1. Genetics and Genomics
  2. Neuroscience

Avoidance of hydrogen sulfide is modulated by external and internal states in Caenorhabditis elegans

  1. Longjun Pu
  2. Lina Zhao
  3. Jing Wang
  4. Clementine Deleuze
  5. Lars Nilsson
  6. Johan Henriksson
  7. Patrick Laurent  Is a corresponding author
  8. Changchun Chen  Is a corresponding author
  1. Department of Molecular Biology, Umeå University, Sweden
  2. Umeå Centre for Molecular Medicine, Umeå University, Sweden
  3. Wallenberg Centre for Molecular Medicine, Umeå University, Sweden
  4. Laboratory of Neurophysiology, ULB Neuroscience Institute (UNI), Université Libre de Bruxelles (ULB), Belgium
  5. The Laboratory for Molecular Infection Medicine Sweden (MIMS), Umeå University, Sweden
  6. Integrated Science Lab (Icelab), Umeå University, Sweden
https://doi.org/10.7554/eLife.92964.4
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Cite this article
  1. Longjun Pu
  2. Lina Zhao
  3. Jing Wang
  4. Clementine Deleuze
  5. Lars Nilsson
  6. Johan Henriksson
  7. Patrick Laurent
  8. Changchun Chen
(2025)
Avoidance of hydrogen sulfide is modulated by external and internal states in Caenorhabditis elegans
eLife 12:RP92964.
https://doi.org/10.7554/eLife.92964.4
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eLife Assessment

These valuable studies explore the consequences of exposure to the toxin hydrogen sulfide (H2S) on the behavior and physiology of C. elegans. The work finds that behavioral changes evoked by H2S exposure are modulated by several regulatory pathways known to influence chemosensory-evoked locomotor behavior, but there is incomplete data to support the authors' claim of comprehensive mechanistic insight into the consequences of H2S exposure. Nevertheless, the findings may be informative for those studying organismal stress responses and the effects of mitochondrial ROS on behavior and physiology.

https://doi.org/10.7554/eLife.92964.4.sa0

Abstract

Hydrogen sulfide (H2S) acts as an energy source, a toxin, and a gasotransmitter across diverse biological contexts. We use the robust locomotory responses of Caenorhabditis elegans to high levels of H2S to elucidate the molecular mechanisms underlying its acute and adaptive responses. We find that the H2S-evoked behavioral response is shaped by multiple environmental factors including oxygen (O2) levels and nutritional state and is modulated by various pathways such as insulin, TGF-β, and HIF-1 signaling, as well as by input from O2-sensing neurons. Prolonged exposure to H2S activates HIF-1 signaling, leading to the upregulation of stress-responsive genes, including those involved in H2S detoxification. This promotes an adaptive state in which locomotory speed is reduced in H2S, while responsiveness to other stimuli is preserved. In mutants deficient in HIF-1 signaling, iron storage, and detoxification mechanisms, animals display a robust initial response but rapidly enter a sleep-like behavior characterized by reduced mobility and diminished responsiveness to subsequent sensory stimuli. Furthermore, while acute production of mitochondria-derived reactive O2 species (ROS) appears to initiate the avoidance response to H2S, persistently high ROS promotes an adaptive state, likely by activating various stress-response pathways, without substantially compromising cellular H2S detoxification capacity. Taken together, our study provides comprehensive molecular insights into the mechanisms through which C. elegans modulates and adapts its response to H2S exposure.

Introduction

During the late Proterozoic era, the rise in oxygen (O2) levels eliminated hydrogen sulfide (H2S) from most habitats. Yet, low O2 levels, along with high concentrations of hydrogen sulfide (H2S) and carbon dioxide (CO2), can persist in enclosed environments where bacteria actively decompose organic matter (Olson and Straub, 2016). H2S can easily permeate biological membranes and interfere with various cellular processes. One of the most detrimental effects of H2S is the disruption of cellular respiration by remodeling the mitochondrial electron transport chain and inhibiting cytochrome c oxidase (COX; Cooper and Brown, 2008; Khan et al., 1990; Nicholls and Kim, 1982; Romanelli-Cedrez et al., 2024).

The detection and response to chemical signals are crucial for various organisms to interact effectively with their environment. Animals living in H2S-rich environments have evolved mechanisms to either avoid or adapt to these conditions, conferring them survival advantages. Species that encounter periodic increases in H2S often exhibit avoidance behaviors. For example, benthic species undertake daily vertical migrations between sulfidic and non-sulfidic waters (Abel et al., 1987; Salvanes et al., 2011). Certain metazoan species have seized the ecological opportunities presented by H2S-rich environments through the development of cellular adaptations. Live-bearing fish species, for instance, have evolved COX subunits that are more resistant to H2S and more efficient at H2S detoxification (Kelley et al., 2016; Pfenninger et al., 2014).

Free-living nematodes feed on bacteria that thrive in decaying organic matter, such as compost. This complex and dynamic environment, where species like Caenorhabditis elegans are commonly found, may exhibit significant fluctuations in H2S levels over short distances (Adams et al., 1979; Budde and Roth, 2011; Morra and Dick, 1991; Patange et al., 2025; Rodriguez-Kabana et al., 1965; Romanelli-Cedrez et al., 2024). Evidence suggests that C. elegans has evolved tolerance to prolonged H2S exposure by reprogramming gene expression through H2S-induced stabilization of the hypoxia-inducible factor HIF-1 (Budde and Roth, 2010; Budde and Roth, 2011; Horsman et al., 2019; Ma et al., 2012; Miller et al., 2011; Powell-Coffman, 2010; Topalidou and Miller, 2017), and by developing an H2S-resistant electron transport chain (Romanelli-Cedrez et al., 2024). In particular, low concentrations of H2S are not only well tolerated but can also be beneficial. For instance, exposure to 50 ppm H2S has been shown to extend the lifespan and enhance thermotolerance (Fawcett et al., 2015; Miller and Roth, 2007). Moreover, endogenous H2S production in C. elegans modulates its interactions with actinobacteria (Patange et al., 2025). However, exposure to concentrations exceeding 150 ppm proved lethal (Budde and Roth, 2010; Horsman et al., 2019).

In addition to long-term adaptive mechanisms, C. elegans exhibits an acute behavioral response to H2S exposure (Budde and Roth, 2011). In this study, we investigated whether and how the nematode C. elegans acutely avoids environments rich in H2S, how it adapts at the molecular and physiological levels to H2S exposure, and how these adaptations alter the avoidance response.

Results

C. elegans increases locomotory activity in response to acute H2S exposure

In response to aversive cues, C. elegans initiates a pirouette followed by an increase of its locomotory speed to escape the noxious stimuli. When acutely exposed to 150 ppm H2S, the laboratory N2 strain exhibited an increased locomotory speed and enhanced turning behavior, while generally decreasing the frequency of reversals, as a strategy to escape the noxious stimuli (Figure 1A–C, Figure 1—video 1). The maximum speed was reached after 6–8 min in H2S and returned to the baseline over a period of approximately 1 hr (Figure 1D). The reduced locomotory speed after 30 min in 150 ppm H2S was reversible when these animals were immediately challenged with another stimulus, such as near-UV light (Figure 1E), suggesting that neuromuscular function remains preserved. The magnitude and dynamics of the H2S-induced avoidance responses were concentration-dependent. Under our assay conditions, 50 ppm H2S did not elicit an acute speed increase, and responses to 75 ppm were variable. In contrast, exposure to 150 ppm H2S consistently triggered a robust and reproducible escape response (Figure 1F), consistent with earlier reports that H2S at 50 ppm exerts beneficial effects, while H2S at 150 ppm is toxic to C. elegans (Budde and Roth, 2010; Fawcett et al., 2015; Horsman et al., 2019; Miller and Roth, 2007). To investigate whether low levels of H2S might be preferable or even attractive to the animals, we exposed wild-type animals to a linear gradient of H2S (Figure 1G; See methods for details; Bretscher et al., 2008; Chang et al., 2006; Gray et al., 2004). Animals were allowed to move freely on a thin layer of bacteria within a microfluidic chamber containing an H2S gradient ranging from 150 ppm at one end to 0 ppm at the other. As expected, C. elegans robustly avoids 150 ppm H2S but does not accumulate at the extreme end of 0 ppm (Figure 1G and H). The highest proportion of animals was found in the region with approximately 40 ppm H2S. These data suggest that H2S acts as a potent repellent for C. elegans at high concentrations but as an attractant at low levels.

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Acute locomotory responses of C. elegans to hydrogen sulfide.

(A) Changes in the locomotory speed of WT animals (N2 laboratory strain) evoked by a switch from 7% O2 to 150 ppm H2S balanced with 7% O2, followed by a return to 7% O2. (B) Proportion of WT animals undergoing reorientation movements (omega turns) during a switch from 7% O2 to 150 ppm H2S balanced with 7% O2, followed by a return to 7% O2. (C) Fraction of WT animals doing backward locomotion (reversals) during a switch from 7% O2 to 150 ppm H2S balanced with 7% O2, followed by a return to 7% O2. (D) Locomotory speed of WT was recorded for 2 min at 7% O2, followed by 148 min at 150 ppm H2S balanced with 7% O2, and then at 7% O2. (E) Locomotory speed of WT animals in response to near-UV light (435 nm, 0.7 mW/mm2) exposure, before and directly after 30-min preincubation in 150 ppm H2S balanced with 7% O2. Locomotory activity was recorded for 2 min before, during, and after UV light exposure. (F) Locomotory speed of WT animals to different concentrations of H2S balanced with 7% O2. Red bars on the x-axis represent two intervals (4–5 min and 11–12 min, labeled a and b, respectively) used for statistical analysis. **** = p < 0.0001, ** = p < 0.01, * = p < 0.05, ns = not significant, Mann–Whitney U test. (G) Distribution of WT animals in a microfluidic device after 25 min of aerotaxis. The gas inputs and the five chamber sections used for scoring are indicated. (H) Distribution of WT animals in a gradient of H2S (150 ppm to 0 ppm). The bins correspond to different sections of the microfluidic chamber described in (G). N = 4 aerotaxis assays.

Candidate gene screen identifies signaling pathways required for H2S-evoked locomotion

To investigate the molecular mechanisms underlying the H2S-evoked locomotory response in C. elegans, we conducted a candidate gene screen for mutants that failed to increase their locomotory speed upon exposure to an acute rise of H2S level to 150 ppm. Our analysis prioritized genes that are either known to be, or potentially involved in, sensory responses to gaseous stimuli in both C. elegans and mammals (Supplementary file 1). We began by examining mutants affecting globins, guanylate cyclases, and cyclic nucleotide-gated (CNG) channels, which are involved in O2 or CO2 sensing in C. elegans, as well as wild isolates and other previously characterized mutants with defective responses to gas stimuli (Supplementary file 1, Bretscher et al., 2008; Gray et al., 2004; Hallem and Sternberg, 2008; Persson et al., 2009). Given the essential role of K+ channels in acute hypoxia sensing in mammals, we also screened all potassium (K+) channel mutants (Gao et al., 2017; Weir et al., 2005). To explore the neuronal components involved in H2S-evoked avoidance, we assayed mutants with defects in ciliogenesis or cilia-mediated signaling, as well as animals deficient in neurotransmission involving classical neurotransmitters, neuropeptides, or biogenic amines. Finally, considering that mitochondrial function is central to H2S detoxification, we included mitochondrial electron transport chain (ETC) mutants and key factors implicated in H2S and superoxide clearance. Together, these functionally diverse categories allowed us to survey a wide spectrum of potential mechanisms contributing to the behavioral response to H2S.

In the screen, a collection of mutants displayed reduced speed response, whereas only a few exhibited a lack of omega-turn and/or reversal responses to H2S (Supplementary file 1). The reversal inhibition upon H2S exposure and its rebound following H2S withdrawal appeared to vary substantially across experiments for N2, suggesting that differences in population density, food availability, and animals’ physiological state prior to the assay may matter. Nevertheless, our screen revealed that signaling pathways, including cGMP, insulin, and TGF-β signaling, contributed to H2S-evoked avoidance (Supplementary file 1). A naturally occurring variant of the neuropeptide receptor gene npr-1, previously implicated in the modulation of O2 and CO2 responses, also affected locomotory speed in H2S (Supplementary file 1). In addition, mitochondrial components, including those involved in respiration and H2S detoxification, played critical roles in triggering and maintaining escape behavior in H2S. In contrast, we did not find evidence that globins, K+ channels, or biogenic amine signaling contributed significantly to the behavioral avoidance to H2S (Supplementary file 1).

cGMP signaling in ASJ cilia contributes to H2S avoidance

One group of mutants exhibiting reduced locomotory speed in H2S had defects in ciliogenesis (Figure 2A–C). This group included daf-19 mutants lacking cilia, dyf-3 mutants with reduced cilia length, and dyf-7 mutants in which ciliogenesis occurs but the cilia are not anchored to their sensilla and are not exposed to the external environment (Heiman and Shaham, 2009; Murayama et al., 2005; Starich et al., 1995; Swoboda et al., 2000). These findings suggest that ciliogenesis and exposure of cilia to the external environment are essential for animals to escape H2S exposure. In addition, these mutants exhibited consistently high reversal rate, likely associated with their reduced locomotory speed (Figure 2—figure supplement 1A–F). Cilia-mediated sensory responses often involve the activation of guanylate cyclases and the opening of cGMP-gated channels (Ferkey et al., 2021). We observed significantly attenuated locomotory speed in response to H2S in tax-4 or tax-2 mutants, which lack key subunits of cilia-enriched cGMP-gated channels (Figure 2D and E). In a comprehensive survey of all guanylate cyclase mutants, the daf-11 strain emerged as the only mutant showing speed response defects in the avoidance of H2S (Figure 2D, Supplementary file 1). Similar to the cilia mutants, animals deficient in daf-11 had low speed, attenuated omega turns, and constantly high reversal rate (Figure 2D, Figure 2—figure supplement 1G, H). Selective expression of daf-11 in ASJ neurons, but not in other neurons, partially restored the speed response to acute H2S (Figure 2F, Figure 2—figure supplement 2A–D). In addition, the H2S response defect of tax-4 mutants was also rescued by selective expression of its cDNA in ASJ neurons (Figure 2G), suggesting that the receptor guanylate cyclase DAF-11 and the CNG channel TAX-4 act at least partially in ASJ neurons to promote acute avoidance to H2S. To validate the role of ASJ neurons in H2S responses, we blocked neurotransmission from ASJ using the catalytic domain of Tetanus Toxin (TeTx), which impairs neurosecretion by specifically cleaving synaptobrevin (Schiavo et al., 1992). Cell-specific expression of TeTx in ASJ significantly inhibited the speed response to H2S (Figure 2H). Surprisingly, using the Ca2+ sensor GCaMP6s to monitor a direct response of ASJ neurons to H2S, we observed no H2S-induced Ca2+ transients under any of the tested conditions, whereas CO2-evoked Ca2+ increases were readily detected (Figure 2I, Figure 2—figure supplement 2E). These findings suggest that although ASJ neuronal activity and neurosecretion contribute to the H2S responses, ASJ neurons are unlikely to play a role in directly detecting H2S.

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Neurosecretion from ASJ neurons contributes to H2S avoidance.

(A–H) Locomotory speed responses to a switch from 7% O2 to 150 ppm H2S balanced with 7% O2 for animals of the indicated genotype: WT and daf-19(m86)(A); WT and dyf-3(m185)(B); WT and dyf-7(m539)(C); WT, daf-11(m47), and tax-4(p678)(D); WT and tax-2(p691) (E); WT, daf-11(m47), and transgenic daf-11(m47) expressing daf-11 genomic DNA in ASJ neurons (two independent lines) (F); WT, tax-4(p678), and transgenic tax-4(p678) expressing tax-4 cDNA in ASJ neurons (two independent lines) (G); WT and transgenic WT expressing the catalytic domain of the tetanus toxin (TeTx) in ASJ neurons (H). For the comparison of acute locomotory speed responses between strains, red bars on the x-axis represent two intervals (4–5 min and 11–12 min, labeled a and b, respectively, in panels D, F, and G) used for statistical analysis. **** = p < 0.0001, *** = p < 0.001, ** = p < 0.01, * = p < 0.05, ns = not significant, Mann–Whitney U test. (I) Calcium transients evoked in ASJ neurons of WT animals in response to a switch from 7% O2 to 150 ppm H2S balanced with 7% O2 for animals (N=42). The slow decline in GCaMP6s fluorescence is due to photobleaching.

Starvation modulates H2S avoidance

In the candidate gene screen, another set of mutants with reduced speed responses to H2S had previously been shown to be defective in response to CO2 (Hallem and Sternberg, 2008). Specifically, the H2S-evoked speed response was reduced in mutants deficient in nutrient-sensitive signaling, including the insulin receptor DAF-2, the TGF-β ligand DAF-7, and the nuclear hormone receptor NHR-49 (Figure 3A–C, Figure 3—figure supplement 1A). Similarly, mutants lacking other CO2 response modulators, such as the calcineurin subunits TAX-6 and CNB-1, also failed to respond to H2S (Figure 3—figure supplement 1B, C). In addition to the abolished speed response, we also observed that the H2S-evoked omega-turn response was nearly abolished in daf-2 mutants, and a high proportion of animals exhibited persistently high reversal rate regardless of H2S stimulation (Figure 3—figure supplement 1D–F). The H2S avoidance defects observed in daf-2 and daf-7 single mutants were fully suppressed in daf-2; daf-16 and daf-7; daf-3 double mutants (Figure 3A and B). Therefore, it is likely that insulin and TGF-β signaling modulate H2S responses by regulating the expression of relevant genes via DAF-16 and DAF-3 transcription factors, respectively. Given that H2S-evoked speed response was modulated by the nutrient-sensitive pathways, we further explored whether the locomotory response to H2S was sensitive to the nutrient state. Similar to the response to CO2 (Bretscher et al., 2008; Hallem and Sternberg, 2008), we observed that a 24-hr period of starvation suppressed the speed response to H2S (Figure 3D). However, the CO2 sensor GCY-9 was dispensable for H2S-evoked avoidance (Figure 3—figure supplement 1G–I). These findings suggest that while acute avoidance of CO2 and H2S share common modulatory pathways, the responses are initiated through distinct mechanisms.

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Insulin, TGF-β, and high O2 signaling pathways antagonize H2S avoidance.

(A, B) Locomotory speed responses to a switch from 7% O2 to 150 ppm H2S balanced with 7% O2 for animals of the indicated genotype: WT, daf-2(e1370), daf-16(mgDf47), and daf-2(e1370); daf-16(mgDf47) double mutants (A); WT, daf-7(e1372), daf-3(mgDf90), and daf-7(e1372); daf-3(mgDf90) double mutants (B). For the daf-2 assays, WT and daf-2 mutants were maintained at 15 °C. L4 animals were picked and shifted to 25 °C until day-1 adults, then assayed at room temperature. (C) Hypothetical model for the regulation of H2S avoidance by a starvation program involving the insulin, TGF-β, and NHR-49 pathways. (D) Locomotory speed response to a switch from 7% O2 to 150 ppm H2S balanced with 7% O2 in fed and starved WT animals. (E, F) Locomotory speed responses to a switch from 7% O2 to 150 ppm H2S balanced with 7% O2 for animals of the indicated genotype: WT and wild isolates (CB4858 and CB4855) (E); WT and npr-1(ad609) (F). (G) Hypothetical model showing how O2 regulates H2S avoidance via the inhibitory RMG circuit activity, which is modulated by O2 sensory inputs and NPR-1 signaling. (H–L) Locomotory speed responses to a switch from 7% O2 to 150 ppm H2S balanced with 7% O2 for animals of the indicated genotype: WT, npr-1(ad609), and transgenic npr-1(ad609) expressing npr-1 in RMG neurons using Cre-LoxP system (Macosko et al., 2009) (H); WT, npr-1(ad609), gcy-35(ok769), and npr-1(ad609); gcy-35(ok769) double mutants (I); WT, npr-1(ad609), tax-4(p678) and npr-1(ad609); tax-4(p678) double mutants (J); WT, npr-1(ad609), qaIs2241(genetic ablation of AQR, PQR, and URX neurons) and npr-1(ad609); qaIs 2241 (K); WT, npr-1(ad609), npr-1(ad609); gcy-35(ok769) double mutants and transgenic npr-1(ad609); gcy-35(ok769) double mutants expressing gcy-35 cDNA under gcy-37 promoter, which drives gcy-35 expression in O2 sensing neurons (L). For the comparison of acute locomotory responses between strains, red bars on the x-axis represent two intervals (4–5 min and 11–12 min, labeled a and b, respectively, in panel E) used for statistical analysis. ****=p < 0.0001, ***=p < 0.001, **=p < 0.01, *=p < 0.05, ns = not significant, Mann–Whitney U test.

The O2 sensing circuit antagonizes H2S avoidance in wild isolates

Wild strains of C. elegans thrive in environmental niches characterized by variable O2, CO2, and H2S levels (Adams et al., 1979; Bretscher et al., 2008; Budde and Roth, 2011; Gea et al., 2004; Hallem and Sternberg, 2008; Morra and Dick, 1991; Oshins et al., 2022; Patange et al., 2025; Rodriguez-Kabana et al., 1965). Compared to the laboratory reference strain N2, wild isolates display robust responses to high O2 but reduced sensitivity to CO2 stimulation (Beets et al., 2020; Bretscher et al., 2008; Carrillo et al., 2013; Hallem and Sternberg, 2008; Kodama-Namba et al., 2013; McGrath et al., 2009). It is thought that the unique O2 and CO2 responses of N2 strain evolved during its domestication on solid media at atmospheric gas concentrations (Sterken et al., 2015). We hypothesized that wild C. elegans strains might also exhibit an attenuated response to H2S, and therefore included a set of wild strains in our candidate gene screen. Consistent with this hypothesis, wild isolates including CB4855, CB4856, and CB4858 exhibited reduced locomotory speed in H2S (Figure 3E, Figure 3—figure supplement 1J).

A naturally occurring variation in the neuropeptide receptor NPR-1 predominantly determines the differences in locomotory responses to O2 and CO2 between the laboratory N2 strain and wild isolates. The NPR-1(215 F) variant found in wild isolates is less active than the NPR-1(215 V) found in N2 animals (de Bono and Bargmann, 1998). We wondered whether the variations in the npr-1 gene also contribute to differences in the acute response to H2S. The absence of npr-1 was sufficient to reduce the speed response to H2S, suggesting that active NPR-1 signaling promotes H2S avoidance (Figure 3F). NPR-1(215 V) primarily acts within the RMG interneurons to modulate responses to environmental stimuli by inhibiting signal output from these neurons (Figure 3G; Laurent et al., 2015; Macosko et al., 2009). Cell-specific expression of the highly active npr-1(215 V) variant in RMG neurons effectively restored the H2S-evoked locomotory speed in npr-1 null mutant worms (Figure 3H). Furthermore, optogenetic stimulation of RMG interneurons using channelrhodopsin-2 (ChR2) significantly dampened the locomotory response to H2S in N2 animals, suggesting that high RMG activity inhibits H2S responses (Figure 3—figure supplement 1K).

 The perception of 21% O2 involves the soluble guanylate cyclases GCY-35/GCY-36 and the downstream cyclic nucleotide-gated (CNG) channels TAX-4/TAX-2 in the URX, AQR, and PQR O2-sensing neurons, which relay the signals to RMG (Busch et al., 2012; Cheung et al., 2005; Couto et al., 2013; Gray et al., 2004; Laurent et al., 2015; Persson et al., 2009; Zimmer et al., 2009). We investigated whether disrupting the O2 sensing machinery would have the same effect as inhibiting RMG signaling pathway by NPR-1(215 V). Mutating gcy-35 and tax-4, or genetically ablating the O2 sensing neurons URX, AQR, and PQR by cell-specific expression of the pro-apoptotic gene egl-1 restored H2S-evoked speed response in npr-1 null mutants (Figure 3I–K). Specific expression of gcy-35 cDNA in O2 sensing neurons was sufficient to repress this response to H2S in npr-1; gcy-35 double mutants (Figure 3L). Finally, enhancing the presynaptic activity of URX, AQR, and PQR neurons by the expression of a gain-of-function PKC-1(A160E) (Dekker et al., 1993; Hiroki et al., 2022) significantly dampened H2S-induced locomotory speed (Figure 3—figure supplement 1L). Taken together, these observations indicate that activation of the O2-sensing circuit inhibits the neural pathway required for H2S-evoked avoidance response.

H2S exposure reprograms gene expression in C. elegans

Wild isolates that were relatively insensitive to H2S exposure (Figure 3E, Figure 3—figure supplement 1J) likely evolved adaptations to persist in environments where H2S levels may transiently increase (Budde and Roth, 2011; Patange et al., 2025; Rodriguez-Kabana et al., 1965; Romanelli-Cedrez et al., 2024). These adaptations to H2S are mediated, at least in part, by the reprogramming of gene expression (Miller et al., 2011). To identify the genes whose expression was induced by prolonged H2S exposure, we conducted a comparative analysis of transcriptome profiles in animals exposed to 50 ppm or 150 ppm H2S for 1 hr, 2 hr, or 12 hr. RNA-seq analysis revealed that exposure to either 50 ppm or 150 ppm H2S for one hour was sufficient to trigger significant changes in gene expression, with 518 or 304 genes showing differential expression, respectively (Figure 4A, Supplementary file 2). Genes induced by H2S exposure for 1 or 2 hr displayed considerable overlap (Figure 4B, Figure 4—figure supplement 1A). Gene Ontology (GO) analysis revealed that biological processes such as defense against bacteria and cysteine biosynthesis from L-serine were significantly enriched after H2S exposure (Figure 4C and D, Figure 4—figure supplement 1B, C). As expected, we observed a robust induction of genes involved in H2S detoxification (Figure 4E and F; Horsman and Miller, 2016; Miller et al., 2011; Niu et al., 2011; Vora et al., 2022). H2S is detoxified in mitochondria by sulfide:quinone oxidoreductase SQRD-1 to produce persulfide, which is then metabolized by sulfur dioxygenase ETHE-1 to generate sulfite. Thiosulfate transferase further oxidizes sulfite to produce thiosulfate (Hildebrandt and Grieshaber, 2008). Glutathione S-transferases (GSTs) remove accumulated sulfur molecules during H2S oxidation (Jackson et al., 2012). Notably, gst-19 and sqrd-1 were among the most significantly upregulated genes after one or two hours of exposure to either 50 ppm or 150 ppm H2S, while ethe-1 showed a weak increase (Figure 4E and F, Figure 4—figure supplement 1D, E, Supplementary file 2).

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Prolonged H2S exposure reprograms gene expression in C. elegans.

(A) Venn diagram displaying the number of differentially expressed genes after 1 hour exposure to 50 ppm or 150 ppm H2S balanced with 7% O2 in WT animals (adjusted p<1e-10). (B) Venn diagram displaying the number of differentially expressed genes after 1-, 2-, and 12 hr exposure to 150 ppm H2S balanced with 7% O2 in WT animals (adjusted p<1e-10). (C, D) Significantly enriched GO categories for differentially expressed genes with adjusted p<1e-10 in WT animals exposed to 50 ppm (C) or 150 ppm (D) H2S balanced with 7% O2 for 1 hr. (E, F) Volcano plots showing the differentially expressed genes with adjusted p<1e-10 in WT animals exposed to 50 ppm (E) or 150 ppm (F) H2S balanced with 7% O2 for 1 hr. A set of genes involved in H2S detoxification, cysteine metabolism, and stress response was highlighted in red.

H2S exposure is known to activate the HIF-1 pathway (Budde and Roth, 2010; Budde and Roth, 2011; Horsman et al., 2019; Ma et al., 2012; Miller et al., 2011; Powell-Coffman, 2010; Topalidou and Miller, 2017). In line with this, a proportion of genes with HIF-1 regulated promoters displayed increased expression after 1 or 2 hr of H2S exposure (Figure 4—figure supplement 1F). Among these were the aforementioned detoxifying enzymes GST-19, SQRD-1, and ETHE-1, as well as CYSL-1, CYSL-2, and CDO-1, which are involved in cysteine metabolism (Supplementary file 2). Consistent with earlier studies, we also observed differential expression of a set of SKN-1 targets in response to chronic H2S exposure (Figure 4—figure supplement 1G; Horsman et al., 2019; Miller et al., 2011; Niu et al., 2011). This included the heat shock protein genes hsp-16.2 and hsp-16.41 (Figure 4E and F, Figure 4—figure supplement 1D, E). Surprisingly, we detected a rapid increase in the expression of genes associated with intracellular H2S production, including the methanethiol oxidase-encoding gene semo-1 and the mercaptopyruvate sulfurtransferase-encoding gene mpst-3 (Figure 4E and F, Figure 4—figure supplement 1D, E; Philipp et al., 2022; Qabazard et al., 2014). Elevated expression of 3-mercaptopyruvate sulfurtransferase has similarly been reported in sulfide spring fish, suggesting a conserved response mechanism to H2S (Kelley et al., 2016; Mathew et al., 2011; Qabazard et al., 2014). However, the functional implications of increased expression of H2S synthesis genes remain unclear.

When exposed to H2S for 12 hr, the differentially expressed genes showed reduced overlap with those identified at earlier time points (Figure 4B, Figure 4—figure supplement 1A), suggesting that distinct defense strategies may be employed at various stages of H2S exposure. Furthermore, substantial overlap was observed between the sets of genes induced by 50 ppm and 150 ppm H2S after 1 hr or 2 hr exposure. However, this overlap decreased significantly after 12 hr exposure (Figure 4—figure supplement 1H–J). Additionally, we noticed that 150 ppm H2S specifically triggered the expression of the HIF-1 regulator rhy-1 and the HIF-1 target nhr-57 at all exposure time points, a response that was less pronounced at 50 ppm H2S (Figure 4E and F, Figure 4—figure supplement 1D, E, K and L). Together, these findings suggest that multiple defense mechanisms are induced to adapt to the stress imposed by H2S, which can diverge over time and at different H2S concentrations.

Locomotory speed response to H2S is modulated by HIF-1-induced detoxification

Given that HIF-1 signaling is robustly activated shortly after H2S exposure (Figure 4—figure supplement 1F; Budde and Roth, 2010; Budde and Roth, 2011; Miller et al., 2011), we next sought to determine how HIF-1 signaling might contribute to H2S-evoked avoidance response and began by examining the effects of HIF-1 stabilization. Acclimating animals in 1% O2 for 12 hr or longer significantly decreased the speed response to H2S, indicating that HIF-1 stabilization inhibits the H2S-evoked avoidance behavior (Figure 5A). To confirm this observation, we examined several mutants with stabilized HIF-1. The conserved proline-4-hydroxylase PHD/EGL-9 and the von Hippel-Lindau (VHL) tumor suppressor protein are required for HIF-1 degradation (Kaelin and Ratcliffe, 2008; Semenza, 2010). Disruption of either egl-9 or vhl-1 leads to HIF-1 stabilization under normoxic conditions. Similar to the effect of prolonged hypoxia, both mutants exhibited a markedly reduced locomotory speed in H2S (Figure 5B). Interestingly, the omega-turn and reversal responses to H2S were also substantially inhibited in egl-9 mutants (Figure 5—figure supplement 1A, B). In contrast, the locomotory speed response to 1% O2 remained largely unaffected in egl-9 deficient animals (Figure 5—figure supplement 1C), suggesting a selective impairment of the H2S response. In addition, expressing the non-degradable forms of HIF-1, hif-1(P621A or P621G), under the pan-neuronal unc-14 promoter or under the endogenous hif-1 promoter, was sufficient to inhibit H2S-evoked speed responses (Figure 5C). Together, these findings indicate that the activation of HIF-1 signaling during prolonged H2S or hypoxia exposure mediates an adaptive response associated with reduced behavioral responses to H2S, and that HIF-1 stabilization in neurons alone is sufficient to promote this behavioral adaptation.

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Acute response to H2S is modulated by HIF-1 signaling.

(A) Locomotory speed responses of WT animals to a switch from 7% O2 to 150 ppm H2S balanced with 7% O2 after incubation of the animals for 0, 12, 24, or 36 hr in 1% O2. (B, C) Locomotory speed responses to a switch from 7% O2 to 150 ppm H2S balanced with 7% O2 for animals of the indicated genotype: WT, egl-9(sa307), and vhl-1(ok161) (B); WT and transgenic animals expressing the non-degradable form of HIF-1 (P621A or P621G) under the pan-neuronal unc-14 promoter or the endogenous hif-1 promoter, respectively (C). (D–F) Avoidance responses of WT and hif-1(ia4) to a switch from 7% O2 to 150 ppm H2S balanced with 7% O2, including locomotory speed (D), reorientation (omega-turn) (E), and reversal (F). (G, H) Locomotory speed responses to a switch from 7% O2 to 150 ppm H2S balanced with 7% O2 for animals of the indicated genotype: WT, egl-9(sa307), hif-1(ia4), and egl-9(sa307); hif-1(ia4) double mutants (G); WT, vhl-1(ok161), hif-1(ia4), and vhl-1(ok161); hif-1(ia4) double mutants (H). (I) Average locomotory speed during 2 min of exposure to near-UV light for WT, hif-1(ia4), and egl-9(sa307) before (Ctrl = control) and after 30 min of exposure to 150 ppm H2S balanced with 7% O2. (J–L) Locomotory speed responses to a switch from 7% O2 to 150 ppm H2S balanced with 7% O2 for animals of the indicated genotype: WT and sqrd-1(tm3378) (J); WT and ethe-1(yum2895) (K); WT, cysl-1(ok762), cysl-2(ok3516), and cysl-3(yum4) (L). For the comparison of acute locomotory responses between strains, red bars on the x-axis represent two intervals (4–5 minutes and 11–12 minutes, labeled a and b, respectively, in panels A, B, C, and L) used for statistical analysis. ***=p < 0.001, **=p < 0.01, *=p < 0.05, ns = not significant, Mann–Whitney U test.

On the other hand, animals lacking HIF-1 signaling have previously been shown to display increased sensitivity to H2S and become paralyzed rapidly upon exposure (Budde and Roth, 2010; Horsman et al., 2019). Consistent with this, hif-1 mutants exhibited a brief increase in speed upon H2S exposure, which rapidly declined to baseline levels (Figure 5D). They also displayed a stronger initial omega-turn response and greater inhibition of the reversal rate than WT (Figure 5E and F), confirming that animals deficient in HIF-1 signaling are sensitized to H2S exposure. This transient avoidance behavior was also observed in egl-9; hif-1 and vhl-1; hif-1 double mutants (Figure 5G and H), suggesting that this phenotype is linked to the absence of HIF-1 signaling. Importantly, hif-1 mutants displayed a robust speed response to other stimuli, such as near-UV light and 1% O2 stimulation (Figure 5I, Figure 5—figure supplement 1D), suggesting that the brief avoidance response is specific to H2S rather than reflecting a general locomotory defect. Further supporting increased sensitivity to H2S in hif-1 mutants, the locomotory speed response of hif-1 mutants to subsequent near-UV light stimulation was nearly abolished after 30 min of H2S exposure, whereas egl-9 mutants and WT animals remained fully responsive (Figure 5I). The reduced movement and lack of responsiveness to other stressors in hif-1 mutants resemble the stress-induced sleep-like behavior previously observed in response to noxious heat (Byrne Rodgers and Ryu, 2020). Prompted by this observation, we next explored whether HIF-1-induced detoxification genes were also required to maintain H2S-evoked behavioral responses (Figure 5—figure supplement 1E). Similar to hif-1 mutants, sqrd-1 and ethe-1 mutants displayed brief responses to H2S, with locomotory activity rapidly returning to baseline levels (Figure 5J and K, Figure 5—figure supplement 1F–I). However, both mutants responded normally to 1% O2, suggesting a specific hypersensitivity to H2S toxicity (Figure 5—figure supplement 1J). Furthermore, we examined the contribution of additional genes whose expression was upregulated in response to H2S, including the sulfhydrylase/cysteine synthase-encoding genes cysl-1, cysl-2, and cysl-3, as well as semo-1. Disrupting any of these genes phenocopied the responses observed in hif-1, ethe-1, or sqrd-1 mutants, while they displayed normal responses to 1% O2 (Figure 5L, Figure 5—figure supplement 1D, K–P). These observations suggest that while the HIF-1-induced H2S detoxification system is not essential for initiating the response to H2S, it is required to maintain animals’ locomotory activity in the presence of H2S.

Labile iron pool sustains the locomotory activity in H2S

Among the most differentially regulated genes upon exposure to H2S, we observed consistent downregulation of ftn-1 and, to a lesser extent, ftn-2, whereas smf-3 expression was increased under specific H2S conditions (Figure 6A and B, Figure 6—figure supplement 1A–D). The ferritin-encoding genes ftn-1 and ftn-2, along with the iron transporter smf-3, are critical for maintaining intracellular labile iron homeostasis in C. elegans (Ackerman and Gems, 2012; Gourley et al., 2003; Rajan et al., 2019; Romney et al., 2011), suggesting that H2S exposure may interfere with cellular iron storage. It has been shown that disrupting ftn-1 increases labile iron pools in the cytoplasm, while its overexpression reduces these levels (Anderson and Leibold, 2014; Romney et al., 2011). We found that ftn-1 mutants had enhanced speed response to H2S, whereas ftn-1 overexpression attenuated H2S-induced speed increase without damaging acute response to 1% O2 (Figure 6C and D, Figure 6—figure supplement 1E). In addition, smf-3 mutants, which impair iron uptake, exhibited a reduced speed response to H2S (Figure 6D). These results support the idea that H2S exposure disrupts iron homeostasis.

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Labile iron pool sustains the locomotory activity in H2S.

(A, B) Volcano plots showing the relative expression of the genes involved in the regulation of iron homeostasis in WT animals after 2 hr exposure in 50 ppm (A) or 150 ppm (B) H2S balanced with 7% O2. (C–G) Locomotory speed responses to a switch from 7% O2 to 150 ppm H2S balanced with 7% O2 for animals of indicated genotypes or treatments: WT and animals overexpressing ftn-1 genomic DNA under its own promoter (#1 and #2 indicate two independent lines) (C); WT, smf-3(ok1305), and ftn-1(ok3625) (D); WT, and WT pretreated with 100 μM 2,2′-Bipyridyl (BP) or with 5 mg/ml ferric ammonium citrate (FAC) in the presence of food for 16 hr (E); WT, hif-1(ia4), and hif-1(ia4) mutants pretreated with 5 mg/ml ferric ammonium citrate (FAC) for 16 hr (F); WT, hif-1(ia4), ftn-1(ok3625), and hif-1(ia4); ftn-1(ok3625) double mutants (G). (H) Hypothetical model of the interactions between labile iron pool and HIF-1 signaling. For the comparison of acute locomotory speed responses between strains or conditions, red bars on the x-axis represent two intervals (4–5 min and 11–12 min, labeled a and b, respectively, in panels C, D, and E) used for statistical analysis. ****=p < 0.0001, ***=p < 0.001, **=p < 0.01, *=p < 0.05, ns = not significant, Mann–Whitney U test.

To complement our genetic analysis, we pharmacologically depleted labile irons. In the presence of the iron chelator 2,2’-dipyridyl (BP), the H2S-evoked speed response was fully lost (Figure 6E), whereas locomotory speed in response to hypoxia was maintained (Figure 6—figure supplement 1F). We noticed that, similar to hif-1 mutants, both smf-3 mutants and BP-treated animals exhibited a strong initial omega-turn response to H2S (Figure 6—figure supplement 1G, I), suggesting that these animals are hypersensitive to H2S toxicity. One possibility is that low iron levels compromise H2S detoxification capacity, as the detoxification enzyme ETHE-1 requires iron for its activity (Kabil and Banerjee, 2012; Pettinati et al., 2015). By contrast, increasing iron levels by feeding ferric ammonium citrate (FAC) or by disrupting ftn-1 significantly sustained animals’ locomotory speed in H2S (Figure 6D and E), likely by delaying H2S-mediated iron depletion. However, the reorientation and reversals were not significantly affected (Figure 6—figure supplement 1G–J). This impact of FAC supplementation and ftn-1 disruption on speed response was largely lost in the absence of hif-1 (Figure 6F and G), suggesting that the effects of iron require HIF-1-dependent detoxification (Figure 6H). Yet, increased iron availability modestly improved the locomotory activity of hif-1 mutants in H2S (Figure 6F and G), presumably by compensating for the low iron levels in hif-1 mutant animals (Rajan et al., 2019; Romney et al., 2011).

Mitochondrial function modulates H2S-evoked locomotion responses

The mitochondrial electron transport chain (ETC) is not only the primary target of H2S toxicity, but is also an essential component of the H2S detoxification pathways (Cooper and Brown, 2008; Khan et al., 1990; Nicholls and Kim, 1982; Romanelli-Cedrez et al., 2024). Based on this, we hypothesized that ETC plays a central role in the locomotory response to H2S. Indeed, mutating the genes critical for ETC function, including gas-1, clk-1, mev-1, and isp-1, impaired the H2S-evoked speed and reversal responses (Figure 7A–D, Figure 7—figure supplement 1A–D). These data suggest either that dysfunctional ETC induces an adaptive response to cellular stressors or that a functional ETC is required to trigger and support H2S-evoked avoidance behavior. Consistent with the later model, exposing animals to rotenone, an ETC complex I inhibitor, robustly inhibited the speed responses to H2S at all time points we tested (Figure 7E). Similar to ETC mutants, WT animals exposed to rotenone for 2 hr also showed reduced basal locomotory activity (Figure 7E).

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Mitochondrial function is required for acute response to H2S.

(A–D) Locomotory speed responses to a switch from 7% O2 to 150 ppm H2S balanced with 7% O2 for animals of the indicated genotype: WT and gas-1(fc21) (A); WT and clk-1(qm30) (B); WT and mev-1(kn1) (C); WT and isp-1(qm150) (D). (E) Locomotory speed responses to a switch from 7% O2 to 150 ppm H2S balanced with 7% O2 for WT animals pretreated with 10 μM rotenone for 0 min, 10 min, 30 min, 60 min, or 120 min. For the comparison of acute locomotory speed responses between strains or conditions, red bars on the x-axis represent two intervals (4–5 min and 11–12 min) used for statistical analysis. ***=p < 0.001, **=p < 0.01, *=p < 0.05, ns = not significant, Mann–Whitney U test. (F–G) Locomotory speed responses to a switch from 7% O2 to 150 ppm H2S balanced with 7% O2 for animals of the indicated genotype: WT, sod-1(tm776), sod-2(ok1030), and sod-3(tm760) (F); WT, sod-4(gk101), sod-5(tm1146), and sod-1(tm783); sod-2(ok1030); sod-3(tm760); sod-4(gk101); sod-5 (tm1146) (G). For the comparison of acute locomotory speed responses between strains or conditions, red bars on the x-axis represent the time interval (6–10 min) used for statistical analysis. ***=p < 0.001, **=p < 0.01, *=p < 0.05, ns = not significant, Mann–Whitney U test. (H) Hypothetical model of the role of mitochondria in response to toxic levels of H2S. In this model, mitochondria play a dual role in H2S-evoked avoidance behavior. A transient burst of mitochondrial ROS triggered by high H2S levels initiates locomotory avoidance, whereas sustained ROS elevation activates stress-responsive pathways, including HIF-1, NRF2/SKN-1, and DAF-16/FOXO, promoting adaptation to prolonged H2S exposure.

In contrast to prolonged rotenone exposure, we noticed that transient rotenone exposure substantially increased animals’ basal locomotory speed (Figure 7E; Onukwufor et al., 2022), suggesting that acute ETC interruption evokes behavioral avoidance, in a manner similar to that observed with other noxious stimuli. Since rotenone rapidly and persistently stimulates mitochondrial ROS production (Ochi et al., 2016; Ramsay and Singer, 1992; Zorov et al., 2014), the transient locomotory increase during short rotenone exposure (Figure 7E) is likely driven by these ROS bursts, whereas reduced locomotory activity during prolonged exposure is presumably caused by chronically elevated ROS production. As toxic levels of H2S also induce ROS (Jia et al., 2020), this raised the possibility that H2S-evoked behavioral responses were modulated by the mitochondrial ROS. Therefore, we sought to further explore how mitochondrial ROS contributes to speed response to high H2S. We focused on superoxide dismutases (SODs), key enzymes in superoxide detoxification. The C. elegans genome encodes five sod genes: sod-1 and sod-5 encode cytosolic Cu/ZnSODs, sod-2 and sod-3 encode mitochondrial MnSODs, and sod-4 encodes the extracellular Cu/ZnSOD isoforms (Doonan et al., 2008; Zubovych et al., 2010). All sod single mutants exhibited a robust initial response to 150 ppm H2S (Figure 7F and G). Interestingly, the mitochondrial SOD mutants sod-2 and sod-3, as well as sod-5, showed an initial burst of locomotory activity followed by a rapid decline in speed in the presence of H2S (Figure 7F and G), but maintained a high locomotory speed in response to 1% O2 (Figure 7—figure supplement 1E). In the quintuple sod-1; sod-2; sod-3; sod-4; sod-5 mutant, which lacks all superoxide dismutases and presumably accumulates higher ROS levels, the H2S-evoked speed and omega-turn responses were nearly abolished, and the proportion of animals exhibiting reversals remained consistently high, a pattern resembling that of egl-9 and ETC mutants (Figure 7G, Figure 7—figure supplement 1F, G). Similar to sod-2 and sod-3, the quintuple sod mutant exhibited robust speed response to 1% O2 (Zhao et al., 2022). In addition, the quintuple mutants responded normally to near-UV light after 30 min of H2S exposure (Figure 7—figure supplement 1H), indicating that their neuromuscular function remains largely preserved. These observations suggest that either constitutively high ROS levels in quintuple mutants dampen H2S-evoked ROS transients, or they promote adaptation to H2S-induced stress, presumably via activation of stress-responsive pathways such as HIF-1, NRF2/SKN-1, and DAF-16/FOXO, ultimately reducing the locomotory responses (Figure 7H; Lee et al., 2010; Lennicke and Cochemé, 2021; Patten et al., 2010).

Discussion

Animals’ behavior and physiology are profoundly influenced by the environments in which they evolved. The laboratory strain N2, which is adapted to low atmospheric CO2 and H2S concentrations, robustly avoids both gases (Beets et al., 2020; Bretscher et al., 2008; Carrillo et al., 2013; Hallem and Sternberg, 2008; Kodama-Namba et al., 2013; McGrath et al., 2009). Upon acute exposure to H2S above 75 ppm, N2 exhibits an avoidance behavior characterized by reorientation and increased locomotion. This response is significantly reduced in wild isolates. These divergent responses between the laboratory strain and wild isolates are primarily driven by variations in the npr-1 gene, which encodes a neuropeptide receptor, suggesting that C. elegans has undergone rapid evolutionary adaptation to its environment.

The behavioral response to a specific sensory stimulus is usually shaped by an interplay of multiple environmental and physiological cues. We observe that conditions dampening C. elegans’ response to CO2 also impair the response to H2S. Specifically, the speed response to H2S is significantly inhibited at high O2 levels, and in mutants with defective insulin or TGF-β pathway that signal starvation. However, despite these shared modulations, CO2 and H2S responses involve distinct molecular mechanisms. For example, CO2 responses are mediated by the guanylate cyclase GCY-9 in BAG neurons (Hallem et al., 2011), which is dispensable for H2S responses. In addition, acute exposure to H2S induces a delayed acceleration compared to CO2. These data suggest that while responses to the two gases share a downstream neural circuit that can be dynamically modulated by the state of O2 sensing circuit, they are triggered by distinct mechanisms.

In a candidate gene survey, we excluded the potential involvement of globins, K+ channels, biogenic amines, and most guanylate cyclases in H2S responses. However, we observed that H2S avoidance requires the activity of guanylate cyclase DAF-11 in ASJ neurons, as well as neurosecretion from these neurons. Since the DAF-11 pathway and neurosecretion from ASJ neurons regulate developmental programs that modify sensory functions in C. elegans (Murakami et al., 2001), it is not surprising that daf-11 mutants display pleiotropic phenotypes including impaired H2S and CO2 responses (Hallem and Sternberg, 2008). In addition, we did not detect an H2S-evoked calcium transient in ASJ neurons. Therefore, although ASJ neurons and DAF-11 activity are clearly required, how H2S avoidance is triggered remains to be elucidated.

Consistent with previous reports (Horsman et al., 2019; Miller et al., 2011), H2S exposure substantially alters the gene expression in C. elegans, including those involved in H2S detoxification and iron homeostasis pathways. The repression of ftn-1 and induction of smf-3 observed during H2S exposure suggests that H2S depletes intracellular labile iron. Iron is a critical cofactor required for the activity of many enzymes and supports a wide range of cellular processes (Dev and Babitt, 2017; Hentze et al., 2004). One important enzyme that requires labile iron as a cofactor is PHD/EGL-9, which targets HIF-1 for degradation. Changes in iron availability are therefore expected to modulate EGL-9 activity, which in turn alters the expression of HIF-1-regulated genes (Liochev, 1996; Myllyharju and Kivirikko, 1997; Read et al., 2021; Xu and Møller, 2011). However, ETHE-1, a key H2S detoxification enzyme upregulated upon HIF-1 stabilization, also requires iron for its persulfide dioxygenase activity (Kabil and Banerjee, 2012; Pettinati et al., 2015). Therefore, H2S-evoked responses under varying iron availability are determined by the combined effects of labile iron on HIF-1 signaling and on the H2S detoxification enzymes, among other factors. For instance, although reduced iron availability in smf-3 or BP-treated animals may promote the HIF-1 induced detoxification pathways by inactivating EGL-9, the actual enzymatic clearance of H2S is impaired due to low ETHE-1 activity. Supporting this, smf-3 mutants and BP-treated animals display increased sensitivity to H2S, including enhanced initial omega-turn responses and rapid inhibition of locomotion. In contrast, increased iron availability in ftn-1 mutants or FAC-treated animals likely delays iron depletion during H2S exposure, preserving ETHE-1 detoxification capacity and postponing the onset of HIF-1-mediated adaptations that are associated with reduced locomotion. This may explain the sustained high locomotory speed observed in ftn-1 mutants or FAC-treated animals. In the absence of HIF-1, iron supplementation only partially improves the locomotory response to H2S, likely because the H2S detoxification system including ETHE-1 cannot be transcriptionally induced. The modest effect observed may instead reflect correction of iron deficiency in hif-1 mutants.

At concentrations below 50 ppm, H2S is well tolerated or even preferred, with beneficial effects such as improved thermotolerance (Fawcett et al., 2015; Miller and Roth, 2007; Qabazard et al., 2014). Notably, behavioral avoidance is absent below 50 ppm H2S, suggesting that escape behavior is triggered only when H2S is not efficiently detoxified. This led us to hypothesize that animals with enhanced detoxification capacity would show reduced avoidance of otherwise toxic H2S levels. Indeed, the speed response to H2S is attenuated when the detoxification program is upregulated under conditions of constitutive activation of HIF-1, such as in egl-9 or vhl-1 mutants, after prolonged exposure to 1% O2, or when HIF-1 signaling is specifically activated in neurons. These observations support the idea that the initial avoidance response to H2S is triggered by neuronal detection of acute toxicity, while the subsequent decline in speed reflects a rapid activation of adaptive mechanisms, particularly through stabilization of HIF-1. The fact that wild-type animals remain responsive to other stimuli after prolonged H2S exposure suggests that reduced locomotory speed in H2S reflects active desensitization rather than general paralysis. However, persistently high H2S exposure is likely to exhaust cellular defense systems, leading to toxicity and paralysis. By contrast, the rapid loss of locomotion observed in hif-1 mutants and detoxification-defective mutants is mediated by a mechanism distinct from adaptation. Their loss of responsiveness to other stimuli after H2S exposure suggests that these animals likely become sensitized and rapidly intoxicated by H2S due to impaired detoxification. Therefore, H2S exposure promotes a behavioral program that includes an initial reorientation and acceleration responses followed by a progressive adaptation driven by cellular detoxification processes. A similar behavioral program has been observed in C. elegans during noxious heat exposure, which induces short-term heat avoidance followed by long-lasting cytoprotective adaptation and a gradual reduction in avoidance (Byrne Rodgers and Ryu, 2020).

The interaction of H2S with mitochondrial ETC is multifaceted, acting as an electron donor at low concentrations and becoming a potent toxicant at high levels (Szabo et al., 2014), in part by promoting superoxide generation through complex IV inhibition and reverse electron transport at complex I (Cooper and Brown, 2008; Jia et al., 2020; Khan et al., 1990; Nicholls and Kim, 1982; Romanelli-Cedrez et al., 2024). We propose that mitochondria play a dual role in H2S-evoked locomotory avoidance. On one hand, the mitochondrial ETC contributes to H2S detoxification and promotes adaptation. On the other hand, toxic levels of H2S remodel the ETC, leading to increased ROS production, which may serve as a trigger for the avoidance response. Supporting the idea that acute mitochondrial ROS generation initiates avoidance of high H2S levels, short-term rotenone exposure, known to promote mitochondrial ROS formation (Ochi et al., 2016; Ramsay and Singer, 1992; Zorov et al., 2014), substantially increases locomotory speed (Onukwufor et al., 2022). Meanwhile, the speed response to high H2S is fully suppressed by rotenone. This inhibition could result either from excessive mitochondrial ROS generated by rotenone, which may dampen the H2S-triggered ROS spike, or from direct complex I inhibition, which may disrupt other H2S signaling processes required to initiate avoidance. However, persistent mitochondrial ROS production appears to suppress high locomotory speed and inhibit responsiveness to H2S, as observed after 2 hr rotenone exposure, in mitochondrial ETC mutants, and in animals lacking all superoxide dismutases. One likely explanation is that mitochondrial ROS can activate a variety of stress-responsive pathways, including HIF-1, NRF2/SKN-1, and DAF-16/FOXO signaling (Lee et al., 2010; Lennicke and Cochemé, 2021; Patten et al., 2010), priming animals’ adaptation to prolonged stress rather than causing toxicity. This is supported by the observation that even though SOD-deficient animals do not display strong initial locomotory responses to H2S, they remain responsive to other stimuli after 30 min of H2S exposure, suggesting that high ROS levels do not compromise general viability or the H2S detoxification capacity. Therefore, we favor a model in which mitochondrial ROS exert a biphasic effect on H2S-induced avoidance, facilitating H2S avoidance under acute conditions and contributing to locomotory inhibition when it is chronically elevated. Overall, lack of a clear sensory machinery, the slow increase of locomotory speed in H2S (Figure 1D), the rotenone-evoked speed responses, and the strong modulation of H2S responses by mitochondrial ETC inhibition suggest that H2S may not be directly perceived by C. elegans. Instead, acute avoidance to H2S is likely initiated by ROS-induced toxicity.

In summary, this study unveils a novel behavior of C. elegans in their avoidance to H2S. The speed response to H2S is intricately shaped by environmental context, integrating inputs from other sensory cues such as food availability and O2 levels. Our findings suggest that H2S-induced locomotion arises from acute remodeling of mitochondrial ETC and associated ROS production, while highlighting the vital role of the HIF-1-induced detoxification pathways and iron homeostasis in protecting against H2S-induced mitochondrial toxicity. Further work is needed to validate this model and to elucidate the neural circuits mediating the behavioral response to ROS-induced toxicity.

Materials and methods

Strains

The model organism C. elegans was used in this study. The N2 wild-type strain and mutant worms were obtained from the Caenorhabditis Genetics Center and the National BioResources Project Japan and maintained using standard protocols (Brenner, 1974). Strains used in this study are listed in Supplementary file 1 and Supplementary file 3.

Preparation of H2S gas mixture

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Different H2S concentrations were created as previously described (Miller and Roth, 2007). Briefly, the H2S-containing gas mixture was prepared by diluting 5000 ppm H2S in nitrogen (N2) with 7% O2 balanced with N2. The gas flow was tuned by Sierra Smart-Trak 100 mass flow controllers. H2S concentration in the mixture was measured using two H2S detectors (MSA ALTAIR 2 X gas detector for H2S and Clip Single Gas Detector, SDG, CROWCON). The pre-defined gas mixtures of 7% O2, 1% O2, and 5% CO2 with N2 were purchased from Air Liquide Gas AB. 5000 ppm H2S stock in N2 was obtained from Linde Gas AB. Gas mixtures in all experiments were hydrated before use.

Molecular biology

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The Multisite Gateway system (Thermo Fisher Scientific, United States) was used to generate expression vectors. Promoters, including gcy-37 (2.7 kb), trx-1 (1 kb), ftn-1(2 kb), daf-11(3 kb), odr-3(2.7 kb), gpa-11(3 kb), sra-6(3 kb), odr-1(2.4 kb), flp-21(4.1 kb), and ocr-2(2.4 kb) were amplified from N2 genomic DNA and cloned into pDONR P4P1 (Thermo Fisher Scientific) using BP clonase. gcy-35, tax-4, and pkc-1 cDNAs were amplified using the first strand cDNA library as the template, while daf-11 and ftn-1 genomic sequences were amplified using genomic DNA as the template and cloned into pDONR 221 (Thermo Fisher Scientific, 12536017) using BP reaction. To generate the gain-of-function mutation of pkc-1 (A160E), Q5 Mutagenesis Kit (NEB) was used according to manufacturer instructions. All expression vectors were generated using LR reaction. The primer sequences were displayed in Supplementary file 4.

To generate transgenic animals, the daf-11 expression vectors were injected at the concentration of 20 ng/μl supplemented with 50 ng/μl of a coelomocyte co-injection marker (unc-122p::GFP) and 30 ng/μl of 1 kb DNA ladder. For the tax-4 gene, the injection mixtures were prepared using 40 ng/μl of tax-4 expression vectors, 50 ng/μl of a coelomocyte co-injection marker, and 10 ng/μl of 1 kb DNA ladder. The rest of the expression constructs were injected at 50 ng/μl together with 50 ng/μl of a coelomocyte marker.

CRISPR/Cas9 genome editing

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Genes were disrupted using CRISPR/Cas9-mediated genome editing as described (Dokshin et al., 2018; Ghanta and Mello, 2020). The strategy involved the utilization of homology-directed insertion of custom-designed single-strand DNA template (ssODN), which had two homology arms of 35 bp flanking the targeted PAM site. Between two homology arms, a short sequence containing a unique restriction enzyme cutting site as well as in-frame and out-of-frame stop codons was included. The insertion of the ssODN template would delete 16 bases of coding sequence, ensuring the proper gene disruption. To prepare the injection cocktail, 0.5 μl of Cas9 protein (IDT) was mixed with 5 μl of 0.4 μg/μl tracrRNA (IDT, United States) and 2.8 μl of 0.4 μg/μl crRNA (IDT). The mixture was incubated at 37 °C for at least 10 min before 2.2 μl of 1 μg/μl ssODN (or 500 ng dsDNA) and 2 μl of 0.6 μg/μl rol-6 co-injection marker were added. Nuclease-free water was used to bring the final volume to 20 μl. The injection mixture was centrifuged for 2 min before use.

Behavioral assays

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H2S-evoked locomotion activity was monitored as described previously (Laurent et al., 2015; Zhao et al., 2022). Briefly, OP50 bacteria obtained from the Caenorhabditis Genetics Center were seeded on the assay plates 16 hr before use. The border of the bacterial lawn was removed using a PDMS stamp. Day-1 adult hermaphrodites were used in all assays unless otherwise specified. For each assay, 25–30 day-1 adult animals were picked onto assay plates, allowed to settle down for 15 min, and subsequently sealed within microfluidic chambers. A syringe pump (PHD 2000, Harvard Apparatus) was employed to deliver gas mixtures into the microfluidic chamber at a constant flow of 3 ml/min. The rapid switch between different gas mixtures was controlled by Teflon valves coupled with ValveBank Perfusion Controller (AutoMate Scientific). The locomotory activity at different gas mixtures was monitored using a high-resolution camera (FLIR) mounted on a Zeiss Stemi 508 microscope. Videos were captured at a rate of 2 frames per second, starting with a 2-min recording in 7% O2, followed by 10 or 11 min in H2S, and ending with 7% O2. For the long-term recording in H2S, video capture commenced with a 2-min period in 7% O2, followed by a duration of 148 min in 150 ppm H2S, and concluded with a final 2-min interval in 7% O2. For each strain or condition, three to four replicates were performed.

Near-UV light experiments were conducted to assess whether animals remained responsive following a 30-min incubation in 150 ppm H2S balanced with 7% O2. For each experiment, 6-min videos were recorded. The first 2 min captured the baseline locomotion of the animals under white light, followed by 2 min of exposure to near-UV light (435 nm, 0.7 mW/mm2), and concluded with 2 min of white light. 3–4 replicates were performed, with approximately 15 animals recorded per replicate.

The H2S gradient experiment was performed in a PDMS chamber connected to a pump delivering gas at 0.5 ml/min. To establish a gradient ranging from 150 ppm to 0 ppm H2S, we pumped 150 ppm H2S balanced with 7% O2 into one side and 7% O2 into the opposite side. After 25 min, the number of animals in each of the five sections was counted (Figure 1G). Four replicates were performed, each using 30 day-1 adult animals.

To assess the effects of rotenone on H2S-evoked locomotory response, day-1 adult animals were subjected to 10 μM rotenone for 10 min, 30 min, 1 hr, and 2 hr on the drug-containing plates. Subsequently, the rotenone-treated animals were assayed on the standard assay plates without the drug. To explore the impact of ferric ammonium citrate (FAC) (F5879, Sigma-Aldrich) and 2,2′-Bipyridyl (BP) (D216305, Sigma-Aldrich) on behavioral response to H2S, L4 animals were exposed to 100 μM FAC or 5 mg/ml BP for 16 hr in the presence of bacterial food. The FAC or BP-treated animals were assayed on FAC or BP-containing plates, respectively.

Optogenetic experiments were performed as previously outlined (Zhao et al., 2022). Transgenic L4 animals were exposed to 100 μM all-trans retinal (ATR; R2500, Sigma) in the dark for 16 hr. The ATR-fed animals were assayed under continuous illumination of 70 μW/mm2 blue light, which was switched on at the start of recording at 7% O2. Blue light was emitted from an ultra-high-power LED lamp (UHP-MIC-LED-460, Prizmatix), and the light intensity was determined using a PM50 Optical Power Meter (ThorLabs). To minimize the effect of transmitted light on the transgenic animals, a long-pass optical filter was used to eliminate the lights with short wavelengths during picking. All videos of behavioral analysis were analyzed using a home-made MATLAB (RRID:SCR_001622) program Zentracker (https://github.com/wormtracker/zentracker; wormtracker, 2015). For all behavior analyses, at least three assays were performed for each strain with more than 80 worms in total.

Ca2+ imaging

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L4 animals expressing the GCaMP6s sensor in ASJ neurons were picked 24 hr before imaging. The agarose pads were prepared 1 hr before the experiment. Animals were incubated in 15 µl of 10 mM levamisole for 40 min prior to imaging. Between 15 and 20 animals were immobilized using 10 mM levamisole diluted in a concentrated OP50 bacterial suspension and mounted on a 2% agarose pad in M9 buffer on a glass slide. The animals were then covered with a microfluidic chamber. Imaging lasted for 16 min. O2 (7%) was pumped into the microfluidic chamber during the first and last 4 min. From min 4 to 12, a mixture of 7% O2 and 150 ppm H2S was pumped.

Imaging was performed on a Nikon AZ100 microscope equipped with a TwinCam adapter (Cairn Research, UK), fitted with two DMK 33 U monochrome cameras (The Imaging Source, Germany). A 2 x AZ Plan Fluor objective with 1 x zoom was used, with an exposure time of 300 ms. Excitation of GCaMP and RFP was provided by a CoolLED pE-300 (CoolLED, UK). The TwinCam adapter was mounted with emission filters for green fluorescent protein (510/20 nm) and red fluorescent protein (590/35 nm), along with a DC/T510LPXRXTUf2 dichroic mirror. Imaging data were analyzed using Neuron Analyzer, a custom-written MATLAB (RRID:SCR_001622) program for processing image stacks (code available at https://github.com/neuronanalyser/neuronanalyser; neuronanalyser, 2015).

RNA extraction and sequencing

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To obtain RNA samples for RNA-seq, 30 young adult animals were picked to each fresh plate and allowed to lay eggs for 2 hr, after which the adult animals were removed and eggs were allowed to develop into day-1 adults. Day-1 animals were subsequently challenged with H2S for three different time periods (1 hr, 2 hr, and 12 hr). For each time period, five plates of day-1 animals were exposed to 50 ppm H2S, 150 ppm H2S, or 7% O2 as a control group. Subsequently, the animals were immediately collected and rinsed three times with M9 buffer. After washing, the worm pellet was frozen in liquid nitrogen. Animals were homogenized using Bullet Blender (Next Advance) in the presence of Qiazol Lysis Reagent and 0.5 mm Zirconia beads at 4 °C. RNA was prepared using RNeasy Plus Universal Mini Kit (QIAGEN). Six independent RNA samples were prepared for each condition. The 2100 Bioanalyzer instrument (Agilent) was used for the RNA quality control with a microfluidic chip specific for RNA (Agilent RNA 6000 Nano Kit). Then 1 μg of qualified RNA samples in RNase-free ddH2O was used for library preparation. The library was constructed and sequenced by Novogene.

RNA-seq analysis

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The RNA-seq data were aligned using STAR v2.7.9a (Dobin et al., 2013; RRID:SCR_004463) and gene expression counts extracted by featureCounts v2.0.3 (Liao et al., 2014; RRID:SCR_012919). Differential expression was calculated using DESeq2 (Love et al., 2014; RRID:SCR_000154). GO analysis was performed using the EnrichR R package (Kuleshov et al., 2016) on genes having p.adj<1e-10. For brevity, several highly similar GO categories were manually omitted (full EnrichR output available at GitHub repository).

Statistics details

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Stage-synchronized worms were randomly assigned into experimental and control groups for each genotype. Experiments were conducted with blinding to genotype and treatment. Sample sizes in behavioral assays are determined based on established methods and standardized procedures in C. elegans research. All data were included in the analyses. Locomotory speeds between genotypes were compared using in MATLAB (RRID:SCR_001622) scripts.

Materials availability

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All reagents generated in this study, including strains and plasmids, will be made available upon request.

Data availability

Raw sequencing data has been deposited at ArrayExpress (RRID:SCR_002964) # E-MTAB-13296. All data generated or analyzed during this study are included in the manuscript and supporting files. The R code is available at https://github.com/henriksson-lab/ce_h2s_tc (copy archived at Henriksson, 2025).

The following data sets were generated

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Article and author information

Author details

  1. Longjun Pu

    1. Department of Molecular Biology, Umeå University, Umeå, Sweden
    2. Umeå Centre for Molecular Medicine, Umeå University, Umeå, Sweden
    3. Wallenberg Centre for Molecular Medicine, Umeå University, Umeå, Sweden
    Contribution
    Data curation, Formal analysis, Validation, Investigation, Visualization, Methodology
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-0239-8732

  2. Lina Zhao

    1. Department of Molecular Biology, Umeå University, Umeå, Sweden
    2. Umeå Centre for Molecular Medicine, Umeå University, Umeå, Sweden
    3. Wallenberg Centre for Molecular Medicine, Umeå University, Umeå, Sweden
    Contribution
    Data curation, Formal analysis, Validation, Investigation, Visualization, Methodology
    Competing interests
    No competing interests declared

  3. Jing Wang

    1. Department of Molecular Biology, Umeå University, Umeå, Sweden
    2. Umeå Centre for Molecular Medicine, Umeå University, Umeå, Sweden
    3. Wallenberg Centre for Molecular Medicine, Umeå University, Umeå, Sweden
    Contribution
    Data curation, Formal analysis, Validation, Investigation, Visualization, Methodology
    Competing interests
    No competing interests declared

  4. Clementine Deleuze

    Laboratory of Neurophysiology, ULB Neuroscience Institute (UNI), Université Libre de Bruxelles (ULB), Brussels, Belgium
    Contribution
    Data curation, Formal analysis, Supervision, Validation, Investigation, Visualization, Methodology
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0009-0008-9581-9575

  5. Lars Nilsson

    1. Department of Molecular Biology, Umeå University, Umeå, Sweden
    2. Umeå Centre for Molecular Medicine, Umeå University, Umeå, Sweden
    3. Wallenberg Centre for Molecular Medicine, Umeå University, Umeå, Sweden
    Contribution
    Data curation, Formal analysis, Validation, Investigation, Visualization, Methodology
    Competing interests
    No competing interests declared

  6. Johan Henriksson

    1. Department of Molecular Biology, Umeå University, Umeå, Sweden
    2. The Laboratory for Molecular Infection Medicine Sweden (MIMS), Umeå University, Umeå, Sweden
    3. Integrated Science Lab (Icelab), Umeå University, Umeå, Sweden
    Contribution
    Data curation, Formal analysis, Validation, Visualization, Methodology
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-7745-2844

  7. Patrick Laurent

    Laboratory of Neurophysiology, ULB Neuroscience Institute (UNI), Université Libre de Bruxelles (ULB), Brussels, Belgium
    Contribution
    Conceptualization, Resources, Formal analysis, Supervision, Funding acquisition, Validation, Investigation, Visualization, Methodology, Writing – review and editing
    For correspondence
    patrick.laurent@ulb.be
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-5360-5597

  8. Changchun Chen

    1. Department of Molecular Biology, Umeå University, Umeå, Sweden
    2. Umeå Centre for Molecular Medicine, Umeå University, Umeå, Sweden
    3. Wallenberg Centre for Molecular Medicine, Umeå University, Umeå, Sweden
    Contribution
    Conceptualization, Resources, Data curation, Formal analysis, Supervision, Funding acquisition, Validation, Investigation, Visualization, Methodology, Writing – original draft, Project administration, Writing – review and editing
    For correspondence
    changchun.chen@umu.se
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-2233-8996

Funding

Vetenskapsrådet (2021-06602)

  • Johan Henriksson

Fonds De La Recherche Scientifique - FNRS (40020876)

  • Patrick Laurent

Vetenskapsrådet (2018-02216)

  • Changchun Chen

Vetenskapsrådet (2024-04141)

  • Changchun Chen

European Research Council (802653)

  • Changchun Chen

Fonds De La Recherche Scientifique - FNRS (40038114)

  • Clementine Deleuze

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

We thank the Caenorhabditis Genetics Center (funded by NIH Office of Research Infrastructure Programs P40 OD010440) and the National BioResources Project Japan for strains.

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You can cite all versions using the DOI https://doi.org/10.7554/eLife.92964. This DOI represents all versions, and will always resolve to the latest one.

Copyright

© 2023, Pu et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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  1. Longjun Pu
  2. Lina Zhao
  3. Jing Wang
  4. Clementine Deleuze
  5. Lars Nilsson
  6. Johan Henriksson
  7. Patrick Laurent
  8. Changchun Chen
(2025)
Avoidance of hydrogen sulfide is modulated by external and internal states in Caenorhabditis elegans
eLife 12:RP92964.
https://doi.org/10.7554/eLife.92964.4

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