. The futile oxidative cycle in the ER under DTT stress does not result in the activation of the hypoxia response pathway.

Figure EV5

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  • A
    Representative fluorescence images of rips‐1p::GFP animals grown on E. coli OP50 diet without any supplements until the young adult stage, followed by incubation on E. coli OP50 diet containing no supplement (control), 10 mM DTT, 5 mM tert‐butyl hydroperoxide (tBHP), 7.5 mM ferric chloride (FeCl3), and 1 mM paraquat (PQ) for 12 h. Scale bar = 200 μm.
  • B
    Quantification of GFP levels of rips‐1p::GFP animals grown on E. coli OP50 diet without any supplements until the young adult stage, followed by incubation on E. coli OP50 diet containing no supplement (control), 10 mM DTT, 5 mM tBHP, 7.5 mM FeCl3, and 1 mM paraquat for 12 h. ***P < 0.001 and **P < 0.01 via the t‐test. n.s., nonsignificant (n = 14 worms each). In the boxplots, the central bands represent the median value, the boxes represent the upper and lower quartile, and the whiskers represent the minimum and maximum values.
  • C
    Representative fluorescence images of gst‐4p::GFP animals grown on E. coli OP50 diet without any supplements until the young adult stage, followed by incubation on E. coli OP50 diet containing no supplement (control), 10 mM DTT, 5 mM tBHP, 7.5 mM FeCl3, and 1 mM paraquat (PQ) for 12 h. Scale bar = 200 μm.
  • D
    Quantification of GFP levels of gst‐4p::GFP animals grown on E. coli OP50 diet without any supplements until the young adult stage, followed by incubation on E. coli OP50 diet containing no supplement (control), 10 mM DTT, 5 mM tBHP, 7.5 mM FeCl3, and 1 mM paraquat for 12 h. ***P < 0.001 via the t‐test (n = 15 worms each). In the boxplots, the central bands represent the median value, the boxes represent the upper and lower quartile, and the whiskers represent the minimum and maximum values.
  • E–H
    Oxygen consumption rates (OCR) in medium control without cells (E), HeLa (F), LN229 (G), and Cal27 (H) cells. After three OCR measurements, water or DTT was injected in the wells to obtain 0 or 10 mM DTT and is indicated as injection in the plots. The cells were seeded at 2 × 104 cells per well. The data represent the mean and standard error from three wells per condition except for LN229 at 10 mM DTT. The data for LN229 at 10 mM DTT represent the mean (without error bars) from two wells. The error bars for control at 0 mM DTT and Cal27 at 10 mM DTT are not visible as the error bars are shorter than the data symbols.

Source data are available online for this figure.

We then tested whether thiol reductive stress resulted in enhanced oxygen consumption. To this end, we measured oxygen consumption rates (OCR) in human cell lines upon exposure to 10 mM DTT. The addition of DTT resulted in a rapid increase in OCR not only in the cell lines but also in a blank control of medium without cells (Fig EV5E–H). This indicated that the addition of DTT might increase OCR because of the oxidation of DTT, which results in oxygen consumption. However, the cells did not exhibit a further increase in OCR, indicating that DTT did not increase oxygen consumption in human cells. These results suggested that DTT does not activate the hypoxia response pathway by enhancing oxygen consumption.

Hydrogen sulfide (H2S) is known to activate the hypoxia response pathway via CYSL‐1 (Ma et al, 2012). Because DTT also required cysl‐1 for the activation of the hypoxia response pathway (Fig 2), we asked whether DTT resulted in H2S production. To this end, we incubated worm lysate with DTT and measured H2S production. While the worm lysate in the absence of DTT did not lead to H2S production, H2S formation was observed in the presence of DTT (Fig 6A). The worm lysate was required for H2S production by DTT, as the addition of DTT to the buffer control did not result in the production of H2S (Fig 6A). Therefore, it appeared that DTT might result in the production of H2S in C. elegans.

Figure 6. The mechanism of activation of the hypoxia response pathway by DTT.

Figure 6

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  1. Images of lead acetate paper showing H2S production in worm lysate incubated with DTT. Buffer control with DTT and worm lysate without DTT did not show H2S production. Data from three independent experiments is shown. The H2S levels reference chart is shown on the right side.
  2. Model for the mechanism of activation of the hypoxia response pathway by DTT.

Source data are available online for this figure.

Discussion

Studies in the early 1990s indicated that DTT affects the oxidative environment in the ER without impacting other cellular processes (Braakman et al, 1992; Lodish & Kong, 1993; Tatu et al, 1993; Jamsa et al, 1994). Since then, DTT has been used as an ER‐specific stressor in a large number of studies in various model organisms (Frand & Kaiser, 1998; Pollard et al, 1998; Merksamer et al, 2008; Qin et al, 2010; Oslowski & Urano, 2011). Meanwhile, some studies suggested that DTT might broadly impact cellular physiology and affect cellular processes beyond ER (MacKenzie et al, 2005; Sims et al, 2005; Guillemette et al, 2007; Gokul & Singh, 2022; Winter et al, 2022). However, the mechanisms of such physiological impacts of DTT remained unexplored. Our current study shows that DTT activates the hypoxia response pathway and challenges the current model that DTT disrupts only the ER milieu in the cell. Our study also highlights that some of the earlier work that used DTT as an ER‐specific stressor might need reevaluation in light of the role of DTT in activating the hypoxia response pathway.

Thiol antioxidants result in the activation of futile oxidative cycles in the ER, enhancing disulfide bond turnover (Maity et al, 2016). Disulfide bond formation consumes oxygen and produces hydrogen peroxide (Zito, 2015). However, neither oxygen consumption nor hydrogen peroxide production seems to be required for DTT‐mediated hypoxia response pathway activation. Earlier studies have shown that oxidative stress could activate the HIF‐1‐dependent hypoxia response (Lee et al, 2010). However, we did not observe the upregulation of rips‐1 under oxidative stress conditions. Because different stimulants could result in non‐overlapping responses via HIF‐1 (Miller et al, 2011), it is possible that the oxidative stress‐triggered hypoxia response does not involve the upregulation of rips‐1. However, the DTT‐mediated hypoxia response pathway involves the upregulation of rips‐1, suggesting that DTT‐induced activation of the hypoxia response pathway is unlikely because of oxidative stress.

We discovered that DTT results in H2S production when incubated with worm lysate. H2S is known to activate the hypoxia response pathway via CYSL‐1 (Ma et al, 2012). Because DTT and H2S activate the hypoxia response through the same pathway, DTT may activate the hypoxia pathway via H2S production (Fig 6B). However, from our current data, it is not possible to determine whether the amount of H2S produced by DTT is sufficient to induce the hypoxia response pathway. It is also possible that DTT directly acts on CYSL‐1, similar to H2S (Ma et al, 2012), to activate the hypoxia response pathway. Our studies do not rule out the possibility of other mechanisms of activation of the hypoxia response pathway by DTT. It is important to note that reduced thiols in the cell could also enhance HIF‐1 degradation by enhancing the activity of EglN1 (Briggs et al, 2016), the C. elegans egl‐9 homolog. Therefore, thiol antioxidants could have opposing roles in stabilizing HIF‐1, and the outcome might depend on the context or the amount of reduced thiols. Future studies exploring the precise mechanisms of activation of the hypoxia response pathway by DTT in different model systems, including human cell lines, will be very useful in understanding the link between thiol reductive stress and the hypoxia response pathway.

Despite links with various pathological conditions, the physiological effects of reductive stress remain poorly characterized (Handy & Loscalzo, 2017; Pérez‐Torres et al, 2017; Ma et al, 2020; Rajasekaran, 2020). In this study, we show that thiol reductive stress activates the hypoxia response pathway. The activation of the hypoxia response pathway appears to have opposing roles in thiol toxicity in C. elegans. By enhancing the expression of rips‐1 under thiol reductive stress, the hypoxia response pathway predisposes C. elegans to thiol‐mediated toxicity (Fig 6B). Interestingly, SAM‐dependent methyltransferases, that are induced by DTT, confer protection against thiol stress in some organisms (MacKenzie et al, 2005; Dolan et al, 2014; Owens et al, 2015; Manzanares‐miralles et al, 2016). Therefore, the activation of rips‐1 might be a defense response against thiol stress and its toxic effects because of SAM depletion (Gokul & Singh, 2022) could be a side effect. The hypoxia response pathway is known to activate antioxidant responses (Vora et al, 2022), which may enhance thiol toxicity. However, because different stimulants could result in non‐overlapping responses via HIF‐1 (Miller et al, 2011), comparing the HIF‐1 targets activated by hypoxia versus thiol stress will be important. The hypoxia response pathway appears to protect against thiol reductive stress independent of rips‐1 expression (Fig 6B). Hypoxia has been shown to protect against mitochondrial reductive stress by enhancing the production of L‐2‐hydroxyglutarate (Oldham et al, 2015). The hypoxia response pathway also enhances the catabolism of hydrogen sulfide (H2S) by enhancing the expression of sulfide:quinone oxidoreductase (Budde & Roth, 2011; Malagrinò et al, 2019). In future studies, understanding the players downstream of HIF‐1 that protect against thiol stress would help establish the interactions between thiol reductive stress and the hypoxia response pathway.

Materials and Methods

Bacterial strains

The following bacterial strains were used in the current study: Escherichia coli OP50, E. coli HT115(DE3), Pseudomonas aeruginosa PAO1, and Comamonas aquatica DA1877. The cultures of E. coli OP50, E. coli HT115(DE3), and C. aquatica DA1877 were grown in Luria‐Bertani (LB) broth at 37°C. The P. aeruginosa PAO1 cultures were grown in brain heart infusion (BHI) broth at 37°C.

C. elegans strains and growth conditions

Caenorhabditis elegans hermaphrodites were maintained at 20°C on nematode growth medium (NGM) plates seeded with E. coli OP50 as the food source unless otherwise indicated. Bristol N2 was used as the wild‐type control unless otherwise indicated. The following strains were used in the study: JSJ13 jsnIs1[rips‐1p::GFP + myo‐2p::mCherry], ZG31 hif‐1(ia4), RE666 ire‐1(v33), xbp‐1(tm2482), RB772 atf‐6(ok551), RB545 pek‐1(ok275), VC3201 atfs‐1(gk3094), and CL2166 dvIs19[(pAF15)gst‐4p::GFP::NLS]. The following strains were generated in this study: JSJ14 jsnIs1[rips‐1p::GFP + myo‐2p::mCherry];rhy‐1(jsn13);rhy‐1(jsn14), JSJ15 jsnIs1[rips‐1p::GFP + myo‐2p::mCherry];egl‐9(jsn15), JSJ16 jsnIs1[rips‐1p::GFP + myo‐2p::mCherry];vhl‐1(jsn16), JSJ17 jsnIs1[rips‐1p::GFP + myo‐2p::mCherry];cysl‐1(jsn17), and JSJ18 jsnIs1[rips‐1p::GFP + myo‐2p::mCherry];cysl‐1(jsn18). Some of the strains were obtained from the Caenorhabditis Genetics Center (University of Minnesota, Minneapolis, MN). The hif‐1(ia4);rips‐1p::GFP, ire‐1(v33);rips‐1p::GFP, xbp‐1(tm2482);rips‐1p::GFP, atf‐6(ok551);rips‐1p::GFP, pek‐1(ok275);rips‐1p::GFP, and atfs‐1(gk3094);rips‐1p::GFP strains were obtained by standard genetic crosses.

Preparation of NGM plates with different supplements

The following supplements with their product numbers were obtained from HiMedia BioSciences: DTT (#RM525), vitamin B12 (cyanocobalamin) (#PCT0204), ferric chloride (#TC583), pyridoxine hydrochloride (#TC039), NAC (#RM3142) and tert‐butyl hydroperoxide (#RM2022). Paraquat (#856177) and MTT (#M5655) were obtained from Sigma‐Aldrich. The supplements for C. elegans exposure were added to NGM just before pouring them into plates to obtain the desired final concentration.

C. elegans development assays

Synchronized C. elegans eggs were obtained by transferring 20–25 gravid adult hermaphrodites on NGM plates for egg‐laying for 2 h. After 2 h, the gravid adults were removed. The synchronized eggs were incubated at 20°C for 72 h. After that, the animals in different development stages (L1/L2, L3, and L4/adult) were quantified. Representative images of the NGM plates at the time of quantification of development were also captured. At least three independent experiments (biological replicates) were performed for each condition.

RNA interference (RNAi)

RNA interference was used to generate loss‐of‐function RNAi phenotypes by feeding worms with E. coli HT115(DE3) expressing double‐stranded RNA homologous to a target C. elegans gene. E. coli with the appropriate vectors were grown in LB broth containing ampicillin (100 μg/ml) at 37°C overnight and plated onto RNAi NGM plates containing 100 μg/ml ampicillin and 3 mM isopropyl β‐D‐thiogalactoside (IPTG). The RNAi‐expressing bacteria were allowed to grow overnight at 37°C on RNAi plates. The worms were synchronized on RNAi plates, and the eggs were allowed to develop at 20°C for 72 h. The RNAi clones were from the Ahringer RNAi library and were verified by sequencing.

Forward genetic screens for mutants with upregulated rips‐1p::GFP expression levels

Ethyl methanesulfonate (EMS) mutagenesis screens (Singh, 2021) were performed using the rips‐1p::GFP strain. Approximately 2,500 synchronized late L4 larvae were treated with 50 mM EMS for 4 h and then washed three times with M9 medium. The washed animals (P0 generation) were then transferred to 9‐cm NGM plates containing E. coli OP50 and allowed to lay eggs (F1 progeny) overnight. The P0s were then washed away with M9 medium, while the F1 eggs remained attached to the bacterial lawn. The F1 eggs were allowed to grow to adulthood. The adult F1 animals were bleached to obtain F2 eggs. The F2 eggs were transferred to E. coli OP50 plates containing 50 nM vitamin B12 and incubated at 20°C for 72 h. After that, the plates were screened for animals that had high GFP levels. Approximately 50,000 haploid genomes were screened, and three mutants were isolated. All of the mutants were backcrossed six times with the parental rips‐1p::GFP strain before analysis.

Forward genetic screens for mutants that lacked DTT‐mediated rips‐1p::GFP upregulation

All the mutagenesis steps were identical to the screen for mutants with upregulated rips‐1p::GFP expression levels until the F2 eggs stage. The F2 eggs were transferred to E. coli OP50 plates and incubated at 20°C for 72 h. After that, the adult F2 animals were washed away with M9 and transferred to E. coli OP50 plates containing 10 mM DTT. After 10 h of incubation, the plates were screened for animals that lacked GFP induction. Approximately 50,000 haploid genomes were screened, and two mutants were isolated. Both the mutants were backcrossed six times with the parental rips‐1p::GFP strain before analysis.

Whole‐genome sequencing (WGS) and data analysis

The genomic DNA was isolated as described earlier (Singh & Aballay, 2017; Gokul & Singh, 2022). Briefly, the mutant animals were grown at 20°C on NGM plates seeded with E. coli OP50 until starvation. Four 9‐cm E. coli OP50 plates were used for each strain to obtain a sufficient number of animals. The animals were rinsed off the plates with M9, washed three times, incubated in M9 with rotation for 2 h to eliminate food from the intestine, and washed three times again with distilled water, followed by storage at −80°C until genomic DNA extraction. Genomic DNA extraction was performed using the Gentra Puregene Kit (Qiagen, Netherlands). DNA libraries were prepared according to a standard Illumina (San Diego, CA) protocol. The DNA was subjected to WGS on an Illumina NovaSeq 6000 sequencing platform using 150 paired‐end nucleotide reads. Library preparation and WGS were performed at the National Genomics Core, National Institute of Biomedical Genomics, Kalyani, India.

The whole‐genome sequence data were analyzed using the web platform Galaxy as described earlier (Gokul & Singh, 2022). Briefly, the forward and reverse FASTQ reads, C. elegans reference genome Fasta file (ce11M.fa), and the gene annotation file (SnpEff4.3 WBcel235.86) were input into the Galaxy workflow. The low‐quality ends of the FASTQ reads were trimmed using the Sickle tool. The trimmed FASTQ reads were mapped to the reference genome Fasta files with the BWA‐MEM tool. Using the MarkDuplicates tool, any duplicate reads (mapped to multiple sites) were filtered. Subsequently, the variants were detected using the FreeBayes tool that finds small polymorphisms, including single‐nucleotide polymorphisms (SNPs), insertions and deletions (indels), multi‐nucleotide polymorphisms (MNPs), and complex events (composite insertion and substitution events) smaller than the length of a short‐read sequencing alignment. The common variants among different mutants were subtracted. The SnpEff4.3 WBcel235.86 gene annotation file was used to annotate and predict the effects of genetic variants (such as amino acid changes). Finally, the linkage maps for each mutant were generated using the obtained variations.

C. elegans RNA isolation and quantitative reverse transcription‐PCR (qRT‐PCR)

Animals were synchronized by egg laying. Approximately 40 gravid adult animals were transferred to 9‐cm E. coli OP50 plates without DTT and allowed to lay eggs for 4 h. The gravid adults were then removed, and the eggs were allowed to develop at 20°C for 72 h. Subsequently, the synchronized adult animals were collected with M9, washed twice with M9, and then transferred to 9‐cm E. coli OP50 plates containing 10 mM DTT. The control animals were maintained on E. coli OP50 plates without DTT. After the transfer of the animals, the plates were incubated at 20°C for 10 h. Subsequently, the animals were collected, washed with M9 buffer, and frozen in TRIzol reagent (Life Technologies, Carlsbad, CA). Total RNA was extracted using the RNeasy Plus Universal Kit (Qiagen, Netherlands). A total of 1 μg of total RNA was reverse‐transcribed with random primers using the PrimeScript™ 1st strand cDNA Synthesis Kit (TaKaRa) according to the manufacturer's protocols. qRT‐PCR was conducted using TB Green fluorescence (TaKaRa) on a LightCycler 480 II System (Roche Diagnostics) in 96‐well‐plate format. Twenty microliter reactions were analyzed as outlined by the manufacturer (TaKaRa). The relative fold‐changes of the transcripts were calculated using the comparative CT(2−ΔΔCT ) method and normalized to pan‐actin (act‐1, ‐3, ‐4) as described earlier (Singh & Aballay, 2019a). All samples were run in triplicate and repeated at least three times. The sequence of the primers is provided in Appendix Table S2.

Fast‐killing assays on P. aeruginosa PAO1

For fast‐killing assays, P. aeruginosa PAO1 cultures were grown by inoculating individual bacterial colonies into 4–5 ml of BHI broth and incubated for 10–12 h on a shaker at 37°C. Then, the cultures were diluted 10 times in BHI broth, and 50 μl of the diluted culture was spread on a 3.5‐cm BHI agar plate. After spreading bacteria, plates were incubated at 37°C for 24 h. After that, the plates were allowed to cool to room temperature, and worms synchronized on NGM plates seeded with C. aquatica DA1877 with and without 5 mM DTT at the L4 stage were transferred to them. Plates were kept at room temperature and scored for live and dead worms at indicated times of transfer.

H2S production assay

The H2S production capacity assay was adapted from an earlier study (Statzer et al, 2022) and modified for measuring H2S production by DTT. Briefly, synchronized wild‐type N2 young adult C. elegans hermaphrodites were washed off the plates with M9. The worms were washed three times with M9 to remove residual bacteria. The semisoft pellet of worms was weighed to ensure an equal number of worms in each sample (100 μg of worm pellet per sample) and mixed with 800 μl of 1× passive lysis buffer (Promega, #E1910). This was followed by eight cycles of sonication with 3 s on and 30 s off at an amplitude of 40%. The suspension was centrifuged at 12,000 × g at 4°C for 10 min. The supernatant (800 μl) was transferred to new microcentrifuge tubes. The fuel mix was prepared by adding 20 mM DTT and 2.5 mM pyridoxine hydrochloride in phosphate buffer saline. The fuel mix (200 μl) was added to the worm lysate to make the final DTT and pyridoxine hydrochloride concentrations 4 and 0.5 mM, respectively. The tubes containing worm lysate and DTT were sealed with H2S sensing lead acetate strips (HiMedia BioSciences # WT041) and incubated at 37°C for 12 h. After 12 h of incubation, lead acetate strips were observed to determine H2S production.

Impact of oxidative stress on rips‐1p::GFP expression

Egg laying of rips‐1p::GFP and gst‐4p::GFP animals was carried out by transferring gravid adult hermaphrodites on NGM plates seeded with E. coli OP50 for 4 h. Worms were removed after egg laying, and the plates containing eggs were incubated at 20°C. After 72 h, synchronized adult worms were transferred to E. coli OP50 plates containing 1 mM paraquat, 5 mM tert‐butyl hydroperoxide, 10 mM DTT, and 7.5 mM ferric chloride for 12 h. After that, the animals were prepared for fluorescence imaging.

Fluorescence imaging of C. elegans

Fluorescence imaging was carried out as described previously (Singh & Aballay, 2019b; Gokul & Singh, 2022). Briefly, the fluorescence reporter strains were picked using a non‐fluorescence stereomicroscope to avoid potential bias. The animals were anesthetized using an M9 salt solution containing 50 mM sodium azide and mounted onto 2% agarose pads. The animals were then visualized using a Nikon SMZ‐1000 fluorescence stereomicroscope. The fluorescence intensity was quantified using Image J software.

Cell culture

Human cell lines, HeLa (cervical cancer), LN229 (glioblastoma), and Cal27 (oral squamous cell carcinoma), were obtained from the National Center of Cell Science (NCCS), Pune. The cells were cultured in a humidified incubator at 37°C and 5% CO2 in Dulbecco's modified eagle medium (DMEM), supplemented with 10% fetal bovine serum (FBS). The culture medium was replenished every third day. The study with human cell lines was approved by the institute ethics committee, PGIMER, with approval number IEC‐01/2023‐2648.

MTT cell viability assay

To assess cell viability, HeLa, LN229, and Cal27 cell lines were exposed to 10 mM DTT for 24 h, followed by measurement of their survival using MTT (3‐(4,5‐Dimethylthiazol‐2‐yl)‐2,5‐Diphenyltetrazolium Bromide) assay as reported previously (Kumar et al, 2017; preprint: Kumar et al, 2021). Briefly, the cells were seeded at an initial density of 5 × 103 per well in 96‐well cell culture plates and treated with 10 mM DTT for 24 h while the control cells remained untreated. After that, the cells were washed to remove DTT. Then, the cells were treated with 10% MTT solution for 4 h in the dark. The formazan crystals formed from MTT by live cells were solubilized by DMSO. The absorbance of the solution was measured at 570 nm wavelength on a microplate reader (Infinite 200 Pro, TECAN). Readings from 12 different wells were obtained for each condition.

Cell lines RNA isolation and qRT‐PCR

HeLa, LN229, and Cal27 cell lines were treated with 10 mM DTT for 24 h. The total RNA of DTT treated and control cells was isolated using the TRIzol method. Total RNA was reverse transcribed to cDNA using the High‐Capacity cDNA Reverse Transcription Kit (Applied Biosystems), as per the manufacturer's instructions. qRT‐PCR was conducted using DyNAmo ColorFlash SYBR Green (ThermoFisher Scientific) on a CFX96 Real‐Time System (Bio‐Rad) in 96‐well‐plate format. Twenty microliter reactions were analyzed as outlined by the manufacturer (Bio‐Rad). The relative fold changes of the transcripts were calculated using the comparative CT(2−ΔΔCT ) method and normalized to beta‐actin. All samples were run in triplicate and repeated at least three times. The sequence of the primers is provided in Appendix Table S2.

Hypoxia staining of human cell lines and quantification

Human cell lines were pretreated with 10 mM DTT for 24 h. After that, DTT was removed, and the cells were treated with Image‐iT™ Green Hypoxia Reagent (Invitrogen) at a concentration of 1 μM for 30 min at 37°C. The reagent was then removed, and the cells were incubated in either DMEM (control) or DMEM containing 10 mm DTT for 4 h. After that, cell images were captured using a fluorescence microscope (Leica) at the excitation/emission wavelength of 488/520 nm. Simultaneously, cells were separately cultured in 96‐well plates and stained with the Image‐iT™ Green Hypoxia Reagent as described above. The fluorescence of the cells was then measured in a fluorescence plate reader (Infinite 200 Pro, TECAN) at the excitation/emission wavelengths of FITC (488/520 nm) with the following parameters: Z‐position 20,000 μm, gain 100, excitation bandwidth 9 nm, emission bandwidth 20 nm, and the number of flashes 25. As a control for the redox environment, the cells were treated with 10 mM NAC for 24 h and stained with the Image‐iT™ Green Hypoxia Reagent, like the treatment with DTT. The pH of NAC was adjusted to 7.0 using a sodium hydroxide solution (NaOH). The fluorescence of the cells was measured in a fluorescence plate reader as described for DTT.

Oxygen consumption rate (OCR) measurement

Human cell lines HeLa, LN229, and Cal27 were seeded at a density of 2 × 104 cells per well in a Seahorse XFe24 Cell Culture Microplate (102340‐100; Agilent, Santa Clara, CA). Cells were incubated overnight for adherence in a culture media composed of DMEM and 10% FBS at 37°C and 5% CO2. Before OCR measurements, the cells were washed twice in DMEM, devoid of sodium bicarbonate (pH adjusted with NaOH), and incubated in a BOD incubator at 37°C for 1 h. The cells were in DMEM without FBS and sodium bicarbonate at a final volume of 450 μl per well. The culture medium controls contained only the medium without cells. The plate with cells was then placed into a Seahorse XFe24 Analyzer (Agilent, Santa Clara, CA), and the OCR measurements were carried out at 37°C. The sensor cartridge was hydrated overnight before using for OCR measurements. After three readings for basal OCR, 50 μl of 100 mM DTT was injected into different wells to obtain a final concentration of 10 mM DTT. For 0 mM DTT controls, 50 μl of water was injected into wells. Three technical replicates were set for each condition in each experiment. Two independent experiments were carried out.

Quantification and statistical analysis

The statistical analysis was performed with Prism 8 (GraphPad). All error bars represent mean ± standard deviation (SD) unless otherwise indicated. The two‐sample t‐test was used when needed, and the data were judged to be statistically significant when P < 0.05. In the figures, asterisks (*) denote statistical significance as follows: *P < 0.05, **P < 0.01, ***P < 0.001, as compared with the appropriate controls.

Author contributions

Ravi: Conceptualization; data curation; formal analysis; validation; investigation; visualization; methodology. Ajay Kumar: Resources; data curation; formal analysis; validation; investigation; methodology. Shalmoli Bhattacharyya: Supervision. Jogender Singh: Conceptualization; resources; data curation; formal analysis; supervision; funding acquisition; validation; investigation; visualization; writing – original draft; project administration; writing – review and editing.

Disclosure and competing interests statement

The authors declare that they have no conflict of interest.

Supporting information

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Source Data for Figure 6

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Acknowledgements

We thank Shweta Saini (IISER Mohali) for assistance with C. elegans forward genetic screens and Aditi Bhagat (PGIMER Chandigarh) for assistance with cell culture experiments. Some strains used in this study were provided by the Caenorhabditis Genetics Center (CGC), which is funded by the NIH Office of Research Infrastructure Programs (P40 OD010440). This work was supported by the Ramalingaswami Re‐entry Fellowship (Ref. No. BT/RLF/Re‐entry/50/2020) awarded by the Department of Biotechnology, India, Science and Engineering Research Board (SERB) Startup Research Grant (Ref. No. SRG/2020/000022) awarded by DST, India, and IISER Mohali intramural funds.

The EMBO Journal (2023) 42: e114093

Data availability

The whole‐genome sequence data for JSJ14‐JSJ18 have been submitted to the public repository, the Sequence Read Archive, with BioProject ID PRJNA933286 (https://www.ncbi.nlm.nih.gov/bioproject/PRJNA933286). All data generated or analyzed during this study are included in the manuscript and the Source Data files.

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Associated Data

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    Supplementary Materials

    Appendix

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    Data Availability Statement

    The whole‐genome sequence data for JSJ14‐JSJ18 have been submitted to the public repository, the Sequence Read Archive, with BioProject ID PRJNA933286 (https://www.ncbi.nlm.nih.gov/bioproject/PRJNA933286). All data generated or analyzed during this study are included in the manuscript and the Source Data files.

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