Strains used in this study are detailed in Table 1 . C. elegans was grown on nematode-growth media (NGM) plates seeded with E. coli OP50 at 20°C, according to standard procedures (30), unless mentioned otherwise.

C. elegans was grown at 20°C until L4 stage on NGM plates seeded with OP50 and then 100 μg/ml of FUDR was added. Animals were transferred to 25°C. Dead animals were counted every 24-48 hours. Animals that did not respond to touch with a pick were considered dead. 2-3 replicates with 20-30 animals on each plate were used for each experiment. Animals dying of disrupted vulva or crawling on walls were censored for analysis, which was performed using Prism 9 (Graphpad Inc.). Longevity assays were performed 3 independent times.

Animals were washed off growth or infection plates twice using sterile water and collected in 1.5 ml tubes containing 1,000 μl TRIzol (Sigma-Aldrich) for lysis. 100 μl chloroform was added and centrifuged at 12,000 rpm to separate the aqueous phase containing RNA. The aqueous phase was transferred to Purelink RNA mini kit (ThermoFisher Scientific) columns and purified total RNA was obtained following manufacturer’s instructions. cDNA was made using iScript cDNA synthesis kit (Bio-Rad). RT-qPCR was performed using SYBR Green Super mix (Bio-Rad) using a ViiA7 Real-Time qPCR system (Applied Biosystems). Test genes were normalized to neuronal reference gene snb-1. Fold change was calculated using the Pfaffl method (31).

Total RNA was collected from C. elegans using Purelink RNA kit as described above. Total RNA from 4 biological replicates for each condition was sent to BGI Genomics for library preparation and sequencing using BGI-seq 500. BGI provided clean reads in FASTQ format. FASTQ files were analyzed by DolphinNext high-throughput sequence analysis software (32). Steps followed in DolphinNext were as follows: FastQC was used to create quality control outputs. FASTQ files were aligned to the reference C. elegans genome using STAR. STAR was used to count or filter out common RNAs (eg. rRNA, miRNA, tRNA, piRNA). RSEM was used to align reads to reference transcripts and estimate gene and isoform expression levels. Genome-wide Bam analysis was done by RseQC. DEBrowser v1.22.5 was used for batch effect correction and normalization. Data analysis including generation of volcano plots and heat maps was done using DE analysis within DEBrowser v1.22.5. Adjusted p-value (P_adj) ≤ 0.01 was considered significant for differential gene expression. Gene Ontology representation analysis was performed using the enrichment analysis tool in Wormbase (33) and g:Profiler (34).

To generate Pnhr-42::mCherry, the nhr-42 promoter region (2 kbp upstream of the transcription start site) was amplified and cloned into entry vector pTOPO for Gateway^® recombination (ThermoFisher Sci.). This amplified region was recombined into destination vector pDest-16 (generous gift by Dr. Michael Francis, UMass Chan Medical School) expressing mCherry to generate the expression vector. The expression vector was sequenced to verify proper insertion of the nhr-42 promoter upstream of the mCherry coding sequence. Transformation was carried out by gonad microinjection following standard procedures (35).

Images were taken using a Lionheart FX automatic microscope (BioTek Instruments) at 4x and 20x magnification. 30-50 C. elegans were used for each experimental condition. 10-20 animals at a time were anesthetized using 100 mM NaN₃ on a 2% agar pad for imaging. All images were captured at the same exposure and intensity. Greyscale images were used for image analysis. A region of interest (ROI) was drawn around the pharynx, which showed fluorescent expression of mCherry. Mean fluorescence intensity (MFI) of the ROI was determined by using the Analyze>Measure function in ImageJ (NIH). MFI values were plotted in Prism 9 for statistical analysis. Fluorescence microscopy experiments were repeated 2 independent times.

C. elegans was grown at 20°C until L4 stage on NGM plates seeded with OP50. 20-30 animals were placed on TSA plates containing a full lawn of S. aureus or BHI plates for E. faecalis infection. 8 h after infection, animals were washed off with water and collected in centrifuge tubes. Lipid staining using Oil Red O was performed as described (36). 10-20 animals were used for imaging. Animals were mounted on agarose pads and visualized using a Lionheart microscope at 4x and 20x magnification and the staining intensity was determined using ImageJ. Greyscale.tiff images were used for quantification. A relative threshold for the detection of red color staining was set manually (Image>Threshold>). The set threshold was constant for all images quantified. A region of interest (ROI) was drawn around each whole animal. The percent area stained inside the ROI by Oil Red O for each whole animal was calculated (Analyze>Measure>Area%). For intestine-specific quantification, the intestine (visualized by brightfield microscopy) was used as the ROI. Percent of area (Area%) stained by Oil Red O per animal was plotted and statistical analysis were performed in Prism.

C. elegans was grown at 20°C until L4 stage on NGM plates seeded with OP50 or RNAi clones as mentioned. Thereafter, 30-40 animals were transferred to TSA plates containing a full lawn of S. aureus. At the desired timepoint 10 live animals were collected in centrifuge tubes containing autoclaved water. Colony forming units were determined as in (8). 5-7 biological replicates (10 animals per replicate) were used for each condition.

GraphPad Prism 9 was used for statistical analysis. Survival curves were compared using Log-Rank (Mantel-Cox test). Unpaired, two-sample two-tailed t-tests were performed to compare with a single reference. For multiple comparisons, one-way ANOVA was used to establish significance followed by Šidák’s post-hoc test. P ≤ 0.05 was considered significant.

We previously showed that hlh-30 induces around 600 genes during S. aureus infection (8). However, HLH-30 is predicted to induce just 10% of these genes by directly binding their promoters ( Figure 1A ) (8). We hypothesized that indirect regulation of key host defense genes by hlh-30 could be mediated by the 17 transcription factors it induces during infection (8). To identify which are important in the intestinal epithelium for host defense, we performed intestine-specific silencing by RNAi. As expected, we found genes whose knockdown impaired host survival (e.g., nhr-55, Figure S1A ) suggesting that they may be important for inducing the intestinal host response to infection. However, we were surprised to identify one gene (nhr-42), whose knockdown significantly promoted host survival ( Figure S1D ), suggesting that nhr-42 may function as a host response repressor.

Public datasets contain evidence that HLH-30 may bind directly to the nhr-42 promoter ( Figure 1B ) (37–39), suggesting that nhr-42 may be a direct target of HLH-30. A fluorescent nhr-42 transcriptional reporter showed expression mainly in the pharynx of noninfected animals ( Figure 1C ), in agreement with a previous report (40). The same reporter exhibited increased activity in the pharynx after S. aureus infection ( Figures 1D, E, I ), suggesting that nhr-42 expression may be induced transcriptionally. In contrast, silencing of hlh-30 abrogated induced reporter activity after infection ( Figures 1F–I ). These results suggested that the nhr-42 promoter is active in the pharynx, and that its activity increases during infection, in the same tissue, in an HLH-30-dependent manner. This result is consistent with our prior RNA-seq data (8).

To gain better understanding of the timing of nhr-42 induction, we performed time-resolved infection followed by RT-qPCR. This revealed that nhr-42 expression is relatively stable in noninfected animals, and independent of HLH-30 ( Figure 1J ). During infection, nhr-42 expression was elevated throughout the experiment and peaked around 11 h in wild type animals, but in hlh-30 (–) mutants nhr-42 expression remained low ( Figure 1K ). These data showed that in noninfected animals nhr-42 expression does not require HLH-30, but that its early induction and peak about 11 h during infection are HLH-30-dependent.

Although we observed nhr-42 promoter activity mainly in the pharynx, nhr-42 could function in other tissues to repress host defense. To systematically identify the tissues in which nhr-42 functions, we performed tissue-specific knockdown of nhr-42 by RNAi ( Figure 2 ). nhr-42 knockdown in the whole animal strongly enhanced host survival ( Figure 2A ), providing a benchmark. As previously, intestinal knockdown enhanced survival of infection ( Figure 2B ), as also did epidermal (hypodermal) silencing ( Figure 2C ). In contrast, muscle knockdown showed no significant effect ( Figure 2D ). These data suggest that nhr-42 represses host survival of infection mainly from the intestinal epithelium, with contributions from the epidermis.

Consistently, a total knockout of nhr-42 provided strong protection against S. aureus ( Figure 2E ) and Enterococcus faecalis, a distantly related Gram-positive human pathogen ( Figure 2F ). However, loss of nhr-42 did not affect defense against Pseudomonas aeruginosa, a Gram-negative human pathogen ( Figure S2 ). These results show remarkable specificity of nhr-42 function.

Because nhr-42 induction by S. aureus is hlh-30-dependent, we hypothesized that they might function in the same genetic pathway. hlh-30;nhr-42 double mutants exhibited the same susceptibility to S. aureus as hlh-30 single mutants, completely suppressing the enhanced survival phenotype of nhr-42 mutants ( Figure 2G ). Remarkably, nhr-42 loss did not affect lifespan on nonpathogenic E. coli, and again hlh-30 was epistatic to nhr-42 ( Figure 2H ). In a linear genetic pathway model, these results would imply that hlh-30 functions downstream or parallel to nhr-42; however, hlh-30 expression was unaffected in nhr-42 mutants (see below). Given the results so far, we rather favored a model in which hlh-30 functions upstream of nhr-42 to induce host defense genes that nhr-42 represses.

To gain further insight into the function of nhr-42, we performed whole-animal RNA-seq of noninfected and infected animals ( Figure 3A ). We compared the transcriptomes of mutants to wild type under both conditions to identify differentially expressed genes ( Figures 3 , S3 ; Table S1 ).

292 transcripts were expressed more highly in noninfected nhr-42 mutants compared to wild type ( Figures 3B ; Table S1 ). We found only minor overlap between this set of genes overexpressed in nhr-42 mutants and HLH-30-induced genes in infected animals (8)( Figure 3D ). Nonetheless, noninfected nhr-42 mutants showed patterns that were easily identified by gene ontology (GO) over-representation analysis ( Table S2 ). Specifically, relative to wild type, nhr-42 mutants showed increased expression of genes related to various innate immunity and host defense categories (including Defense Response to Fungus) and decreased expression of genes related to the response to unfolded protein ( Figure 3E ; Table S2 ). This remarkable enrichment for host defense genes reveals that NHR-42 functions to repress host defense in noninfected animals, which is consistent with the observed enhanced survival of infection that is caused by nhr-42 loss. Together, these results suggest that, in noninfected animals, NHR-42 represses host defense genes that are mostly HLH-30-independent.

During infection, a more complex picture emerged. nhr-42 mutants showed increased expression of 525 transcripts ( Figure 3C ; Table S1 ), enriched for genes related to reproduction (including cell cycle and embryo development) and DNA damage response, of unclear relevance to infection biology ( Table S2 ; Figure 3F ). Perhaps easier to rationalize, they also expressed higher levels of genes related to signal transduction, response to stress, defense response (including innate immune response), apoptosis, and autophagy (including macroautophagy, negative regulation of autophagy, and TOR signaling), which are all processes that have been linked to host defense in C. elegans by prior research.

In contrast, infected nhr-42 mutants showed decreased expression of 218 transcripts relative to wild type ( Table S2 ). The most outstanding over-represented categories were related to lipid metabolism, including fatty acid biosynthesis and lipid catabolism ( Figure 3G ). This result prompted us to investigate the relationships among infection, lipid stores, and nhr-42.

The RNA-seq analysis showed that nhr-42 mutants expressed lower levels of seven known or proposed lipid catabolism genes ( Figure 4A ). These include predicted enoyl-CoA hydratase ech-7 and acyl-CoA oxidases acox1.3 and acox1.4, which are potentially involved in fatty acid β-oxidation. Also downregulated in nhr-42 mutants was Abhydrolase domain-containing homolog abhd-3.2, which is predicted to be a phospholipase involved in catabolism of medium chain fatty acids. However, genes involved in lipid biosynthesis were also repressed in infected nhr-42 mutants ( Table S2 ). To determine the biological significance of these observations, we assessed the lipid status of nhr-42 mutants by neutral lipid droplet staining with Oil Red O (ORO) (41, 42).

Noninfected nhr-42 mutants showed a similar level of ORO staining as wild type ( Figure 4B–I ), indicating that nhr-42 does not affect the overall levels of lipid droplets at baseline. After infection, wild type animals showed strongly decreased ORO staining ( Figures 4B, C, J–L ), whereas in nhr-42 mutants staining was significantly preserved ( Figures 4B, C, M–O ). This indicated that nhr-42 is required for full lipid loss during infection with S. aureus. Furthermore, we observed a similar defect during infection with E. faecalis ( Figure S4 ), showing that nhr-42 is partially required for lipid loss during infection by two archetypal Gram-positive pathogens.

We and others showed that hlh-30 is required for lipid loss during starvation (5, 6) and for host defense against S. aureus infection (8), suggesting that the two functions of HLH-30 may be related. Indeed, it was proposed that HLH-30 induces the expression of lipid catabolism genes that may be responsible for lipid droplet loss, at least during starvation (5). Recall that hlh-30 was epistatic to nhr-42 for infection survival ( Figure 2G ) i.e., hlh-30;nhr-42 double mutants are defective in infection survival, like the hlh-30 single mutants. To determine the relationship between this phenotype and lipid loss, we performed ORO staining of the hlh-30;nhr-42 double mutants ( Figure 5 ). hlh-30;nhr-42 double mutants were not significantly different from wild type, hlh-30, or nhr-42 animals at baseline ( Figures 5A–F, M ). After infection with S. aureus, the hlh-30 single mutants showed a much smaller drop in ORO staining compared to wild type, to a level similar to the nhr-42 single mutants ( Figures 5G–I, M ). Moreover, the hlh-30;nhr-42 double mutants were statistically indistinguishable from the single mutants ( Figures 5J–M ). Therefore, hlh-30 is also required for lipid loss during S. aureus infection, and by epistasis participates in the same genetic pathway as nhr-42. However, loss of hlh-30 impairs host infection survival while loss of nhr-42 promotes it; therefore, these mutations have opposite effects on infection survival but the same effect on lipid loss. These data support the notion that nhr-42 mediates the lipid loss function of hlh-30, and thus that HLH-30 does not directly drive lipid loss during infection. In further support of this notion, of the lipase-like (lipl) genes that were proposed to function downstream of HLH-30, only lipl-2 was differentially expressed by RNA-seq, and only in one condition: infected nhr-42 mutants expressed more lipl-2 compared with infected wild type ( Table S1 ). Therefore, the defective lipid loss of nhr-42 mutants is likely not due to decreased lipl gene expression. It follows that infection survival and lipid loss have distinct genetic requirements, and that lipid droplet loss does not explain the enhanced resistance to infection of nhr-42 mutants.

As mentioned, noninfected nhr-42 mutants displayed a strong signature of upregulated host defense genes ( Figure 3E ). Such upregulated genes include antibacterial factor-related genes abf-1 and abf-2, antimicrobial caenacins cnc-2, cnc-4, and cnc-6, and immune response gene irg-5, as well as lec-11 and pals-23, which were previously implicated in host defense ( Figure 6A ) (43–50). RT-qPCR independently confirmed their upregulation, except for irg-5 ( Figure 6B ). To test the hypothesis that antimicrobial gene upregulation may be a major contributor to the enhanced infection survival of nhr-42 mutants, we performed RNAi-mediated knockdown. Silencing of cnc-4, irg-5, and pals-23 had no significant effect in nhr-42 mutants, whereas silencing of abf-2, cnc-2, and lec-11 significantly decreased their enhanced survival ( Figures 6C–H ). abf-2 and cnc-2 knockdowns showed the strongest effects in nhr-42 mutants, but did not affect the wild type ( Figures 6C, D ). Curiously, silencing of lec-11 (which reduced the survival of nhr-42) and of pals-23 (which did not), both enhanced the survival of wild type ( Figures 6G, H ). Thus, specific antimicrobial effectors that are constitutively induced in nhr-42 mutants are required for their enhanced infection survival.

To determine if nhr-42 mutants show enhanced infection survival because of enhanced infection resilience (i.e., same colonization as wild type) or resistance (i.e., lower colonization than wild type), we measured S. aureus accumulation in the C. elegans gut over time. Whereas by 24 h of infection the nhr-42 mutants accumulated significantly less S. aureus than wild type ( Figure 7A ), they tended to accumulate less at earlier time points as well. Control experiments showed that the pharynxes of nhr-42 mutants pumped at rates that are indistinguishable from wild type ( Figure 7B ), suggesting that the decreased accumulation of S. aureus in nhr-42 mutants was not due to decreased oral intake. Of the genes that were required for enhanced survival of nhr-42 mutants, abf-2 is the only one whose product has demonstrated antibacterial activity in vitro (44), and therefore we chose it for further study. Animals treated with abf-2 RNAi showed increased S. aureus load, comparable to wild type ( Figure 7C ), suggesting that abf-2 mediates infection resistance in nhr-42 mutants. As mentioned, noninfected nhr-42 mutants exhibit higher expression of abf-2 than wild type counterparts ( Figures 6A, B ). However, abf-2 expression at 5 h infection was similar in wild type and nhr-42 mutant animals ( Figure 7D ), suggesting that abf-2 may be overexpressed only in noninfected nhr-42 mutants. However, these data do not rule out potentially increased abf-2 expression in nhr-42 mutants at other times of infection.