DTT is known to be toxic to C. elegans and affects its development (Kozlowski et al., 2014). We first studied C. elegans development retardation at different concentrations of DTT. In the presence of Escherichia coli OP50 diet, C. elegans was at the L1 or L2 stage at 5 mM and higher DTT concentrations at 72 hr of hatching, while the control animals without DTT had developed to adulthood at the same time (Figure 1A, B). We also studied development retardation in the presence of DTT on E. coli HT115 bacterium, a strain commonly used for feeding-based RNA interference (RNAi). Surprisingly, we observed that C. elegans developed much better on DTT in the presence of E. coli HT115 diet than E. coli OP50 diet (Figure 1A–C). These results suggested that some bacterial components might be able to modulate DTT toxicity.

To gain an understanding of the potential bacterial components that might modulate DTT toxicity, we studied the development of C. elegans in the presence of DTT on different bacterial diets, including Comamonas aquatica DA1877, Pseudomonas aeruginosa PA14 gacA mutant, Salmonella enterica, and Serratia marcescens. Because P. aeruginosa virulence might affect the development of C. elegans, we used the gacA mutant, which has drastically reduced virulence (Tan et al., 1999). Interestingly, C. elegans showed no developmental defects on C. aquatica DA1877 and P. aeruginosa PA14 diets in the presence of 10 mM DTT (Figure 1D, E). Similarly, C. elegans developed much better on S. enterica and S. marcescens diets compared to the E. coli OP50 diet at lower DTT concentrations (Figure 1—figure supplement 1A–D). These results showed that bacterial diet has a major effect on DTT toxicity, and likely some bacterial diet component(s) modulated DTT toxicity.

Previous studies have shown that relative to the E. coli OP50 diet, a C. aquatica DA1877 diet accelerates C. elegans development primarily because it contains higher levels of vitamin B12 (Watson et al., 2014). Similarly, E. coli HT115 and P. aeruginosa PA14 diets are known to have higher amounts of vitamin B12 than the E. coli OP50 diet (Revtovich et al., 2019; Watson et al., 2014). The amount of dietary vitamin B12 available to C. elegans can be reliably determined by a dietary sensor strain expressing green fluorescent protein (GFP) under the promoter of acyl-coenzyme A dehydrogenase (acdh-1) gene (Watson et al., 2014). Low levels of vitamin B12 result in the accumulation of the short-chain fatty acid propionate, which leads to transcriptional activation of acdh-1 (Watson et al., 2014). Thus, the GFP levels in the acdh-1p::GFP animals indirectly report the levels of dietary vitamin B12 available to C. elegans. When the animals experience low levels of vitamin B12, acdh-1p::GFP is strongly induced, and the level of GFP expression reduces as the animals obtain increasing levels of vitamin B12. Using acdh-1p::GFP animals, we first tested the levels of vitamin B12 in the animals on different bacterial diets compared to the E. coli OP50 diet. As shown in Figure 1F and reported previously (Watson et al., 2014), the animals show very bright GFP expression on the E. coli OP50 diet, while the GFP levels were diminished on E. coli HT115, S. enterica, and S. marcescens diets. On the other hand, the animals showed no GFP expression on C. aquatica DA1877 and P. aeruginosa PA14 diets. The levels of GFP expression in acdh-1p::GFP animals on different bacterial diets correlated perfectly with the developmental defects on these bacterial diets in the presence of DTT (Figure 1G).

The correlation between the GFP levels in acdh-1p::GFP animals and the developmental defects of animals in the presence of DTT on different bacterial diets suggested that vitamin B12 could be the microbial component that alleviates DTT toxicity in C. elegans. To test this, we studied the development of N2 animals on E. coli OP50 diet containing 10 mM DTT and supplemented with 50 nM vitamin B12. In contrast to the development retardation on E. coli OP50 diet containing 10 mM DTT, the animals developed to adulthood on E. coli OP50 diet containing 10 mM DTT supplemented with 50 nM vitamin B12 (Figure 2A, B). We further asked whether the DTT-mediated development retardation could be reversed by supplementation of vitamin B12. To this end, vitamin B12 was added to the L1-arrested animals after 72 hr of hatching on E. coli OP50 plates containing 10 mM DTT. After 48 hr of incubation post vitamin B12 supplementation, more than 95% of the animals developed to adulthood (Figure 2—figure supplement 1A, B). On the other hand, the control animals on E. coli OP50 plates containing 10 mM DTT, but without vitamin B12 supplementation, were still at the L1–L2 stages after 120 hr of hatching. Taken together, these studies showed that vitamin B12 is sufficient to alleviate and revert the toxic effects of DTT.

Vitamin B12 is an essential cofactor for two metabolic enzymes: methylmalonyl-CoA mutase (MMCM-1 in C. elegans), which converts methylmalonyl-CoA to succinyl-CoA in the propionyl-CoA breakdown pathway, and methionine synthase (METR-1 in C. elegans), which converts homocysteine to methionine in the methionine–homocysteine cycle. We asked whether one or both of these enzymes are required for alleviating DTT toxicity. While vitamin B12 supplementation rescued the toxic effects of DTT in mmcm-1(ok1637) animals, it failed to do so in metr-1(ok521) animals (Figure 2C, D). The metr-1(ok521) animals were at the L1 stage on E. coli OP50 diet containing 10 mM DTT regardless of vitamin B12 supplementation (Figure 2C). Further, mmcm-1(ok1637) animals did not show any significant development retardation effects when grown on C. aquatica DA1877 and P. aeruginosa PA14 diets containing 10 mM DTT (Figure 2E, F). On the other hand, metr-1(ok521) animals failed to develop when grown on C. aquatica DA1877 and P. aeruginosa PA14 diets containing 10 mM DTT (Figure 2E, F). Taken together, these studies showed that vitamin B12 could alleviate DTT toxicity via the methionine synthase enzyme.

To understand why DTT toxicity depended on vitamin B12 and the methionine–homocysteine cycle, we carried out a forward genetic screen for mutants that would develop to L4/adult stage at 72 hr posthatching on E. coli OP50 diet containing 10 mM DTT. From a screen of approximately 50,000 ethyl methanesulfonate (EMS)-mutagenized haploid genomes, 12 mutants were isolated that exhibited DTT resistance and developed on E. coli OP50 diet containing 10 mM DTT (Figure 3—figure supplement 1A, B). To identify the causative mutations, we conducted the whole-genome sequencing of all the 12 mutants after backcrossing them six times with the wild-type N2 strain. The sequenced genomes were aligned with the reference genome of C. elegans. After subtraction of the common variants, linkage maps of single-nucleotide polymorphisms (SNPs) were obtained (Figure 3—figure supplement 2). All of the mutants had linkage to chromosome V. Analysis of the protein-coding genes carrying mutations in the linked regions of each mutant revealed that all of the mutants had a mutation in the gene R08E5.3 (Figure 3A, B), which was recently named as rips-1 (Rhy-1-Interacting Protein in Sulfide response) (personal communication with Wormbase). rips-1 is predicted to encode a protein with a SAM-dependent methyltransferase domain. SAM-dependent methyltransferases catalyze diverse methylation reactions where a methyl group is transferred to the substrate from SAM, resulting in the conversion of SAM to S-adenosylhomocysteine (SAH) in the methionine–homocysteine cycle. Fortuitously, the allele gk231506 available in the Caenorhabditis Genetics Center (CGC) has the same molecular change in the protein RIPS-1 as the allele jsn6 obtained in our screen has. rips-1(gk231506) animals showed DTT resistance and developed on E. coli OP50 diet containing 10 mM DTT (Figure 3C, D).

Three of the isolated alleles of rips-1, jsn4, jsn11, and jsn12 had premature stop codon mutations and one allele (jsn8) had a splice acceptor mutation (Figure 3A). All of these mutations are expected to result in a loss-of-function of the gene. To confirm that resistance to DTT toxicity is imparted by loss-of-function mutations in rips-1, we created rips-1(jsn11) animals expressing rips-1 under its own promoter. As shown in Figure 3E, the DTT-resistant phenotype of rips-1(jsn11) animals was completely rescued in these animals. We further determined the expression pattern of rips-1 by expressing GFP driven by its own 918 bp promoter in wild-type N2 animals. As shown in Figure 3F, rips-1 is expressed in the intestine.

We next asked how DTT exerted its toxic effects via rips-1. In the fungus Aspergillus niger, DTT is known to highly upregulate two SAM-dependent methyltransferases (MacKenzie et al., 2005). We asked whether DTT also upregulated the SAM-dependent methyltransferase rips-1, and any other closely related methyltransferases. rips-1 shares high sequence similarity with three other SAM-dependent methyltransferases in C. elegans, R08E5.1 (BLAST e value: 1e−168), R08F11.4 (BLAST e value: 1e−166), and K12D9.1 (BLAST e value: 2e−152). Quantitative reverse transcription-PCR (qRT-PCR) analysis showed that DTT resulted in significant upregulation of the transcript levels of rips-1, R08E5.1, and R08F11.4 (Figure 3G). The enhanced expression of rips-1 was confirmed in a reporter strain expressing GFP under the promoter of rips-1 (Figure 3H, I).

Because DTT resulted in the upregulation of methyltransferases closely related to rips-1, we used RNAi knockdown to study the role of these methyltransferases in the toxic effects of DTT. As expected, knockdown of rips-1 in the N2 animals resulted in a drastic improvement in their development in the presence of DTT (Figure 3—figure supplement 3A, B). Interestingly, knockdown of R08E5.1 resulted in a phenotype similar to rips-1 knockdown. On the other hand, knockdown of R08F11.4 and K12D9.1 improved the development in the presence of DTT only marginally (Figure 3—figure supplement 3A, B). These results suggest that the rips-1-related methyltransferases might also be required for DTT toxicity. It is also possible that the knockdown of these methyltransferases results in the knockdown of rips-1 by cross RNAi. The level of sequence similarity between rips-1 and R08E5.1 (78 % identical) indicates that these genes would undergo complete cross RNAi (Rual et al., 2007). Because we did not recover mutants of any of the rips-1-related methyltransferases in our genetic screen in contrast to the 12 alleles of rips-1, the rips-1-related methyltransferases likely have only minor roles in DTT toxicity. However, our studies do not rule out the involvement of the rips-1-related methyltransferases in DTT toxicity.

To establish that the toxic effects of DTT are indeed due to the upregulation of rips-1, we tested the effects of overexpression of rips-1 on DTT sensitivity. Animals overexpressing rips-1 exhibited enhanced sensitivity to DTT toxicity (Figure 3—figure supplement 3C, D). Taken together, these results suggested that by increasing the expression of rips-1, DTT modulates the methionine–homocysteine cycle resulting in toxic effects that can be reversed by supplementation of vitamin B12 (Figure 3J).

Modulation of the methionine–homocysteine cycle or low dietary vitamin B12 are known to upregulate the mitochondrial UPR (Amin et al., 2020; Wei and Ruvkun, 2020). Since DTT modulates the methionine–homocysteine cycle and requires vitamin B12 to reverse those effects, we asked whether DTT can also lead to the upregulation of the mitochondrial UPR. To this end, we exposed the adult hsp-6p::GFP animals to DTT. Compared to the control animals, animals exposed to 10 mM DTT had upregulation of the mitochondrial UPR (Figure 3—figure supplement 4A, B). Supplementation of vitamin B12 rescued the upregulation of mitochondrial UPR by DTT, indicating that DTT-induced mitochondrial UPR via the methionine–homocysteine cycle.

We tested whether the other thiol reagents, β-mercaptoethanol (β-ME) and NAC, caused toxicity via rips-1. Interestingly, rips-1(jsn11) animals exhibited resistance to β-ME toxicity, but not NAC toxicity (Figure 4A, B). We studied the effect of these thiol reagents on the expression levels of rips-1p::GFP. While NAC exposure did not affect the expression of rips-1p::GFP, β-ME resulted in its dramatic upregulation (Figure 4C, D). Finally, we studied the effects of vitamin B12 supplementation of the toxicities of β-ME and NAC. Vitamin B12 supplementation alleviated β-ME, but not NAC, toxicity (Figure 4E, F). These results suggested that β-ME shares the toxicity mechanism with DTT. On the other hand, NAC causes toxicity by a mechanism different from DTT.

Next, we asked how perturbations of the methionine–homocysteine cycle activity led to DTT toxicity. Since DTT exerts its toxicity via the SAM-dependent methyltransferase RIPS-1, we postulated that either depletion of the metabolites upstream of RIPS-1 activity (methionine and SAM) or accumulation of metabolites downstream of RIPS-1 activity (SAH and homocysteine) or a combination of both would result in DTT toxicity. To discriminate between these possibilities, we carried out methionine supplementation assays in the presence of DTT. We reasoned that if DTT toxicity is because of methionine and/or SAM depletion, supplementation of methionine should attenuate DTT toxicity. On the other hand, if DTT toxicity is caused by the accumulation of SAH and/or homocysteine, supplementation of methionine should enhance DTT toxicity as methionine supplementation would likely enhance levels of SAH and/or homocysteine. The wild-type N2 animals had improved development when 10 mM DTT containing E. coli OP50 diet was supplemented with varying concentrations of methionine (Figure 5A, B). Methionine supplementation alleviated DTT toxicity in a dose-dependent manner (Figure 5A, B). We observed that metr-1(ok521) animals that failed to develop upon vitamin B12 supplementation in the presence of DTT (Figure 2C, D), developed upon supplementation of methionine (Figure 5A, C). These results suggested that depletion of methionine and/or SAM is the primary cause of DTT toxicity.

Next, we studied whether DTT toxicity was an outcome of depletion of methionine per se or its downstream metabolite SAM. The first step in the methionine–homocysteine cycle involves the conversion of methionine into SAM by methionine adenosyltransferase (S-adenosylmethionine synthetase or SAMS in C. elegans). Animals having knockdown of sams-1 have defects in methionine to SAM conversion and exhibit significantly reduced levels of SAM (Walker et al., 2011). Because sams-1 mutants have reduced conversion of methionine to SAM, supplementation of methionine would alleviate DTT toxicity in sams-1 mutants only if DTT toxicity is caused by depletion of methionine and not SAM. Supplementation of methionine did not improve the development of sams-1(ok2946) animals on E. coli OP50 diet containing 10 mM DTT (Figure 5D, E). Further, sams-1(ok2946) animals also failed to develop on 10 mM DTT containing E. coli OP50 diet supplemented with 50 nM vitamin B12 (Figure 5D, E). Taken together, these results suggested that SAM depletion is the primary cause of DTT toxicity.

The genetic and methionine supplementation experiments suggested that SAM depletion could be the primary cause of DTT toxicity. To establish that DTT indeed resulted in the depletion of SAM, we quantified SAM levels in animals with and without exposure to DTT. The exposure of the wild-type N2 animals to DTT resulted in a significant reduction in their SAM levels (Figure 5F). To study whether SAM depletion by DTT depended on the gene rips-1, we quantified SAM levels in rips-1(jsn11) animals. rips-1(jsn11) animals did not have significant changes in SAM levels upon exposure to DTT (Figure 5F). Although nonsignificant, the SAM levels appeared to be higher in rips-1(jsn11) animals upon DTT exposure. It is unclear why DTT exposure resulted in the increased SAM levels in rips-1(jsn11) animals. Nevertheless, these results establish that DTT exposure depletes SAM levels in N2, but not rips-1(jsn11) animals.

SAM is the universal methyl donor in the cell and regulates a vast array of cellular activities, including the synthesis of phosphatidylcholine by methylation of phosphoethanolamine (Vance and Vance, 2004). Several of the SAM functions, including lipogenesis (Walker et al., 2011), gene expression changes (Giese et al., 2020), innate immunity (Ding et al., 2015; Nair et al., 2022), are regulated by phosphatidylcholine. Phosphatidylcholine can also be synthesized from choline by an alternate pathway independent of SAM (Vance and Vance, 2004). Importantly, the SAM-dependent phenotypes that require phosphatidylcholine can be rescued by supplementation of choline (Ding et al., 2015; Giese et al., 2020; Walker et al., 2011). We asked whether choline could also attenuate DTT toxicity. Supplementation of choline rescued the developmental defects in N2 and metr-1(ok521) animals on an E. coli OP50 diet containing 10 mM DTT (Figure 5G, H). The rescue of DTT toxicity was only partial in sams-1(ok2946) animals (Figure 5G, H). These results suggested that phosphatidylcholine is a major SAM product responsible for combating DTT toxicity. However, the partial rescue of DTT toxicity by choline supplementation in sams-1(ok2946) animals suggested that other SAM-related functions may also be involved in attenuating DTT toxicity.

Disturbance in the methionine–homocysteine cycle (accumulation of homocysteine and depletion of SAM) is known to result in ER stress (Hou et al., 2014; Walker et al., 2011; Werstuck et al., 2001). Since DTT causes ER stress, we asked what the contribution of methionine–homocysteine cycle perturbation in the DTT-induced ER stress was. To this end, we studied the effect of vitamin B12 and methionine supplementation on DTT-induced upregulation of hsp-4p::GFP. Supplementation of vitamin B12 and methionine suppressed DTT-induced upregulation of hsp-4p::GFP (Figure 6A, B). Choline supplementation is known to suppress the ER stress caused by the depletion of SAM (Hou et al., 2014). We observed that supplementation of choline resulted in suppression of DTT-induced upregulation of hsp-4p::GFP (Figure 6A, B). Thus, these results suggested that DTT causes ER stress in part by modulation of the methionine–homocysteine cycle.

Next, we explored the role of ER stress in DTT-mediated development retardation. First, we tested whether a lower concentration of DTT, which results in development retardation, would lead to ER stress. Exposure to 5 mM DTT resulted in the upregulation of hsp-4p::GFP to an intermediate level between the control and 10 mM DTT exposure levels (Figure 6—figure supplement 1A, B), indicating that lower concentrations of DTT also cause ER stress. The ER UPR pathways alleviate enhanced proteotoxic stress, and therefore, the animals having mutations in the UPR pathways have enhanced development retardation on chemical stressors such as tunicamycin (Shen et al., 2001). Surprisingly, we observed that in the presence of DTT, the mutants of different UPR pathways showed development retardation similar to the N2 animals (Figure 6C–G). Next, we tested whether vitamin B12 could alleviate DTT toxicity in the different UPR mutants. Similar to the wild-type N2 animals, supplementation of 50 nM vitamin B12 resulted in the rescue of both 5 and 10 mM DTT toxicity in atf-6(ok551) and pek-1(ok275) animals (Figure 6—figure supplement 1C–F). On the other hand, while supplementation of 50 nM vitamin B12 rescued the development of ire-1(v33) and xbp-1(tm2482) animals on 5 mM DTT, it failed to rescue their development on 10 mM DTT (Figure 6H–J and Figure 6—figure supplement 2). Taken together, these results suggested that at lower DTT concentrations, the primary cause of DTT toxicity was the modulation of the methionine–homocysteine cycle, and attenuation of DTT toxicity did not require a functional UPR. On the other hand, at higher DTT concentrations, the toxicity was an outcome of both the modulation of the methionine–homocysteine cycle and the ER proteotoxic stress, and attenuation of the DTT toxicity required a functional IRE-1/XBP-1 UPR pathway.

The following bacterial strains were used: E. coli OP50, E. coli HT115(DE3), P. aeruginosa PA14 gacA mutant, C. aquatica DA1877, S. marcescens Db11, and S. enterica serovar Typhimurium 1344. The cultures of these bacteria were grown in Luria–Bertani (LB) broth at 37°C. The nematode growth medium (NGM) plates were seeded with the different bacterial cultures and incubated at room temperature for at least 2 days before using for experiments.

C. elegans hermaphrodites were maintained at 20°C on NGM plates seeded with E. coli OP50 as the food source (Brenner, 1974) unless otherwise indicated. Bristol N2 was used as the wild-type control unless otherwise indicated. The following strains were used in the study: VL749 wwIs24 [acdh-1p::GFP + unc-119(+)], RB755 metr-1(ok521), RB1434 mmcm-1(ok1637), VC2428 sams-1(ok2946), RE666 ire-1(v33), xbp-1(tm2482), RB772 atf-6(ok551), RB545 pek-1(ok275), SJ4100 zcIs13 [hsp-6p::GFP + lin-15(+)], SJ4005 zcIs4 [hsp-4::GFP], and VC20288 rips-1(gk231506). Some of the strains were obtained from the Caenorhabditis Genetics Center (University of Minnesota, Minneapolis, MN). The xbp-1(tm2482) animals were obtained from National Bioresource Project, Japan.

The following supplements with their product numbers were obtained from HiMedia BioSciences: DTT (#RM525), β-mercaptoethanol (#MB041), NAC (#RM3142), vitamin B12 (cyanocobalamin) (#PCT0204), methionine (#PCT0315), and choline chloride (#TC102). All of the supplements that were added to the NGM plates were dissolved in water to prepare a stock solution. The supplements were added to the nematode growth media just before pouring into plates to obtain a desired final concentration.

Synchronized C. elegans eggs were obtained by transferring 15–20 gravid adult hermaphrodites on NGM plates for egg-laying for 2 hr. After 2 hr, the gravid adults were removed. The synchronized eggs were incubated at 20°C for 72 hr. After that, the animals in different development stages (L1/L2, L3, L4, and adult) were quantified. Since ire-1(v33) animals develop slightly slower than wild-type animals, the development stages in ire-1(v33) animals were quantified after 84 hr of incubation at 20°C. For choline and methionine supplementation assays, the synchronized eggs were incubated at 20°C for 4 and 5 days, respectively, before quantifying the development stages. 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.

RNAi was used to generate loss-of-function RNAi phenotypes by feeding nematodes E. coli strain HT115(DE3) expressing double-stranded RNA homologous to a target gene. E. coli with the appropriate vectors were grown in LB broth containing ampicillin (100 μg/ml) at 37°C overnight and plated onto NGM plates containing 100 μg/ml ampicillin and 3 mM isopropyl β-D-thiogalactoside (RNAi plates). The RNAi-expressing bacteria were allowed to grow overnight at 37°C on RNAi plates without DTT. On the other hand, the RNAi plates with 10 mM DTT were kept at room temperature for 2 days upon seeding with bacterial cultures. The wild-type N2 worms were first synchronized on RNAi plates without DTT and allowed to develop at 20°C for 3 days. The gravid adults obtained from RNAi plates without DTT were transferred to corresponding RNAi plates containing 10 mM DTT and allowed to lay eggs for 2 hr. The gravid adults were removed, and the eggs were allowed to develop at 20°C for 72 hr before quantifying the development stages. The RNAi clones were from the Ahringer RNAi library and were verified by sequencing.

To study whether the DTT-mediated development retardation could be reversed by supplementation of vitamin B12, 2 µl of 750 µM vitamin B12 (total 1.5 nmol) was added to the center of a plate with L1-arrested animals after 72 hr of hatching on E. coli OP50 with 10 mM DTT. To the control plate with L1-arrested animals after 72 hr of hatching on E. coli OP50 with 10 mM DTT, 2 µl of water was added to the center. After that, the plates were incubated at 20°C for 48 hr, followed by the quantification of development stages.

An EMS mutagenesis selection screen (Singh, 2021) was performed using the wild-type N2 strain. Approximately 2500 synchronized late L4 larvae were treated with 50 mM EMS for 4 hr 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 10 mM DTT and incubated at 20°C for 72 hr. After that, the plates were screened for animals that developed to L4 or adult stages. Approximately 50,000 haploid genomes were screened, and 12 mutants were isolated. All of the mutants were backcrossed six times with the parental N2 strain before analysis.

The genomic DNA was isolated as described earlier (Singh and Aballay, 2017). Briefly, the mutant animals were grown at 20°C on NGM plates seeded with E. coli OP50 until starvation. The animals were rinsed off the plates with M9, washed three times, incubated in M9 with rotation for 2 hr to eliminate food from the intestine, and washed three times again with M9. 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 whole-genome sequencing (WGS) on an Illumina HiSeq sequencing platform using 150 paired-end nucleotide reads. Library preparation and WGS were performed at Clevergene Biocorp Pvt. Ltd, Bengaluru, India.

The whole-genome sequence data were analyzed using the web platform Galaxy. 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 SNPs, insertions and deletions (indels), multinucleotide polymorphisms, 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 variation.

For rescue of the DTT-resistance phenotype of rips-1(jsn11) animals, the rips-1 gene along with its promoter region (918 bp upstream) was amplified from genomic DNA of N2 animals. The gene, including its stop codon, was cloned in the pPD95.77 plasmid containing the SL2 sequence before GFP. For generating the rips-1p::GFP reporter strain, the promoter region of the rips-1 gene (918 bp upstream) was amplified from genomic DNA of N2 animals and cloned in the pPD95.77 plasmid before GFP. rips-1(jsn11) animals were microinjected with rips-1p::rips-1::SL2::GFP plasmid along with pCFJ90 (myo-2p::mCherry) as a coinjection marker to generate rescue strains, rips-1(jsn11);jsnEx1[rips-1p::rips-1::SL2::GFP + myo-2p::mCherry] and rips-1(jsn11);jsnEx2[rips-1p::rips-1::SL2::GFP + myo-2p::mCherry]. To obtain rips-1 overexpressing strain, the rips-1(jsn11) allele was outcrossed to wild-type N2 animals to obtain jsnEx1[rips-1p::rips-1::SL2::GFP + myo-2p::mCherry] animals. N2 wild-type animals were microinjected with rips-1p::GFP plasmid along with pCFJ90 as a coinjection marker to generate rips-1p::GFP reporter strain. The rips-1p::rips-1::SL2::GFP and rips-1p::GFP plasmids were used at a concentration of 50 ng/µl, while the coinjection marker was used at a concentration of 5 ng/µl. The plasmids were maintained as extrachromosomal arrays, and at least two independent lines were maintained for each plasmid. The rips-1p::GFP array was integrated using UV irradiation to obtain jsnIs1[rips-1p::GFP + myo-2p::mCherry] followed by six times backcrossing with the wild-type N2 animals.

Animals were synchronized by egg laying. Approximately 35 N2 gravid adult animals were transferred to 9 cm E. coli OP50 plates without DTT and allowed to lay eggs for 4 hr. The gravid adults were then removed, and the eggs were allowed to develop at 20°C for 72 hr. 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 4 hr. 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 oligo dT primers using the Transcriptor High Fidelity cDNA Synthesis Kit (Roche) 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 and Aballay, 2019a). All samples were run in triplicate.

Animals were synchronized by egg laying. Approximately 40 N2 and rips-1(jsn11) gravid adult animals each were transferred to 9 cm E. coli OP50 plates without DTT and allowed to lay eggs for 4–5 hr. For every experiment, synchronization on six 9 cm E. coli OP50 plates (three plates for DTT treatment and control each) was carried out for each strain. The gravid adults were then removed, and the eggs were allowed to develop at 20°C for 72 hr. The animals were then collected, washed with M9 buffer, and transferred to 9 cm E. coli OP50 plates containing 10 mM DTT and incubated at 20 C for 12 hr. The control animals were maintained on 9 cm E. coli OP50 plates without DTT. After that, the animals were collected and washed with M9 buffer three to four times. The semisoft pellet of the animals was weighed (50–70 mg of worm pellet) and stored at −80°C until further processing. The samples were processed for metabolite extraction as described earlier (Nair et al., 2022) with appropriate modifications. Briefly, the worm pellets were thawed from −80°C and mixed with an ice-chilled acetonitrile–methanol–water (3:5:2) mixture containing 0.1% formic acid, followed by eight cycles of sonication with 3 s on and 30 s off at an amplitude of 40%. The suspension was centrifuged at 15,000 × g at 4°C for 10 min. Then, the supernatant was transferred to a fresh tube and was vacuum dried and stored at −80°C until processed for mass spectrometry.

For mass spectrometry, the sample was reconstituted in ice-chilled water with 0.1% formic acid. The volume of water used to reconstitute the samples depended on the weight of the preprocessed semisoft pellet of the animals, such that 1 µl of water was added per mg of worm pellet. The reconstituted mixture was centrifuged at 15,000 × g for 2 min, and 5 μl was injected for LC–MS analysis. The separation of metabolites was achieved on an Acquity UPLC BEH130 C18 column (2.1 × 100 mm, 1.7 µm particles, Waters) at room temperature. The two mobile phases consisted of (A) water with 0.1% formic acid and (B) acetonitrile with 0.1% formic acid. Linear gradients from 100% A to 90% A for 6 min were used for separation at a flow rate of 0.2 ml/min. The effluent was then introduced into the electrospray ionization source of the Synapt G2-Si HD mass spectrometer (Waters). The acquisitions were carried out in the positive ion mode. SAM chloride dihydrochloride was purchased from Sigma-Aldrich (#A7007) and was used as a standard. SAM levels were quantified by using the peak intensity at 399.1 m/z upon integration of SAM chromatogram peak.

The rips-1p::GFP animals were synchronized by egg laying on E. coli OP50 plates and incubated at 20°C for 72 hr. Subsequently, the animals were transferred to E. coli OP50 plates containing 10 mM DTT, 2 mM β-ME, and 20 mM NAC and incubated at 20°C for 8 hr. The control animals were maintained on E. coli OP50 plates without the chemical supplements. After that, the animals were prepared for fluorescence imaging.

The acdh-1p::GFP animals were synchronized by egg laying on different bacterial diets without DTT. The animals were incubated at 20°C for 72 hr. After that, the animals were prepared for fluorescence imaging.

The hsp-6p::GFP animals were synchronized by egg laying on E. coli OP50 plates without DTT and incubated at 20°C for 72 hr. Subsequently, the animals were transferred to E. coli OP50 plates containing 0, 10, and 10 mM DTT supplemented with 50 nM vitamin B12 followed by incubation at 20°C for 24 hr. After that, the animals were prepared for fluorescence imaging.

The hsp-4p::GFP animals were synchronized by egg laying on E. coli OP50 plates without DTT and incubated at 20°C for 72 hr. Subsequently, the animals were transferred to E. coli OP50 plates containing 0, 5, 10, 10 mM DTT supplemented with 50 nM vitamin B12, 10 mM DTT supplemented with 25 mM methionine, and 10 mM DTT supplemented with 80 mM choline chloride followed by incubation at 20°C for 24 hr. After that, the animals were prepared for fluorescence imaging.

Fluorescence imaging was carried out as described previously (Singh and Aballay, 2019b). Briefly, 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.

The statistical analysis was performed with Prism 8 (GraphPad). All error bars represent the standard deviation (SD). 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.