To study the role of DBL-1 in specific host responses to bacteria at behavioral, molecular, and physiological levels, we first established a panel of bacteria that would facilitate genetic and molecular studies over time: previous studies have been limited in range of challenge and used pathogens that killed animals in hours or a few days. The control bacteria chosen was Gram-negative Escherichia coli OP50, a commonly used strain for laboratory culture of C. elegans. The panel of test bacteria comprised Gram-negative and Gram-positive bacteria that are opportunistic pathogens in humans and are found in the natural habitat of C. elegans (Samuel et al., 2016). We selected three Gram-negative test strains, Serratia marcescens (D1), Enterobacter cloacae (ATCC 49141), and Klebsiella oxytoca (ATCC 49131), and three Gram-positive test strains, Bacillus megaterium (ATCC 14581), Enterococcus faecalis (ATCC 51299), and Staphylococcus epidermidis (ATCC 49134).

We first asked if these bacteria alter lifespan of wild-type C. elegans. While pathogenic bacteria can reduce C. elegans lifespan, other microbial diets can extend lifespan of C. elegans populations (Stuhr and Curran, 2020). We noted an extended lifespan of wild-type animals on two Gram-negative bacteria (E. cloacae and K. oxytoca, by 4 d and 2 d, respectively). However, we found that wild-type animals on Gram-negative S. marcescens have lifespans comparable to E. coli-fed animals. Interestingly, we observed an extended lifespan of wild-type animals on all three Gram-positive bacteria (B. megaterium, E. faecalis, and S. epidermidis) (Supplementary file 1).

Loss of DBL-1 has previously been shown to reduce lifespan of animals exposed to fungus Drechmeria coniospora, Gram-negative strains E. coli and S. marcescens Db11, and Gram-positive E. faecalis (Mallo et al., 2002; Zugasti and Ewbank, 2009; Tenor and Aballay, 2008). The reduced lifespan of animals lacking DBL-1 on E. coli OP50 was not observed when animals were grown on 5-fluorodeoxyuridine (FUdR), which blocks DNA synthesis of both bacteria and the nematodes (Mallo et al., 2002). It has also been shown that DBL-1 is required for the probiotic Lactobacillus spp. Lb21-mediated lifespan extension of C. elegans subjected to pathogenic methicillin-resistant Staphylococcus aureus (MRSA) (Mørch et al., 2021). To determine if DBL-1 is required in maintaining lifespan of animals subjected to our bacterial panel, we compared lifespans of dbl-1(-) and wild-type animals exposed to the control or test bacteria on plates containing FUdR. Loss of DBL-1 did not alter lifespan of animals fed on plates containing E. coli OP50 and FUdR (Figure 1A), consistent with previous work (Mallo et al., 2002). dbl-1 mutant animals did not have the lifespan extension seen in wild-type populations on E. cloacae or K. oxytoca: lifespans of dbl-1 mutant populations were the same on these two Gram-negative bacteria as on E. coli (Figure 1B and C, Supplementary file 1). However, loss of DBL-1 resulted in a significantly shortened lifespan on S. marcescens compared to E. coli control bacteria, unlike the wild-type (by 6 d, Supplementary file 1). This reduced lifespan of dbl-1(-) animals was significantly lower than that of wild-type animals fed on S. marcescens, which is consistent with a previous report that used the more virulent S. marcescens strain Db11 (Figure 1D; Mallo et al., 2002). The lifespan of dbl-1(-) animals was modestly extended upon exposure to Gram-positive B. megaterium and E. faecalis compared to the E. coli-fed population’s lifespan, but was significantly less extended than the wild-type lifespan (by about a day, Figure 1E and F, Supplementary file 1). Lastly, dbl-1 mutant animals displayed a significantly decreased lifespan on S. epidermidis compared to E. coli-fed populations’ lifespan (by almost 2 d), which was not observed in wild-type animals fed on S. epidermidis (Figure 1G, Supplementary file 1).

In summary, we showed that exposure to all bacteria in the test panel except S. marcescens resulted in lifespan extension of wild-type animals. Loss of DBL-1 reduced lifespan in all test bacteria in this panel to varying degrees. dbl-1(-) mutant animals have the lowest average lifespan on S. marcescens, significantly lower than the same strain on control E. coli OP50. In contrast to the wild type, dbl-1(-) animals fed S. epidermidis have reduced lifespans compared to the same strain on control E. coli OP50. Wild-type lifespan extension is lost in dbl-1(-) mutant animals fed E. cloacae and K. oxytoca, but their average lifespan is comparable to the same strain grown on E. coli OP50. On B. megaterium and E. faecalis, the lifespan of dbl-1(-) mutants was extended but to a lesser extent than the wild type (Figure 1B–G). Together, these results provide evidence that DBL-1 signaling confers protection against some bacteria and contributes to the lifespan extension observed in response to most bacteria in our panel.

One cause of reduced survival upon exposure to pathogenic bacteria is disruption of intestinal integrity (Sim and Hibberd, 2016). To examine if this is the underlying reason for DBL-1-dependent differences in animal survival in response to various bacteria, we investigated the integrity of the intestinal barrier of animals fed on different bacteria over time. We compared the intestinal barrier function of wild-type and dbl-1(-) animals exposed to control and test bacteria when 50% of the population on the test bacteria was still alive using a cell-impermeable blue dye that remains in the intestinal lumen in healthy animals (Gelino et al., 2016). Animals with intact intestinal barrier function will contain dye only within the intestinal lumen, but animals with intestinal barrier dysfunction will have leakage of the dye from the intestine into the body cavity. Consistent with previous reports, wild-type populations on E. coli exhibited an increased percentage of animals with dye outside the intestine over time, indicating an age-dependent reduction of intestinal integrity (Gelino et al., 2016). dbl-1-mutant populations grown on E. coli also displayed a decline in intestinal integrity that was not significantly different from the decline observed in the wild-type populations at the tested time points (Figure 1—figure supplement 1A). Exposure of wild-type animals to any of the test bacteria in the panel did not further decrease intestinal integrity compared to E. coli (Figure 1—figure supplement 1). Similar to wild-type animals, exposure of dbl-1(-) mutants to the test bacteria did not further significantly decrease their intestinal integrity. Therefore, DBL-1 is not required for intestinal integrity or its age-related decline. Furthermore, this result suggests the lifespan changes observed in dbl-1 mutant populations are not caused by loss of intestinal integrity.

Lifespan can also be reduced by increased bacterial colonization in the C. elegans gut (Portal-Celhay et al., 2012). To determine if the decrease in lifespan of dbl-1(-) animals is correlated with an increase in gut colonization, we quantified bacterial load in the intestine of wild-type and dbl-1(-) animals fed S. marcescens, the bacterial strain in the selected panel that had the most deleterious lifespan effect on dbl-1(-) populations. No significant differences in bacterial load were observed between wild-type and dbl-1(-) animals that were exposed to S. marcescens (Figure 1—figure supplement 2). This suggests that bacterial colonization is not the underlying cause for reduction in dbl-1(-) populations’ survival upon exposure to S. marcescens.

We next asked if our panel of bacteria affect feeding behavior in C. elegans. C. elegans reduce their food intake in response to bacteria they perceive as harmful (O’Quinn et al., 2001). To determine if loss of DBL-1 alters animals’ perception of test bacteria as potential threat, we assessed pharyngeal pumping of wild-type and dbl-1(-) populations fed on control and test bacteria. Loss of DBL-1 does not alter the pharyngeal pumping rate of animals fed on the control E. coli OP50 bacteria (Figure 2A and B). Overall, animals fed the Gram-negative bacteria did not have major changes in pharyngeal pumping rates (Figure 2A). Wild-type animals fed on E. cloacae exhibited a small but significant decrease in pharyngeal pumping that was not further decreased in dbl-1(-) animals. Pharyngeal pumping of wild-type animals was not significantly reduced after exposure to K. oxytoca, but animals lacking DBL-1 had a mild but significant decrease in pumping rate. Wild-type and dbl-1(-) animals fed on S. marcescens had a similar pumping rate as on the control bacteria (Figure 2A). There is no clear correlation between lifespan phenotypes and feeding rate for populations fed on the Gram-negative bacteria. For wild-type animals fed on the three Gram-positive bacteria, though, the pharyngeal pumping rate was dramatically decreased, consistent with the lifespan extension these strains conferred to wild-type C. elegans (Figure 2B, Supplementary file 1). Interestingly, the feeding rate of dbl-1(-) animals is further reduced from the wild-type rate on E. faecalis and S. epidermidis. However, there is no reproducibly significant decrease in the pharyngeal pumping rate of dbl-1(-) animals fed on B. megaterium in comparison to that of the wild type (Figure 2B). These results indicate that while the feeding reduction caused by exposure to the Gram-positive bacteria is partially independent of DBL-1, a stronger pharyngeal pumping reduction in dbl-1(-) populations occurs in response to some bacteria, providing support to the idea that loss of DBL-1 sensitizes animals to certain bacterial stressors.

C. elegans can also respond to potentially pathogenic bacteria or harmful environments by avoiding such challenges (Pradel et al., 2007; Beale et al., 2006; Anderson and McMullan, 2018). To further investigate the role of DBL-1 in perceiving harmful environments, we performed an avoidance assay. First, we measured the avoidance response of wild-type animals fed on the test bacteria over the first 2 d of adulthood and compared it with the avoidance response on the control bacteria. Wild-type animals do not avoid E. coli (Figure 3A). We found that wild-type animals do not avoid E. cloacae but do avoid Gram-negative K. oxytoca and S. marcescens (Figure 3B–D). While wild-type animals displayed a moderate to strong response to Gram-positive B. megaterium, they exhibited mild or no avoidance of E. faecalis and S. epidermidis (Figure 3E–G). This indicates that wild-type animals perceive K. oxytoca, S. marcescens, and B. megaterium as harmful.

Next, we tested avoidance behavior of dbl-1(-) populations exposed to the test bacteria and compared it with wild-type animals fed on the same test bacteria. Loss of DBL-1 did not result in an avoidance response to the control bacteria (Figure 3A). Loss of DBL-1 resulted in a significantly stronger avoidance response to all three tested Gram-negative bacteria (Figure 3B–D). Interestingly, the avoidance response of dbl-1(-) animals upon exposure to Gram-positive B. megaterium, E. faecalis, and S. epidermidis was similar to wild type (Figure 3E–G). The protective behavioral response displayed by dbl-1(-) populations on bacteria that wild-type animals do not avoid suggests that animals lacking DBL-1 perceive the tested Gram-negative (but not Gram-positive bacteria) to be more harmful than wild-type animals do. This also suggests that DBL-1 signaling normally suppresses avoidance depending on the specific bacterial exposure.

Because we observed strong bacteria-specific avoidance responses in animals lacking DBL-1, we next asked if the canonical DBL-1 signaling pathway is required to attenuate this response. Canonical signaling occurs by DBL-1 ligand binding to receptors SMA-6 and DAF-4, which activate downstream Smad transcription factors SMA-2, SMA-3, and SMA-4 (Savage et al., 1996). A non-canonical DBL-1 pathway, which does not signal through SMA-2 and SMA-4, is required for C. elegans to respond to the fungus D. coniospora (Zugasti and Ewbank, 2009). To identify the Smads required for DBL-1 pathway-mediated bacterial avoidance, we measured the avoidance response of sma-2(-), sma-3(-), and sma-4(-) strains fed on the test Gram-negative or Gram-positive bacteria and compared them with the avoidance response on the control bacteria. On the control bacteria, loss of sma-2 and sma-3 did not result in significantly increased avoidance. However, populations with sma-4(e727) showed a moderate but significant avoidance response to the control bacteria (Figure 3A). Because sma-4(e727), a strong loss-of-function point mutation (Q246Stop), initially displayed a stronger avoidance response, we confirmed this effect was reproducible in the sma-4(jj278) null strain, in which most of the open reading frame was deleted using CRISPR (McKillop et al., 2018). sma-2(-), sma-3(-), and both sma-4(-) populations displayed strong, reproducible avoidance responses to all Gram-negative bacterial strains tested that were comparable to the dbl-1(-) populations’ response (Figure 3B–D).

In comparison, the response of Smad mutant populations to the panel of Gram-positive bacteria was notably different. Loss of sma-2, sma-3, or sma-4 resulted in avoidance responses on the Gram-positive B. megaterium and E. faecalis similar to the wild type and dbl-1(-) responses (Figure 3E and F). This indicates that avoidance elicited in response to these bacterial strains is independent of not only DBL-1, but also of the Smad machinery. However, both sma-4 mutant strains but not dbl-1, sma-2, or sma-3 strains had a stronger, moderate avoidance response on the Gram-positive S. epidermidis in comparison to the wild type (Figure 3G). Together, our findings indicate that while canonical DBL-1 signaling does not play a major role in responding to E. coli, a standard Gram-negative food source, and Gram-positive bacteria, it plays a major role in suppressing animal avoidance responses to the tested Gram-negative bacteria.

We next asked if animals alter expression of the DBL-1 Smads in response to the panel of opportunistic pathogens. We tested gene expression levels of sma-2, sma-3, and sma-4 in wild-type and dbl-1 mutant backgrounds in response to our panel of bacteria. Gene expression was assessed by real-time PCR experiments that were performed in triplicate in three independent trials. Expression of mRNA from these three Smad genes was not changed in dbl-1(-) animals on control E. coli OP50 compared to the wild type (Figure 4—figure supplement 1A). The relative levels of sma-2 mRNA were consistently decreased in the dbl-1 mutant background upon exposure to all test bacterial strains except S. epidermidis, where sma-2 mRNA levels were increased in both the wild-type and dbl-1(-) backgrounds (Figure 4A and B and Figure 4—figure supplement 1A and B). The relative levels of sma-3 mRNA were not reproducibly different between dbl-1(-) and wild-type backgrounds exposed to the same control or test bacteria (Figure 4—figure supplement 1C and D). Whereas sma-3 expression levels remained largely unchanged in wild-type animals when exposed to Gram-negative and Gram-positive bacteria, exposure to E. faecalis or S. epidermidis resulted in increased expression of sma-3 in the dbl-1(-) background compared to control bacteria (Figure 4C and D). On test Gram-negative bacteria, sma-4 expression was similar in the dbl-1(-) background as in the wild type (Figure 4E, Figure 4—figure supplement 1E). However, sma-4 was significantly induced in response to Gram-positive bacteria in both wild-type and dbl-1 mutant backgrounds, albeit less in the dbl-1(-) populations (Figure 4F and Figure 4—figure supplement 1F). Together, these results suggest that the DBL-1 Smads are differently regulated at the level of gene expression by molecular pathways that are responsive to specific bacterial challenges. sma-2, but not sma-3, requires DBL-1 for full expression on all bacterial food sources except S. epidermidis. Neither sma-2 nor sma-3 expression is affected to a great extent by test bacteria. In contrast, sma-4 expression is not altered by Gram-negative bacteria, but is responsive to Gram-positive bacteria in the panel, possibly by both DBL-1 and DBL-1-independent mechanisms. The Smads may be regulated post-transcriptionally to contribute to the differential avoidance responses to various bacteria. Investigation of downstream genes regulated by Smads will further our understanding of the role of Smads in differential avoidance responses.

We next asked if DBL-1 signaling activity is altered in response to the bacterial panel. To address this question, we fed the bacterial panel to wild-type or dbl-1(-) animals expressing an integrated fluorescent DBL-1 pathway reporter called RAD-SMAD. This reporter consists of the GFP gene under the control of multiple copies of a Smad-binding element sequence (Tian et al., 2010). The expression of this reporter is robust in hypodermal nuclei of L2 stage wild-type animals, and changes in DBL-1 signaling positively correlate with hypodermal expression of RAD-SMAD (Tian et al., 2010; Fernando et al., 2011; Gleason et al., 2014; Liu et al., 2015; Savage-Dunn et al., 2019). We measured the hypodermal fluorescence intensity of the RAD-SMAD-expressing L2 populations fed control or test Gram-negative bacteria. Compared to the wild-type expression of RAD-SMAD fluorescence on E. coli control bacterial conditions (Figure 5A), reporter fluorescence in the wild-type background on Gram-negative bacteria E. cloacae and K. oxytoca was strikingly increased (Figure 5B, C, and H). While Gram-negative S. marcescens increased expression of the DBL-1 reporter in the population, it did not reach significance (p=0.49, Figure 5D and H).

In stark contrast, RAD-SMAD hypodermal fluorescence in wild-type animals was lost upon exposure to test Gram-positive bacteria. Because of the general absence of measurable hypodermal fluorescence, we compared the fraction of wild-type RAD-SMAD population exhibiting detectable GFP fluorescence when exposed to control or test Gram-positive bacteria. RAD-SMAD hypodermal fluorescence in wild-type populations was significantly reduced upon exposure to Gram-positive B. megaterium (0% had any detectable expression [n = 44]) or S. epidermidis (6% [n=66]) (Figure 5E, G, and I). RAD-SMAD hypodermal fluorescence was detected only in half the wild-type population fed E. faecalis (50% [n=52]), and fluorescence levels were much fainter in those animals with detectable RAD-SMAD in the hypoderm (Figure 5F and I, Supplementary file 2).

We then asked if the changes in the DBL-1 pathway reporter in response to the bacterial panel were dependent on the DBL-1 ligand. In dbl-1(-) L2 animals fed on E. coli, RAD-SMAD fluorescence in epidermal nuclei was not detectable or very faint, consistent with published reports (Supplementary file 2). Reporter intensity in dbl-1(-) animals on all Gram-negative bacteria remained mostly undetectable or faint, though a few animals surprisingly had wild-type fluorescence levels (Supplementary file 2). For animals lacking DBL-1 fed on any of the Gram-positive bacterial conditions, hypodermal RAD-SMAD fluorescence was undetectable or very faint. In general, these results indicate that animals modulate DBL-1 signaling depending on the specific bacterial challenge that animals encounter. In our panel, animals increased DBL-1 signaling in response to specific Gram-negative bacteria but repressed DBL-1 signaling in response to the tested Gram-positive bacteria. Of note, the increased expression of the RAD-SMAD reporter in some DBL-1-deficient animals on Gram-negative bacteria suggests that DBL-1 pathway signaling can be stimulated by pathogen response downstream of the DBL-1 ligand.

We then asked if this differential modulation of DBL-1 signaling activity translated to bacterial-specific downstream transcriptional responses. To identify the role of DBL-1 in differentially regulating transcription of downstream genes, we performed RNA sequencing using wild-type and dbl-1(-) animals exposed to control E. coli OP50, Gram-negative S. marcescens, or Gram-positive E. faecalis, building on established work that shows these bacteria alter innate immunity gene expression responses in wild-type C. elegans (Mallo et al., 2002; Alper et al., 2007; Liang et al., 2007; Roberts et al., 2010; Lakdawala et al., 2019). Animals were hatched and grown on E. coli OP50, synchronized as L4s, and then fed on the control or test bacteria for 48 hr before analysis. In animals lacking DBL-1 and fed control bacteria, 83 genes were significantly downregulated and 49 genes were significantly upregulated compared to the wild type (p<0.01, Figure 6A). Using WormCat, we identified enrichment of differentially expressed genes in stress response and extracellular material categories (Figure 6A). Some genes and gene classes previously reported to be regulated by DBL-1 at different developmental stages were also regulated by DBL-1 in the 2-day adult populations in this study (Figure 6—source data 1; Mallo et al., 2002; Liang et al., 2007; Roberts et al., 2010; Lakdawala et al., 2019).

In dbl-1(-) animals fed on Gram-negative S. marcescens, 102 genes were downregulated and 117 genes were upregulated. Gene enrichment analysis revealed that these genes belonged to genes regulated by multiple stresses, CUB-like domain genes, and C-type lectins (Figure 6B). Almost half of all genes that were identified as highly expressed in wild-type populations in response to S. marcescens were also identified in the dbl-1(-) populations, suggesting that DBL-1 plays a lesser role in the organismal response to S. marcescens (Figure 6—figure supplement 1A–C).

In dbl-1(-) animals fed on Gram-positive E. faecalis, 63 genes were downregulated and 64 genes were upregulated compared to the wild type. These genes were assigned to the functional category of genes regulated by multiple stresses upon analysis using WormCat (Figure 6C). This lower number of highly regulated genes in dbl-1(-) animals fed on E. faecalis is consistent with the reduced DBL-1 reporter fluorescence—and therefore decreased DBL-1 pathway signaling—in wild-type animals on E. faecalis (Figures 5 and 6C). While there was some overlap in the genes regulated in response to E. faecalis independent of DBL-1, we observed a higher number of genes differentially regulated in response to E. faecalis exposure in wild-type populations (Figure 6—figure supplement 1D–F).

Notably, some highly regulated genes in the wild-type strain were common in response to both pathogenic bacterial strains, but many more were specific to the bacterial challenge (Figure 6—figure supplement 1G). In the dbl-1(-) strain, there were also genes that were differentially regulated compared to the control that were either common to both S. marcescens and E. faecalis responses or unique to one or the other challenge (Figure 6—figure supplement 1H). Using WormCat, gene enrichment analysis of genes regulated in response to S. marcescens or E. faecalis exposure revealed differential regulation of sets of genes associated with pathogen and stress responses, among other gene classes (Figure 6—figure supplement 1A, B, D, and E; Holdorf et al., 2020). Comparison of the differentially expressed genes between wild-type and dbl-1(-) strains showed many common genes regulated in response to S. marcescens, indicating that their regulation is independent of DBL-1 signaling (Figure 6—figure supplement 1C). On the other hand, there is little overlap between the E. faecalis-enriched genes in the wild-type and dbl-1(-) strains, indicating the uniqueness of genes regulated in a DBL-1-dependent and independent manner (Figure 6—figure supplement 1F). These results suggest that animals use DBL-1 signaling to tailor transcriptional responses specific to the type of bacterial exposure.

We focused on the DBL-1-regulated genes that have known or putative roles in innate immunity (Figure 6D–F). Some genes were induced in wild-type animals upon exposure to S. marcescens or E. faecalis, and this induction was lost in dbl-1(-) animals. We also found some genes to be upregulated only upon loss of DBL-1 and not in the wild-type animals in response to S. marcescens or E. faecalis (Figure 6—figure supplement 1A–F). Some gene classes are highly regulated in response to these bacteria, but the specific genes within these families differed, including lysozyme, aspartyl protease, saposin-like, and C-type lectin genes (Figure 6D–F, Figure 6—figure supplement 1A, B, D, and E). These results suggest that DBL-1 is involved in regulating (positively and negatively) transcription of not only some innate immunity genes specific to the bacterial challenge, but also genes that are commonly regulated upon exposure to a variety of bacteria. This supports a role for DBL-1 signaling in differentially regulating molecular host responses to the bacterial challenge.

To determine if the expression of candidate target immunity genes is regulated by DBL-1 signaling in response to a wider variety of bacterial exposures, we used reporters of select immunity-related genes. Based on our RNA-sequencing dataset and previously published reports, we shortlisted genes based on their differential expression in either Gram-negative or Gram-positive-challenged animals (or both) and their requirement for DBL-1 signaling (or not) for their differential expression. This list of genes included dod-22, F55G11.7, irg-4, dod-24, and ilys-3. We measured and compared intestinal expression of these reporter genes in wild-type and dbl-1(-) backgrounds exposed to control or test bacteria. Animals were grown on E. coli, synchronized as L4s, and fed on control E. coli, Gram-negative, or Gram-positive bacteria for 48 hr before expression of these selected genes was measured and compared. dod-22 is a gene that is known to be regulated by the insulin-like signaling pathway transcription factor DAF-16 (Murphy et al., 2003). It is expressed primarily in the intestine and is involved in defense response to Gram-negative bacteria (Alper et al., 2007). RNA-sequencing analyses revealed dod-22 expression was induced in the wild type, but not in dbl-1(-) animals, only upon exposure to S. marcescens (Figure 6—source data 1). The dod-22 reporter was induced in the wild-type background in the presence of all Gram-negative test bacteria compared to the control but was not induced in response to the panel of Gram-positive bacteria. This induction of the dod-22 reporter on Gram-negative test bacteria was lost in the dbl-1(-) background, though loss of DBL-1 did not affect expression levels on the control E. coli. dod-22 reporter fluorescence remained largely unchanged in dbl-1(-) animals exposed to Gram-positive test bacteria (Figure 7A and B). These results indicate that DBL-1 is not required for the basal expression of dod-22 but is required for dod-22 induction on Gram-negative bacteria, which correlated with the RNA-seq data.

F55G11.7 is majorly expressed in the intestine and has been reported to be involved in innate immune responses to both Gram-negative and Gram-positive bacteria in C. elegans. Previous findings indicate that F55G11.7 is regulated by insulin, MAPK, and DBL-1 signaling pathways (Alper et al., 2007). Expression of F55G11.7 was reduced in dbl-1(-) mutants compared to wild-type fed on S. marcescens in our RNA-seq data (Figure 6—source data 1). Expression of the F55G11.7 reporter did not consistently change upon exposure to all test bacteria in the wild-type background except B. megaterium, where reporter expression was reduced (in two of three trials). We observed a significant reduction of F55G11.7 reporter activity in animals lacking DBL-1 except in response to B. megaterium and E. faecalis compared to wild-type animals. Additionally, a further reduction of F55G11.7 reporter fluorescence in dbl-1(-) animals was observed in response to Gram-negative S. marcescens and Gram-positive B. megaterium and S. epidermidis compared to the response on control bacteria (Figure 7C and D). These findings indicate that expression of F55G11.7 is not induced upon exposure to most of the test bacteria but is generally downregulated upon loss of DBL-1, expanding on previous findings that F55G11.7, which is induced upon S. marcescens Db11 and P. aeruginosa, is regulated by DBL-1 in response to pathogenic bacteria (Alper et al., 2007).

irg-4 is expressed in intestine and head neurons (Alper et al., 2007). It is known to be involved in defense responses to Gram-negative bacteria and has been shown to be regulated by insulin, MAPK, and DBL-1 signaling pathways (Shapira et al., 2006; Troemel et al., 2006; Nandakumar and Tan, 2008; Peterson et al., 2019). Based on our RNA-sequencing data, expression of irg-4 in wild-type animals was induced upon exposure to S. marcescens but was reduced in response to E. faecalis in comparison with the control bacteria. Furthermore, we found irg-4 to be a DBL-1-responsive gene. Expression of irg-4 was reduced in dbl-1(-) compared to wild-type animals fed on control, S. marcescens, and E. faecalis (Figure 6—source data 1). In the wild-type background, we observed a visible, reproducible induction of irg-4 reporter activity in response to Gram-negative S. marcescens, but not to K. oxytoca or E. cloacae (Figure 7E). Expression of the irg-4 reporter in wild-type animals is reproducibly unaltered in response to Gram-positive bacteria (Figure 7F). Loss of DBL-1 did not reproducibly alter irg-4 reporter activity in control conditions. However, irg-4 reporter induction in response to Gram-negative S. marcescens was lost in the dbl-1(-) background (Figure 7E and F). Indeed, irg-4 reporter expression was reduced compared to the wild-type background on all test bacteria except Gram-negative E. cloacae and Gram-positive E. faecalis. These results support that irg-4 is responsive to a broad range of bacteria and is regulated in part by DBL-1 signaling.

dod-24 is mainly expressed in the intestine (Yamawaki et al., 2010). dod-24 is regulated by the insulin-like signaling transcription factor DAF-16 in response to Gram-negative bacteria (Shapira et al., 2006). RNA-sequencing data indicated that dod-24 expression in wild-type animals was induced in response to S. marcescens and dampened in response to E. faecalis compared to wild-type animals fed on control bacteria. These changes in dod-24 expression were lost in dbl-1(-) animals (Figure 6—source data 1). We observed robust expression of dod-24 reporter activity in all tested Gram-negative bacteria, including the control, E. cloacae, K. oxytoca, and S. marcescens. Furthermore, dod-24 reporter activity was greater in response to E. cloacae and S. marcescens than to the control bacteria (Figure 7G). We observed a striking decrease of dod-24 reporter activity in wild-type animals exposed to all tested Gram-positive bacteria (B. megaterium, E. faecalis, and S. epidermidis) compared to the control bacteria (Figure 7H). Loss of DBL-1 resulted in a significant reduction of dod-24 reporter activity in control conditions. dod-24 reporter activity was also drastically reduced in all tested Gram-negative bacterial conditions to levels significantly lower than the wild type (Figure 7G). In the three Gram-positive conditions, loss of DBL-1 resulted in a further decrease of dod-24 reporter fluorescence relative to the wild type (Figure 7H). These results confirm that dod-24 is differentially expressed in response to an array of Gram-negative and Gram-positive bacteria and indicate that DBL-1 signaling plays a major role in regulating dod-24 expression.

ilys-3, which encodes a lysozyme, is expressed in the intestine, pharynx, and coelomocytes. It is induced in response to Gram-positive bacteria (Gravato-Nobre et al., 2005). By RNA-sequencing, we confirmed expression of ilys-3 was induced in wild-type animals exposed to Gram-positive E. faecalis but not Gram-negative S. marcescens. This induction was lost in dbl-1(-) mutants (Figure 6—source data 1). ilys-3 reporter activity in wild-type animals was unchanged or reduced in response to the tested Gram-negative bacteria compared to the control (Figure 7I). We observed induction of ilys-3 reporter activity upon exposure to all tested Gram-positive bacteria (B. megaterium, E. faecalis, and S. epidermidis) (Figure 7J). Loss of DBL-1 did not alter ilys-3 reporter activity in control conditions. The ilys-3 reporter activity remained at relatively low levels in animals lacking DBL-1 exposed to test Gram-negative bacteria (Figure 7I). However, ilys-3 reporter activity also remained at relatively low levels upon loss of DBL-1 in response to Gram-positive bacteria B. megaterium and S. epidermidis but was wild type in response to E. faecalis (Figure 7J), consistent with our RNA-seq results. These results suggest that while DBL-1 is not required for basal levels of ilys-3 expression, it is required for the induction of ilys-3 expression in response to some Gram-positive bacteria.

Together, these results expand on the RNA-sequencing dataset and validate some of the RNA-sequencing results. These data provide a snapshot of the gene expression profile of the host upon exposure to a variety of bacteria. These findings collectively indicate that DBL-1 signaling also tailors defense responses to specific bacteria by regulating antimicrobial gene expression.

Lifespan assay was performed as previously described (Sifri et al., 2003; Reddy et al., 2009; Amrit et al., 2014). Concentrated bacterial cultures were spread on 6-cm-diameter EZ media plates (full lawn plates) containing 50 µg/ml FUdR to cover the surface of the plates entirely. FUdR, which prevents reproduction by blocking DNA synthesis, was used to prevent offspring from confounding the scoring. Wild-type and dbl-1(-) animals (n ≥ 30) were fed on control and test bacteria on full lawn plates at the L4 stage in quadruplicate. The plates were scored for live and dead nematodes every 24 hr until all animals were dead. Animals were scored as dead if they did not respond to gentle touch with a sterilized platinum wire and were removed from the plate. At least three independent trials were performed. Worms that died by desiccating on the walls of the plates were censored from the analysis.

The intestinal barrier function assay was performed as previously described (Kissoyan et al., 2019). Wild-type and dbl-1(-) L4 animals were fed on control and test bacteria on full lawn plates. The assay was performed when about 50% of the population with the lowest mean lifespan remained alive. At least 15 animals were sampled at the specified times to examine intestinal tissue integrity. The intestinal barrier integrity was assessed using a blue dye, erioglaucine disodium salt (5% wt/v), as an indicator of tissue integrity as the animals age. Leaking of this blue dye outside the intestinal lumen indicates reduced intestinal integrity. The animals were washed with S buffer and were incubated in erioglaucine disodium salt solution in a 1:1 ratio for 3 hr. The animals were then washed thrice with S buffer and were mounted on 2% agarose pads on glass slides. 10 µM levamisole was added to paralyze the animals. The animals were imaged on a Nikon DS-Ri2 camera mounted on a Nikon SMZ18 dissecting microscope. The leakiness of the intestine was assessed and scored for no leakage/no Smurf, mild leakage/mild Smurf, and severe leakage/severe Smurf phenotypes. The experiment was performed in at least three independent trials for each experimental condition.

The bacterial colonization assay was performed using a method adapted from Portal-Celhay et al., 2012; Eng and Nathan, 2015. Wild-type and dbl-1(-) L4 animals were fed on S. marcescens on full lawn plates. The assay was performed when about 50% of the population with the lowest mean lifespan remained alive. Five animals in quadruplicates were picked and washed in 5 µl drops of M9 buffer containing 25 mM levamisole to paralyze animals and inhibit their pharyngeal pumping and expulsion. The animals were washed with thrice more with M9 buffer. The washed animals were placed in 1.5 ml Eppendorf tubes containing 120 µl M9 buffer and 100 µl was aliquoted from the tube to and incubated at 37°C overnight on tryptic soy agar plates to yield background bacterial counts. To the animals remaining in the tube, 30 µl M9 buffer containing 1% Triton X-100 was added and the animals were mechanically disrupted by using a pestle. This homogenized worm lysate was incubated overnight at 37°C on tryptic soy agar plates to yield bacterial counts. Bacterial colony-forming units (CFUs) per worm were calculated using the formula: (colony number × dilution factor)/(5 animals × 100 μl lysate plated). Finally, CFU per worm of the background was subtracted from the CFU per worm of the worm lysate to compare bacterial colonization between wild-type and dbl-1(-) populations.

Wild-type and dbl-1(-) L4 animals (n = 12) were fed on control and test bacteria on full lawn plates. The number of contractions of the pharyngeal bulb was counted for 20 s to calculate the pharyngeal pumping rate of animals. Two counts were made and averaged for each animal. Three independent trials were performed in triplicate (Clark et al., 2018).

Microbial avoidance assays were performed as previously described (Chang et al., 2011). Then, 20 µl of the concentrated bacterial cultures were spotted on 6 cm diameter EZ media plates and incubated at 37°C overnight. Wild-type, dbl-1(-), sma-2(-), sma-3(-), and sma-4(-) L4 hermaphrodites were placed on control and test bacterial lawn (n = 30 per condition/trial, performed in triplicate). The plates were scored for number of worms occupying the lawn at the indicated time points. Three independent trials were performed. The avoidance ratio (A) was calculated using the formula: A = number of animals off the lawn/total number of animals.

Animals were synchronized as embryos by bleaching mixed-stage populations (Stiernagle, 2006). Total RNA was extracted from animals at 48 hr after the L4 stage. Total RNA was extracted by the freeze cracking method as previously described (Portman, 2006).

RNA from wild-type and dbl-1(-) adult populations fed on control and test bacteria was extracted in three independent trials. Sequencing libraries from the extracted RNA were generated using the NEBNext RNA Library Prep Kit for Illumina (NEB, USA) following the manufacturer’s recommendations. Also, 1 μg RNA of each sample was used as input material for the RNA sample preparations. Novogene performed RNA sequencing of samples. Differential expression analysis of wild-type compared to dbl-1(-) populations grown on different bacteria was performed using the DESeq R package (1.18.0) (Love et al., 2014). Genes with an adjusted p-value<0.01 found by DESeq were assigned as differentially expressed.

After RNA isolation, cDNA was synthesized and quantitative real-time PCR was performed as previously described (Madhu et al., 2020). 2 µg of total RNA isolated was primed with oligo(dT) and reverse transcribed to yield cDNA using the SuperScript III reverse transcriptase kit as per the manufacturer’s protocol (Invitrogen). Real-time PCR was performed on a QuantStudio3 system (Applied Biosystems by Thermo Fisher Scientific) using the PowerUP SYBR Green master mix (Applied Biosystems) according to the manufacturer’s instructions. Three independent biological trials were performed. Each biological trial was performed in three technical replicates for each condition. Primer sequences are available in Supplementary file 4. QuantStudio Design and Analysis Software v1.5.1 was used to calculate raw C_t values. The C_t values for the target genes were normalized to the housekeeping gene act-1 (actin) (Applied Biosystems by Thermo Fisher Scientific). Fold change in gene expression between experimental sample and the control was determined by using the 2^(-ΔΔC_t⁾ method.

RAD-SMAD reporter strains were placed on full lawns of the control or test bacteria at the L4 stage. L2 progeny were mounted on 2% agarose pads and anesthetized by using 1 mM levamisole, and fluorescence was captured by a Zeiss LSM 900 confocal microscope using a 40× oil objective. At least 10 L2 animals with at least 5 hypodermal nuclei per worm in the focal plane were imaged per condition when fluorescence was visible, giving a moderate effect size as determined by power analysis. The experiment was performed in three independent trials. The microscope conditions were optimized with respect to the control and test conditions and kept consistent within each trial. Mean fluorescence intensities of the five hypodermal nuclei in each animal were calculated, and the average intensity of each strain was determined as previously described using Zeiss ZEN lite software (Savage-Dunn et al., 2019). Data was statistically analyzed by unpaired t-test.

The innate immune reporter strains were transferred to full lawns of the control and test bacteria at the L4 stage and were imaged after 48 hr of exposure. Fluorescence of the reporter strains was captured by a Nikon DS-Ri2 camera mounted on a Nikon SMZ18 dissecting microscope. Animals were mounted on 2% agarose pads and anesthetized with 1 mM levamisole. At least 15 animals were imaged per condition as determined by power analysis with a moderate effect size. The microscope conditions were optimized with respect to the control and test conditions and kept consistent within each trial. However, imaging exposure times were different between some trials to prevent saturation of signal in experimental conditions. Three independent trials were performed. Mean intestinal fluorescence intensities were measured using the Nikon NIS Elements AR v5.02 software.

Lifespans of C. elegans populations were calculated by the Kaplan–Meier method, and statistical analysis was performed using the two-tailed log-rank test. The average pharyngeal pumping rates were analyzed using two-way ANOVA using Tukey’s post hoc test. Bacterial colonization counts (CFU per worm) were compared using the two-tailed unpaired t-test. The intestinal barrier function phenotypes and percentages of animals expressing RAD-SMAD reporter exposed to Gram-positive bacteria were statistically analyzed by the chi-square test. The avoidance ratio was compared by repeated measures ANOVA using Tukey’s post hoc test. qRT-PCR values and mean fluorescence intensities were evaluated using ANOVA with Dunnett’s multiple-comparisons test. RNA-sequencing analysis was performed using the DESeq R package (1.18.0). The resulting p-values were adjusted using the Benjamini and Hochberg’s approach for controlling the false discovery rate.