Organismal survival and evolutionary success require organisms to balance the demands of defense against infection with the demands of development and reproduction. Research from several organisms has demonstrated that there can be trade-offs between the demands for immunity and the demands for growth. For example, enhancing immune responses in plants often leads to impaired growth and lowered crop yields, a trade-off that is regulated by secondary messengers and phytohormones [1][2][3]. Similarly, immune pathways have been shown to impact the development of the nematode Caenorhabditis elegans. For example, upregulation of the PMK-1 p38 MAPK immune pathway, such as by mutations in the genes nipi-3 and olrn-1, is detrimental to development [4][5][6][7][8]. However, there is still much to be learned about the mechanisms by which animals tune the balance between growth and defense in order to promote their overall survival and reproduction.
Recently, we described a genetic switch in C. elegans that regulates a balance between growth and immunity [9,10]. Through forward genetic screening, we found that the pals-22 gene represses a set of immunity-related genes called the Intracellular Pathogen Response (IPR) genes, which are activated upon infection by natural intracellular pathogens of the C. elegans intestine-the Orsay virus and the fungus Nematocida parisii from the microsporidia phylum [11][12][13][14][15]. In the absence of infection, mutations in pals-22 lead to constitutive expression of IPR genes, increased immunity against intracellular pathogens, and slowed growth. Through suppressor screens, we found that mutations in the pals-25 gene reverse pals-22 mutant phenotypes back to wild-type, indicating that pals-25 is required for mediating the effects caused by the loss of pals-22 [9]. Thus, the antagonistic paralogs pals-25 and pals-22 comprise an ON/OFF switch module that increases immunity with an associated cost of slowed growth.
Genes in the pals family encode proteins that contain the ALS2CR12 domain, a divergent amino acid signature of unknown biochemical function [11,16]. Genes associated with immunity often undergo species-specific expansion [17,18], and accordingly, there are 39 pals genes found in C. elegans, and only a single pals gene found each in mice and humans, with no reported function in those organisms. The C. elegans pals-22 and pals-25 genes are co-expressed in an operon, and most pals genes in C. elegans are clustered together in the genome, suggesting that they resulted from gene duplication events. The pals gene family exhibits sequence divergence and species-specific changes in gene number within species of the Caenorhabditis genus. While C. elegans has 39 pals genes, there are 18 pals genes in C. remanei, and only 8 pals genes each in C. briggsae and C. brenneri [16]. Of the 39 pals genes in C. elegans, 26 of them are upregulated by IPR triggers, including the pals-5 gene, which is commonly used as a reporter for the IPR (pals-5p::GFP) [11]. Notably, pals-22 and pals-25 are not induced by infection, leading to the hypothesis that non-induced pals genes serve as regulators of induced pals genes.
Here, through a forward genetic screen for mutants that have constitutive pals-5p::GFP expression in the absence of infection, we identify pals-17 as a new negative regulator of IPR gene expression, acting independently of pals-22. Through targeted RNA interference (RNAi) screening of non-induced pals genes, we identify pals-20 and pals-16 as two positive regulators that act downstream of pals-17 to induce IPR gene expression. Notably, pals-17, pals-20, and pals-16 are not induced by infection, similar to pals-22 and pals-25, supporting the hypothesis that non-induced pals genes are regulators of induced pals genes. We show that PALS-17 and PALS-20 proteins are co-expressed in the intestine. We also find that point mutations in pals-17 lead to slowed growth and increased immunity against natural intracellular pathogens of the intestine, while a full pals-17 deletion leads to developmental arrest. The immunity and growth phenotypes of pals-17 mutants are dependent on the positive IPR regulators pals-16 and pals-20. Consistent with pals-20 being an activator of the IPR, we find that overexpression of pals-20 in a wild-type background promotes resistance to viral infection. Altogether, these results identify PALS-17 as a negative regulator of the IPR similar to PALS-22, and PALS-16 and PALS-20 as positive regulators of the IPR similar to PALS-25. These findings indicate there are two separate modules of IPR regulators including the PALS-22/25 module and PALS-16/17/20 module, highlighting how PALS proteins in C. elegans have extensive control over the balance between growth and immunity, which is critical for organismal success.
To identify novel regulators of the IPR, we performed a forward genetic screen for mutations that cause constitutive expression of the transcriptional IPR reporter pals-5p::GFP (jyIs8). Here, we mutagenized a strain that has loss-of-function mutations in pals-22 and pals-25, two previously identified regulators of the IPR. We performed the screen in this double mutant to prevent isolation of new alleles of pals-22 or pals-25. By screening 33,504 haploid genomes, we recovered four alleles exhibiting constitutive pals-5p::GFP expression, all of which complemented each other. In this study, we focus on the jy74 allele, which shows a strong constitutive expression of the pals-5p::GFP reporter in intestinal epithelial cells in the absence of infection (Fig . Whole-genome sequencing of jy74 mutants identified a mutation in the gene pals-17 (Fig 1E). Specifically, the jy74 allele has guanine 325 mutated into adenine, which is predicted to affect splicing between the third and fourth exons of pals-17. pals-17, like pals-22 and pals-25, belongs to the clade of pals genes that are not induced upon intracellular infection [9][10][11][12]. To confirm that the mutation in pals-17 is responsible for the upregulated expression of the pals-5p::GFP reporter, we introduced a fosmid containing a wild-type copy of pals-17 into pals-22(-) pals-25(-) pals-17(jy74) mutants. We observed a decrease in pals-5p:: GFP expression in animals carrying the pals-17 fosmid array compared to their siblings without the array (Fig 1F). This observation indicates that loss of pals-17 function is responsible for constitutive pals-5p::GFP expression in jy74 mutants.
To further confirm the role of pals-17 in pals-5p::GFP regulation, we treated pals-22 pals-25 double mutant animals with pals-17 RNAi. Upon knockdown of pals-17, we observed increased expression of the pals-5p::GFP reporter, consistent with pals-17 acting as a negative regulator (Fig 1G and 1H and S1 Table). Next, we investigated whether loss of pals-17 induces pals-5p::GFP expression in the presence of functional pals-22 and pals-25. Here, we performed the same RNAi analysis in a wild-type background and observed that pals-5p::GFP expression was significantly induced following pals-17 RNAi treatment (Fig 1I and 1J and S1 Table). We next examined if the loss of pals-17 upregulates ZIP-1::GFP expression. In recent work, we demonstrated that ZIP-1 is a transcription factor that promotes immunity against intracellular pathogens. IPR triggers require ZIP-1 for induction of the pals-5p::GFP reporter and increase ZIP-1::GFP nuclear expression [10,19]. Here, we found that pals-17 RNAi induces expression of ZIP-1::GFP in intestinal nuclei (S1 Fig), similar to other IPR triggers [10,19].
To determine the null phenotype of pals-17, the entire coding region of pals-17 was deleted to generate the pals-17(Δ) allele (S2A Fig) . We found that these pals-17(Δ) null mutants failed to develop past an early larval stage (approximately first larval stage L1 to second larval stage L2), indicating that pals-17 is an essential gene (S2B Fig) . To quantify this effect, we measured the body length of progeny from a balanced pals-17(Δ)/sC1(s2023)[dpy-1(s2170) umnIs21] strain and observed significantly smaller body size for pals-17 homozygotes in comparison to their pals-17 heterozygous siblings, Dpy (wild-type pals-17) siblings, and wild-type control animals (S2C Fig and S1 Table ). Notably, we observed that this larval arrest phenotype is 100% penetrant, as all pals-17(Δ) homozygotes failed to develop into adults (60/60 from three replicates). In summary, our analyses demonstrate that pals-17 is a negative regulator of the IPR reporter and is essential for C. elegans development.
The analysis above indicates that wild-type pals-17 acts as a repressor of IPR gene expression, similar to wild-type pals-22. Activation of the IPR in pals-22 mutants requires the pals-25 gene, which is immediately downstream of pals-22 in the genome as part of an operon [9]. Therefore, we hypothesized that a gene positioned immediately downstream of pals-17 might antagonize pals-17 function, similar to how pals-25 antagonizes pals-22 function. Because pals-20 is the closest neighbor of pals-17 in the genome (although they are not in an operon), we performed RNAi against pals-20 in pals-22(-) pals-25(-) pals-17(jy74) mutants and examined pals-5p::GFP expression. Notably, we observed a decrease in GFP reporter expression in pals-20 (RNAi) animals compared to control RNAi, indicating that pals-20 may be an activator of the IPR (Fig 2A).
To further investigate the function of pals-20, a mutant allele that contains a premature stop codon in the first exon of pals-20 was generated (pals-20(*); see Materials and Methods for more information) (Fig 2B). Similar to pals-25 loss-of-function alleles, the pals-20(*) loss-offunction allele had no apparent phenotype in a wild-type background. Next, we investigated whether pals-20(*) would suppress pals-17 mutant phenotypes. Because a full deletion of pals-17 led to larval arrest, we sought another pals-17 allele, with the goal of creating a mutation that would cause a strong loss-of-function, but allow animals to develop into adulthood. To animals (right), compared to their pals-17(jy74) mutant siblings that do not contain the fosmid (left). Red, green and DIC channels were merged in the composite images. myo-2p::mCherry and myo-3p::mCherry co-injection markers are shown in red. Scale bar, 100 μm. (G-J) pals-17 RNAi increases pals-5p::GFP expression in pals-22(-) pals-25(-) mutants and wild-type animals. Representative images of synchronized L4 stage populations of the pals-22(-) pals-25(-) (G) and wild-type animals (I) grown on the control RNAi vector and pals-17 RNAi plates. Red, green and DIC channels were merged. myo-2p::mCherry coinjection marker is shown in red. Scale bars, 50 μm. (H, J) Box-and-whisker plots of pals-5p::GFP expression levels standardized to TOF following control RNAi and pals-17 RNAi treatments of pals-22(-) pals-25(-) mutants (H) and wild-type animals (J). (C, D, H, J) Lines in box-and-whisker plots represent the medians, box bounds indicate 25 th and 75 th percentiles, and whiskers extend from the box bounds to the minimum and maximum values. Green dots represent individual values for each animal; 600 (C), 800 (D), 1200 (H) and 750 animals (J) were analyzed per strain per each of the three replicates. A Kolmogorov-Smirnov test was used to calculate p-values; **** p < 0.0001. https://doi.org/10.1371/journal.ppat.1011120.g001
achieve this goal, the pals-17(*) allele was generated, which contains an early stop in the third exon in pals-17 (Fig 2B). This mutation causes slowed and asynchronous development but allows some mutant animals to reach adulthood and reproduce (Fig 2C and S1 Table). Here, we found that the pals-20(*) mutation substantially suppressed the developmental delay and asynchronous development of these pals-17(*) mutants. pals-17(*) pals-20(*) double mutants had slightly slower developmental rates than pals-20(*) single mutants and the wild-type control, but they developed significantly faster than pals-17(*) mutants (Fig 2C and S1 Table). To better understand the developmental delay of pals-17 mutants and determine conditions for better synchronization of these animals for further analyses, we measured the gonad development of pals-17 and pals-20 mutant strains. We found that gonad development in pals-17(*) mutants is delayed and highly asynchronous, while pals-17 pals-20 double mutants at the L2 stage have similar gonad areas to wild-type and pals-20 single mutants (S3 Fig and S1 Table ). Taken together, these results show that the loss of pals-20 can suppress the developmental delay and asynchrony of pals-17 mutants.
We next examined the endogenous mRNA expression of IPR genes at different developmental stages in pals-17, pals-20, and pals-17 pals-20 double mutants. First, we performed qRT-PCR analysis in L1 animals. pals-20 single mutants had wild-type levels of IPR gene expression, indicating that pals-20 does not regulate IPR gene expression in a wild-type background (Fig 2D and S1 Table). We found significant upregulation of IPR genes in the pals-17 mutant background, which was almost completely suppressed in pals-17 pals-20 double mutants (Fig 2D and S1 Table). Next, we found that the upregulated IPR gene expression observed in pals-17 mutants was only partially suppressed in pals-17 pals-20 double mutants at the L2 stage (Fig 2E and S1 Table). Finally, at the L4 stage, we surprisingly observed no suppression of IPR gene induction in pals-17 mutants by loss of pals-20 (Fig 2F and S1 Table). Thus, we found that endogenous mRNA levels of multiple IPR genes are highly induced in pals-17 mutants from the L1 to the L4 stage. This induction is dependent on pals-20 early in life but is less dependent on pals-20 later in life.
Because IPR induction in pals-17 mutants is dependent on the pals-20 gene only in early life, we screened for a gene that plays a role in IPR induction later in life. Here, we used RNAi to test several previously uncharacterized pals gene family members, focusing on those that are not strongly induced by intracellular infection [11,16]. These genes included pals-1, pals-13, pals-16, pals-18, pals-19, pals-23, pals-24, pals-31, pals-34, pals-39 and pals-40 (S4A Fig) . In this assay, we used pals-17 pals-20 double mutant animals, which express pals-5p::GFP in the intestine at the adult stage, to look for an RNAi clone that would decrease GFP expression. We found that pals-16 RNAi significantly decreased pals-5p::GFP expression (Fig 3A and 3B and S1 Table), suggesting that pals-16 acts downstream of pals-17 later in life. Of note, PALS-20 and PALS-16 proteins cluster together in a phylogenetic tree of PALS proteins, sharing 52% identity in amino acid sequence (S4 Fig and S2 Table ).
To further examine the role of pals-16, we deleted the entire pals-16 coding region in wildtype animals, pals-17 mutants, and pals-17 pals-20 double mutants (Fig 3C). We found that the deletion of pals-16, called pals-16(Δ), was able to rescue the developmental defects of pals-17 mutants. In contrast, pals-16(Δ) did not have any obvious effects in the other two backgrounds that have wild-type rates of development (Fig 3D and S1 Table). We then measured endogenous mRNA expression of IPR genes with qRT-PCR at L1 and L4 stages to test if and when pals-16(Δ) can suppress pals-17-dependent IPR induction. We found that the loss of pals-16 leads to a decrease in mRNA levels of multiple IPR genes that are upregulated in pals-17 pals-20 double mutants at the L4 stage (Fig 3E and S1 Table). Notably, we found that pals-16(Δ) is also able to suppress IPR gene induction in the pals-17 single mutant background, independent of pals-20. Furthermore, pals-16(Δ) was able to suppress IPR gene expression in pals-17 mutant animals at the L1 stage (Fig 3F and S1 Table). Taken together, our data demonstrate that pals-16(Δ) suppresses IPR induction in pals-17 mutants throughout development and in a manner independent of pals-20.
Next, we used RNA sequencing (RNA-seq) to examine genome-wide transcriptional changes in pals-17, pals-20, and pals-16 mutants. We performed our RNA-seq analysis at the L4 stage because this larval stage had the best RNA quality. First, we compared pals-17 mutants to wildtype animals, growing them for different lengths of time in order to harvest both strains at approximately the L4 stage. Unfortunately, pals-17 mutants have a high degree of individual variability in developmental time (S3 Fig); thus, it was not possible to obtain a pure population of L4 animals. Instead, we harvested a population of animals with the average stage of L4 that also contained some older and younger animals. In RNA-seq analysis of these samples, we found that 3,001 genes were upregulated in pals-17 mutants compared to wild-type animals, which significantly overlap with previously defined IPR genes (70/80 IPR genes) and overlap with genes upregulated in pals-22(jy3) mutants (262/457) (Fig 4A and S1, S3 and S4 Tables) [10]. However, many of the genes upregulated in pals-17 mutants could be differentially expressed due to the wide range of developmental ages found in a population of these mutants (S3 Fig) . To address this issue, we analyzed mRNA expression of pals-17 pals-20 mutants. These animals have upregulated IPR gene expression at the L4 stage similar to pals-17 mutants, but they grow more synchronously within a population, at a rate comparable to wild-type animals (Figs 2 andS3). Here, we found that 190 genes were upregulated in pals-17 pals-20 double mutants compared to the wild type at L4 (Fig 4B). The majority of these genes were also upregulated in pals-22 mutants (188/190), and they significantly overlapped with IPR genes (66/80 IPR genes) (Fig 4B and S1, S3 and S4 Tables). Furthermore, most of the genes upregulated in pals-17 pals-20 double mutants (172/190) were also upregulated in pals-17 single mutants (Fig 4C and S1 Table). To identify genes regulated by pals-16, we also performed RNA-seq on pals-16 pals-17 double and pals-16 pals-17 pals-20 triple mutant animals. We found that the upregulation of genes in pals-17 pals-20 mutants was almost completely suppressed by the loss of pals-16 (182/190) (Fig 4C andS1, S3 and S4 Tables).
To obtain insight into biological processes and cellular structures relevant to the genes upregulated in pals-17 and pals-17 pals-20 mutants, we performed analysis using the WormCat program, which is specifically designed to analyze C. elegans genomics data [20]. We found that some of the genes upregulated in pals-17 mutants belong to the category of genes that encode proteins with an ALS2CR12 domain-the defining hallmark of the PALS protein family [16], many of which are induced as part of the IPR (Figs 4D and S4A). Other categories included genes involved in protein degradation (F-box proteins), transcriptional regulation, reproduction, and signaling (Fig 4D and S1 Table). We analyzed the role of pals-17 in reproduction in a brood size assay, and we found that pals-17 mutants lay significantly fewer eggs (S5 Fig and S1 Table ). Furthermore, this phenotype was suppressed by the loss of pals-20. The ).
Next, we examined the correlation between pals-17-and pals-17 pals-20-dependent genes and gene sets that were previously associated with IPR activation, infection, and stress responses. We performed Gene Set Enrichment Analysis (GSEA) [21,22] and found that pals-17 and pals-17 pals-20 mutants upregulate genes in common with infections by intracellular pathogens as well as known activators and regulators of the IPR (Fig 4F and S1 and S5 Tables). Furthermore, we found that genes upregulated in pals-17 mutants significantly correlate with genes that are differentially expressed (both upregulated and downregulated) in animals infected with other pathogens that are not associated with the IPR. pals-17 pals-20 mutants upregulate genes in common with several other stress responses. In summary, we found that the loss of pals-17 causes extensive changes in the transcriptional profile of animals, leading to the upregulation of the majority of IPR genes and genes associated with previously characterized immune and stress responses.
To examine where these newly identified regulators of the IPR are expressed, we created transcriptional and translational reporters for pals-17 and pals-20. To first determine the tissues where they are expressed, we examined the expression of pals-17p::GFP and pals-20p:: wrmScarlet transcriptional reporters at the L4 stage. Both transgenes were expressed in intestinal tissue, especially in the anterior-most and posterior-most intestinal cells (Fig 5A and5B). Furthermore, we found that both reporters were expressed in the head region. pals-17p::GFP signal was present in head neurons (Fig 5A ), whereas pals-20p::wrmScarlet was sporadically and weakly expressed in the head region (Fig 5B).
To examine where PALS-17 and PALS-20 proteins are expressed and if they colocalize in intestinal cells, we examined PALS-17::GFP and PALS-20::wrmScarlet translational reporters co-expressed in the same animals. We found that these two proteins formed puncta in a subset of intestinal cells, and in most cases, they colocalized with each other (Figs 5C andS6). Furthermore, we observed fluorescent signals from both reporters at the apical border of the intestinal cells, lining the intestinal lumen. PALS-17 and PALS-20 were also diffusely expressed in the cytosol in some intestinal cells (Figs 5C andS6). To confirm that these translational reporters are functional, we tested if the expression of PALS-17::GFP can suppress the growth defects of pals-17 mutants and if PALS-20::wrmScarlet overexpression slows down the development of pals-17 pals-20 double mutants. Here, we found that both constructs are functional because their expression had significant effects on the growth of animals carrying the array, as compared to their non-transgenic siblings (S7 Fig and S1 Table). Unfortunately, we were unable to amplify or synthesize the DNA region upstream of pals-16 to obtain a pals-16 reporter, likely because of the repetitive and complex sequence in this region. In summary, we found that PALS-17 and PALS-20 proteins colocalize in the intestine, which is the tissue invaded by viral and microsporidian pathogens that activate the IPR.
IPR induction in the pals-22 mutant background leads to increased resistance against intestinal intracellular pathogens, namely the Orsay virus and the microsporidian species N. parisii. Therefore, we examined whether pals-17 mutants show increased resistance against these pathogens, and if so, whether this resistance is pals-20-and/or pals-16-dependent. First, we analyzed Orsay virus infection levels after infecting L1 animals for 18 hours and found a significantly lower infection level in pals-17 mutants (Fig 6A and S1 Table). Loss of either pals-20 or pals-16 suppressed this phenotype, suggesting that both genes contribute to the increased viral resistance of pals-17 mutants. We also examined antiviral resistance by infecting L4 animals and measuring their infection levels at 24 hours post-inoculation (hpi). To increase the overall infection level, animals were subjected to rde-1(RNAi) from the L1 stage. rde-1 plays an important role in antiviral immunity as a critical component of the RNAi pathway, but it does not have a known role in regulating the IPR [23]. Again, we saw that pals-17 mutants showed significant resistance to Orsay virus infection, which was rescued by the loss of pals-20 and/or pals- 16 (Fig 6B and S1 Table). This result indicated that pals-20 is still required for immunity at later stages of life, although its role in IPR gene expression seems to be restricted to younger animals.
Next, we analyzed the resistance of pals mutants to the microsporidian species N. parisii. We fed animals N. parisii spores at the L1 and L4 stages, and then examined pathogen load in the intestine at 30 hpi. At this time point, N. parisii develop into multinucleate proliferative forms known as meronts. We observed significantly lower pathogen load in pals-17 mutants at both analyzed stages (Fig 6C and 6D and S1 Table). The intestinal cells of pals-17 pals-20 double mutants that were inoculated with pathogen from the L1 stage also had a significantly lower amount of meronts relative to their size in comparison to the wild-type control. However, the pathogen load of pals-17 pals-20 double mutants was higher than in pals-17 single mutants, indicating partial suppression of this phenotype by pals-20 at the L1/L2 stage. Interestingly, pals-17 pals-20 double mutants inoculated at the L4 stage had wild-type pathogen load levels, again suggesting that pals-20 promotes resistance at later stages of life. A pals-16 mutation fully suppressed the increased pathogen resistance observed in pals-17 mutants at both time points.
In order to assess feeding rate of pals-17 mutants, we fed fluorescent beads to L1 animals and measured bead accumulation in the intestinal lumen. We found that pals-17 mutants have significantly reduced fluorescent bead accumulation compared to wild-type animals (S8A Fig and S1 Table ). Of note, pals-17 pals-20 double mutants also showed significantly lower fluorescent bead accumulation, although these mutants had wild-type infection levels in many assays. This result suggests that feeding rate may not be a determining factor in these infection assays.
To address the concern that the decreased pathogen load in pals-17 mutants, dependent on IPR activators like pals-20, was solely due to decreased feeding rate and thus reduced exposure to pathogens, we examined the pathogen resistance and the feeding rate of a wild-type strain overexpressing a functional PALS-20::wrmScarlet construct. Importantly, unlike in the pals-17 pals-20 double mutant background, we did not observe any effect of PALS-20::wrmScarlet overexpression on the developmental rate of wild-type animals, and these animals had normal rates of fluorescent bead accumulation (S8B Fig and S1 Table ). Then, we tested if PALS-20:: wrmScarlet overexpression had an effect on IPR activation. Here, we found a high induction of IPR reporter expression in pals-20 overexpressing animals in comparison to their wild-type siblings (Fig 6E and 6F and S1 Table). Next, we investigated Orsay virus infection levels in these two groups of animals and found that PALS-20::wrmScarlet overexpression leads to a ). This result confirms the role of pals-20 in controlling immunity against intracellular pathogens.
Next, we investigated whether pals-17 and pals-20 act in the intestine. We created transgenic lines carrying pals-17::gfp and pals-20::wrmScarlet under the intestine-specific promoter vha-6. First, we found that intestinal-specific expression of pals-17 in a pals-17 mutant background caused accelerated development, indicating pals-17 can act in the intestine (S9A Fig and S1 Table ). Also, we found that intestinal-specific expression of pals-20 in a pals-17 pals-20 mutant background caused slightly slowed development (S9B Fig and S1 Table ). Importantly, we found that intestinal-specific expression of pals-20 in a wild-type background was sufficient to significantly reduce the level of Orsay virus infection, while not impairing feeding rate (S9C and S9D Fig and S1 Table ). These results indicate that pals-20 can act in the intestine to regulate pathogen resistance.
In summary, loss of pals-17 leads to significantly lower infection levels with both the Orsay virus and microsporidia. Loss of pals-20 and pals-16 suppress infection phenotypes of pals-17 mutants in different stages of life, and overexpression of pals-20 provides enhanced immunity against infection. Finally, tissue-specific expression of pals-17 and pals-20 indicates that these proteins can function in the intestine.
In this study, we identified three previously uncharacterized genes pals-17, pals-20, and pals-16 as important new regulators of the balance between immunity and development ( Fig 7). The expanded pals gene family in C. elegans consists of two groups of genes: genes that are upregulated upon infection with obligate intracellular viral and fungal pathogens-"Induced pals genes", and genes that do not change expression levels upon infection-"Uninduced pals genes". pals-17, pals-20, and pals-16 all belong to the latter group, together with the previously characterized genes pals-22 and pals-25, which act as antagonistic regulators of the IPR and growth (S4A Fig) [9]. Through a forward genetic screen, we identified pals-17 as a negative regulator of the IPR. We demonstrate that pals-17 is essential for normal development, as deletion of pals-17 leads to larval arrest. Our RNAi suppressor studies identified two other pals family members, pals-20 and pals-16, which function as pals-17 antagonists by promoting IPR activation and inhibiting normal larval development. The negative regulator protein PALS-17 and the positive regulator protein PALS-20 colocalize in intestinal epithelial cells in discrete cytoplasmic puncta, as well as along the apical plasma membrane. Finally, we demonstrate that loss of pals-17 and overexpression of pals-20 protect animals against infection by intracellular pathogens. Because previous studies have only functionally characterized two out of 39 pals genes, our findings have more than doubled the number of pals genes with known functions. and maximum values. A Kruskal-Wallis test was used to calculate p-values; **** p < 0.0001. (E, F) pals-5p::7xGFP reporter expression is induced in animals overexpressing pals-20::wrmScarlet construct. (E) Representative images of synchronized animals 72 hours-after L1 overexpressing pals-20::wrmScarlet myo-2p::mCherry array and their siblings that do not overexpress pals-20 (control). Red, green and DIC channels were merged. myo-2p::mCherry co-injection marker is shown in red. PALS-20:: wrmScarlet is not visible with the exposure setting that was used for imaging. Scale bar, 200 μm. (F) Quantification of pals-5p::7xGFP expression in pals-20::wrmScarlet overexpressing strain and their siblings that do not overexpress pals-20 (control). Green dots in the box-and-whisker plots represent individual values for each analyzed animal; 50 animals were analyzed per strain for each of the three experimental replicates. Box lines represents median value, box bounds indicate 25 th and 75 th percentiles, and whiskers extend to the minimum and maximum values. A Kolmogorov-Smirnov test was used to calculate pvalues; **** p < 0.0001. AU = arbitrary units. (G) Fractions of animals overexpressing pals-20::wrmScarlet construct and their non-transgenic siblings that are infected with Orsay virus. Animals were scored based on the presence or absence of the Orsay virus-specific FISH probe fluorescence (six experimental replicates, at least 300 animals per replicate). Green dots represent the infection level for each replicate; the wild type was set at one for each replicate. Error bars are SD. A one sample Wilcoxon test was used to calculate the p-value; * p < 0.05. https://doi.org/10.1371/journal.ppat.1011120.g006
Strong selective pressures imposed by pathogens are likely the main drivers of the evolutionary expansion of the pals gene family in C. elegans. Such examples of species-specific immune gene family expansions are numerous in eukaryotes [17]. For example, the family of immunity-related GTPase encoding genes in mice likely expanded as a consequence of the arms race with rhoptry kinases from the protozoan parasite Toxoplasma gondii, as well as with other intracellular pathogens including microsporidia [17,[24][25][26]. Furthermore, the plant immunity R genes are expanded in plant genomes and share many characteristics in common with the C. elegans pals gene family, although they are not homologs [9]. In particular, there are similarities in gene expression regulation, protein-protein interactions and phenotypes between the pals-22/pals-25 pair and the R gene pair RPS4/RRS1 in the plant Arabidopsis thaliana [27][28][29][30]. Importantly, R genes regulate a switch between growth and defense, similar to pals genes [1][2][3]9].
The presence of antagonistic paralogs in pals and R gene families highlights the importance of maintaining the balance between immunity and development. In this study, we demonstrate the striking effect that loss of pals-17 has on animal development, which can be reversed by loss of either pals-20 or pals-16. Interestingly, mutations in pals-22 also cause developmental delay, but to a lesser degree than the loss of pals-17 [9][10][11]. Although pals-22(jy1) and pals-17 (*) alleles have premature stop codons in similar regions of pals-22 and pals-17, developmental impairment is more severe in pals-17(*) animals. Furthermore, complete loss of pals-17 causes early developmental arrest.
Other immunity-related genes have been previously implicated in organismal development. For example, overexpression of immune genes, which participate in defense against the bacterial pathogen Pseudomonas aeruginosa, have been shown to impair development of C. elegans. Specifically, hyperactivation of PMK-1 p38 MAPK pathway components in animals that are not exposed to pathogen causes significant delay in development [5,8]. Recently, gut microbiota have been shown to regulate activation of this immune pathway, affecting organismal growth on certain food sources [31]. Furthermore, a null mutation in the gene encoding pseudokinase NIPI-3, an important immunoregulator, leads to developmental arrest [6,7]. NIPI-3 is a negative regulator of bZIP transcription factor CEBP-1, which stimulates transcription of sek-1-a core component of the PMK-1 p38 MAPK immune pathway during development [5,7,32]. Loss-of-function mutations in several genes from the PMK-1 p38 MAPK pathway suppress the larval arrest phenotype of nipi-3 mutants, suggesting that immune hyperactivation has a detrimental effect on development [5]. Similarly, overexpression of a cytokine, tumor necrosis factor (TNF), causes growth impairment in mice, due to improper cartilage development and insufficient bone growth [33]. The elevated levels of different cytokines also play important roles in the intrauterine development of mammals. For example, increased levels of interferon-gamma (IFN-γ), type-I IFNs, TNF-α, and interleukins 1β, 6, and 17 lead to developmental failures in different fetal organs, including the neural tube, brain, lungs and placenta [34][35][36][37][38][39][40][41][42]. These examples demonstrate the deleterious effects that an overstimulated immune system can have on growth and reproduction.
The balance between immunity and development is also dependent on the timing of gene expression. Our data demonstrate that IPR gene upregulation in pals-17 mutants can be completely suppressed by loss of pals-16 at different developmental stages, whereas loss of pals-20 specifically suppresses this phenotype in L1 larvae but not in L4 larvae. Nevertheless, we observed that pals-20 still plays a role in immunity at later stages of development. This result suggests that IPR activation in L1 animals could be essential for priming animals for defense against pathogens that they may encounter later in life. Therefore, the lack of IPR induction in L1 pals-17 pals-20 mutants might be sufficient to cause wild-type susceptibility to intracellular pathogens later in life. Alternatively, it is possible that IPR activation by pals-20 fluctuates throughout life and may not be specific for younger developmental stages. Furthermore, our RNA-seq analyses show a drastic difference in the number of differentially expressed genes between partially synchronized pals-17 and L4 stage pals-17 pals-20 mutant animals. This distinction suggests that there may be some other pals-17-dependent genes that play a role in immunity and are still suppressed by loss of pals-20 at the L4 stage.
Altogether, in this and our previous studies, we have shown that five uninduced pals genes, pals-16, pals-17, pals-20, pals-22 and pals-25, act as ON/OFF switch modules to regulate induced pals genes and other IPR genes [9][10][11]. Our analysis indicates that the OFF switch pals-17 functions independently of the OFF switch pals-22. Further studies could address if other closely related uninduced pals genes represent additional modules that regulate the IPR and development. Having immune modules with some functional overlap likely provides an advantage for hosts defending against infection. Such modules exist among plant R genes, where the same bacterial effector AvrRps4 from Pseudomonas syringae can be recognized by two related but distinct R gene modules-RPS4/RRS1 and RPS4B/RRS1B [29]. However, in other contexts, these modules have non-overlapping function. For example, the PopP2 effector from bacterial species Ralstonia solanacearum is specifically recognized by the RPS4/RRS1 gene pair, but not the RPS4B/RRS1B gene pair [29]. While it is not known what activates pals genes in C. elegans, it is possible that different pals modules in C. elegans are responsible for sensing different pathogen effectors in order to induce the IPR. If it is true that distinct pals modules sense distinct effectors, this specificity would provide an explanation for the expansion of the pals gene family in C. elegans, which appears to have extensive control over the balance between immunity and development.
C. elegans strains were grown on Nematode Growth Media (NGM) agar plates with streptomycin-resistant Escherichia coli OP50-1 at 20˚C [43], unless stated otherwise. Strains used in this study are listed in S6 Table.
To obtain synchronized populations of C. elegans [44], gravid adult animals were washed off plates with M9 media and collected into 15 ml conical tubes. Tubes were centrifuged and the supernatant was removed leaving 3 ml of M9. 1 ml of bleaching solution (500 μl of 5.65-6% sodium hypochlorite solution and 500 μl of 2 M NaOH) was then added to the tube until adult animals were partially dissolved. M9 was added to 15 ml to wash the released embryos. The tubes were immediately centrifuged and the supernatant was removed. The washing step was repeated four times and embryos were resuspended in a final volume of 3 ml of M9. Embryos were placed in a 20˚C incubator with continual rotation for 20-24 h to hatch into L1 animals.
L4 stage pals-22(jy1) pals-25(jy11) mutants carrying the jyIs8[pals-5p::gfp myo-2p::mCherry] transgene were washed off plates with M9 media and collected into 15 ml conical tubes. Following centrifugation and supernatant removal, animals were washed with M9 two times to remove bacteria. The worm pellet was resuspended in 4 ml M9 and 25 μl of ethyl methanesulfonate (EMS) was added to the tube [45]. Animals were incubated at room temperature for 4 h with continual rotation. After incubation, the worms were pelleted and the supernatant was removed. Animals were washed with M9 two times to remove the mutagen. Worms were plated on 10 cm Petri plates with OP50-1 bacteria and incubated at room temperature for 2.5 h to recover. 18 groups of 30 mutagenized animals were then transferred to new 10 cm Petri plates with OP50-1 bacteria and incubated at 20˚C. After 36 h, mutagenized P0 animals were removed and the incubation of F1 progeny was continued. 16,752 F1 animals reached adulthood after three days and they were bleached to obtain their synchronized L1 progeny (thus, we calculate that we screened 33,504 haploid genomes). These F2 progeny were grown and analyzed for expression of pals-5p::GFP at different developmental stages, which led to the isolation of jy74 mutants. These mutants were monitored through several generations to confirm that the GFP ON phenotype was still present. jy74 animals were backcrossed to wild-type strain six times.
Genomic DNA from wild-type and mutant strains was isolated using Qiagen Puregene Core Kit A and submitted to BGI Genomics for whole genome sequencing. The Galaxy platform (https://usegalaxy.org) was used for mapping and variant calling to identify the causative mutation in the jy74 allele. The quality check of the reads was performed using FastQC. Bowtie2 program [46,47] was used to map the reads against the reference C. elegans genome WBCel235. dna.toplevel.fa.gz. Genetic variants were identified using the FreeBayes program. The VcfAlle-licPrimitives program was used to split allelic primitives into multiple VCF lines. Wild-type and mutant VCF datasets were intersected using the VCF-VCFintersect program. Variants were filtered using the filter criteria (QUAL>30)&(DP> = 10)&(isHOM(Gen[0]) in SnpSift Filter [48], and the remaining variants were annotated using the SnpEff eff program [49].
Before imaging, animals were anesthetized with 100 μM levamisole. Samples were imaged using a Zeiss AxioImager M1 compound microscope equipped with Axio Vision 4.8.2 software (Figs 1A, 1B, 1F, 1G, 1I, 2A,3A, 6E, S2B and S3A) or a Zeiss LSM700 confocal microscope run by ZEN2010 software (Figs 5 and S1 and S6).
Following 24 h or 44 h incubation at 20 ˚C, synchronized animals were collected and washed in M9, and fixed in 4% paraformaldehyde for 15 min. GFP fluorescence and time-of-flight (TOF) were measured using the COPAS Biosort machine (Union Biometrica). Fluorescence intensity was standardized to the TOF values, which are a proxy for worm length. Data analysis was performed using GraphPad Prism 9. Raw data are shown in S1 Table . For measurements of the pals-5p::7xGFP reporter expression, fixed animals were imaged using the ImageXpress automated imaging system NanoImager. Fluorescence was measured using FIJI software [50] as a mean fluorescence per animal, which was normalized to background florescence.
Synchronized animals were grown at 20 ˚C for 44 h, collected and washed in M9, and fixed in 4% paraformaldehyde for 5 min. Animals were imaged using the ImageXpress automated imaging system NanoImager (Molecular Devices, LLC) in 96-well plates. The length of each animal was measured using the FIJI program [50]. 50
RNA interference assays were performed using the feeding method [51]. Overnight cultures of HT115 E. coli were plated on RNAi plates (NGM plates supplemented with 5 mM IPTG and 1 mM carbenicillin) and incubated at room temperature for 1-3 days. Synchronized L1 or L4 animals were transferred to these plates and incubated at 20 ˚C. The effects of pals-17 RNAi treatment on animals that were plated at the L1 stage were examined after 44 h incubation (Fig 1G -1J). For pals-20 RNAi treatment, the progeny of L4 animals were examined after three days (Fig 2A). The effects of pals-16 RNAi were examined 72 h after plating L1 animals (Fig 3A and3B). Control RNAi experiments were carried out using vector-alone plasmid L4440.
We designed mutations for pals-17 alleles syb4005 (pals-17(Δ)) and syb3980 (pals-17(*)) and pals-20 allele syb3744 (pals-20(*)); CRISPR/Cas9-mediated genome editing was performed by SunyBiotech. SunyBiotech balanced pals-17(Δ) allele with sC1(s2023). pals-17(Δ) allele is a 1,394 base pair deletion and a 16 base pair insertion. The deletion starts 36 nucleotides upstream of the pals-17 start codon and ends at the 69 th nucleotide after the stop codon; the inserted sequence is 5'-AAAGATCTTTATTTAA-3'. The pals-17(*) allele contains six point mutations that change amino acids L(49) N (50) and Y(51) to three consecutive stop codons; sequence 5'-CTTAACTAC-3' is mutated to 5'-TAGTAATAA-3', creating a SpeI cutting site. pals-20(*) allele contains two point mutations in pals-20 that change amino acid E (15) to stop codon and one silent mutation in the 13 th codon encoding valine; sequence 5'-GTTATCGAA-3' is mutated to 5'-GTCATCTAG-3', creating a XbaI cutting site. The mutations in pals-20(*) should spare the function of the Y82E9BR.5 gene, which completely overlaps with pals-20. Genotyping primers for pals-17 (primers 1 and 2) and pals-20 point mutants (primers 3 and 4) are listed in S7 Table . The pals-17 allele jy167 was created in the pals-20 mutant background; the jy167 allele contains the same mutations as syb3980 and they are both labeled as pals-17(*) in the main text. The deletion alleles of pals-16 in the wild-type, pals-17, and pals-17 pals-20 mutant backgrounds are jy182, jy171 and jy174, respectively; all three alleles are labeled as pals-16(Δ) throughout the text. The jy167, jy182, jy171 and jy174 alleles were all created using the co-CRISPR method with preassembled ribonucleoproteins [52,53]. Cas9-NLS protein (27 μM final concentration) was obtained from QB3 Berkeley; crRNA, tracrRNA and DNA primers were ordered from Integrated DNA Technologies (IDT). The crRNA sequence used for generating the jy167 allele in pals-17 was ATTCAGGTTCCATGTAGTTA; the repair template sequence was 5'-TGGAGAGGCAAGGCGATTTTGCTAGACCAGAGCAGTCAAACTAG-TAATAAATGGAACCTGAATATTATGAGTTTAAGGTATATTTTATTTTGAGAATATA-TAA-3'. This repair template sequence replaces wild-type sequence 5'-CTTAACTAC-3' with mutated sequence 5'-TAGTAATAA-3' that changes amino acids L( 49) N (50) and Y(51) to three consecutive stop codons and adds a SpeI cutting site. The mutated region is flanked with a 41-nucleotide overhang on the 5' end and a 53-nucleotide overhang on the 3' end. AAACGTTTCTACACAGCATG and ACATCGTGACGCCAAATTCA crRNA sequences were used to target regions of the pals-16 gene close to the start codon and the stop codon, respectively. The jy182 allele is a 1,300 base pair deletion in pals-16 in the wild-type background, starting one nucleotide upstream of the pals-16 start codon and ending 14 nucleotides upstream of the stop codon. The jy171 allele is a 1,299 base pair deletion and seven base pair insertion in pals-16 in the pals-17(*) background. The deletion starts at the second nucleotide of the pals-16 start codon and it ends 13 nucleotides upstream of the stop codon; the inserted sequence is 5'-CGGGTGA-3'. The jy174 allele is a 1,302 base pair deletion in pals-16 in the pals-17(*) pals-20(*) background, starting immediately after the pals-16 start codon and ending seven nucleotides upstream of the stop codon. PCR genotyping was performed using primers 5-7 listed in S7 Table. Selected lines were backcrossed three times to the wild-type strain before they were used in experiments.
Gravid adult animals were grown at 20˚C and picked onto 10-cm seeded NGM plates (40 animals per strain; 1 plate per strain). Due to the low brood size of pals-17 mutants, 60 animals were picked for this strain to ensure an adequate sample size of progeny. Animals were then incubated at 20˚C for 2 h. Next, approximately 7 mL of M9 was poured onto the plate and gently swirled for 15-20 s to dislodge all adult animals while leaving eggs fastened to the E. coli OP50-1 lawn. The M9 was removed, and the plates were thoroughly checked under a microscope to ensure that no adult animals remained. After verifying that the eggs were not washed off, the plates were incubated at 20˚C and the time course began. At the 44 h, 64 h, and 72 h time points, 100 animals on each plate were scored for development as being at or below the L3 stage or at or above the L4 stage. Data analysis was performed using GraphPad Prism 9. Raw data are shown in S1 Table.
Total RNA isolation was performed as previously described [54]. For RNA isolation from L1 animals, 35,000 worms were grown per NGM plate for 3 h at 20 ˚C; one or two plates were used per strain per replicate. For RNA isolation from L2 animals, 10,000 worms were grown per NGM plate for 33 h (pals-17(*) mutants) or 24 h (all other strains) at 20 ˚C; four plates were used per strain per replicate. For RNA isolation from L4 animals, 3,000 worms were grown per NGM plate for 72 h (pals-17(*) mutants) or 48 h (all other strains) at 20 ˚C; up to four plates were used per strain per replicate. Animals were washed off plates using M9, then washed with M9 and collected in TRI reagent (Molecular Research Center, Inc.). RNA was isolated using a BCP phase separation reagent, followed by isopropanol and ethanol washes. For RNA-seq analysis, samples were additionally purified using the RNeasy Mini kit (Qiagen).
qRT-PCR analysis was performed as previously described [54]. In brief, cDNA was synthesized from total RNA using iScript cDNA synthesis kit (Bio-Rad). qRT-PCR was performed using iQ SYBR Green Supermix (Bio-Rad) with the CFX Connect Real-Time PCR Detection System (Bio-Rad). Three independent experimental replicates were performed for each qRT-PCR analysis. Each sample was analyzed in technical duplicates. All values were normalized to the expression of the snb-1 control gene, which does not change expression upon IPR activation [11]. The Pffafl method was used for data quantification [55]. The sequences of the primers used in all qRT-PCR experiments are given in S7
Amino acid sequences were obtained from https://wormbase.org/. 38 out of 39 PALS proteins were analyzed; PALS-36 protein was excluded from the analysis because pals-36 is annotated as a pseudogene and the transcript is not available. Eight PALS proteins have more than one isoform: isoform "a" transcripts were used for PALS-3, PALS-7, PALS-11, PALS-19, PALS-23, PALS-37 and PALS-38; the isoform "b" transcript was used for PALS-31. Clustal omega program (https://www.ebi.ac.uk/Tools/msa/clustalo/) was used for sequence alignment of 38 PALS protein sequences [56]. A phylogeny software PhyML 3.0 (http://www.atgc-montpellier. fr/phyml/) based on the maximum likelihood principle with the Smart Model Selection criterion [57,58] was used to create a radial phylogram shown in S4A Fig.
cDNA library preparation and paired-end sequencing were performed at the Institute for Genomic Medicine at the University of California, San Diego. RNA-seq reads were uploaded to the NCBI GEO database with accession number GSE229165 (subseries GSE229142 and GSE229164). Reads were mapped to C. elegans WS235 genome using Rsubread in RStudio. Differential expression analyses were performed using limma-voom function in the Galaxy platform (https://usegalaxy.org/). Genes with counts number lower than one count per million (CPM) were filtered out. Differentially expressed genes had adjusted p-values lower than 0.05 (S3 Table ). Overlaps between datasets shown in Venn diagrams (Fig 4A -4C) were calculated using Multiple List Comparator software (https://molbiotools.com/listcompare.php). The significance of overlaps between datasets was calculated using Overlap Stats software (http:// nemates.org/MA/progs/overlap_stats.html). The default number of genes (17,611) was used for statistical analyses.
Animals were grown at 20˚C and then single-picked onto separate 3.5 cm tissue culturetreated NGM plates upon reaching the L4 stage. Every 24 h, the number of eggs and progeny on each plate was recorded and the gravid adult animal was transferred to a new plate. This process was repeated daily until all animals laid no eggs for three days, at which point the counts were summed to calculate the total progeny for each individual. Three experimental replicates were performed. Data analysis was performed using GraphPad Prism 9. Raw data are shown in S1 Table.
Gene Set Enrichment Analysis (GSEA) software v4.2.3 was used for functional analysis of pals-17(*) and pals-17(*) pals-20(*) compared to wild type (Fig 4F ) using the GSEAPreranked module [21,22]. Differentially expressed genes from RNA-seq expression data for pals-17(*) versus wild type and pals-17(*) pals-20(*) versus wild type were ranked based on Log2 fold changes and converted into a GSEA-compatible file. 93 previously published gene sets were used for comparison [9,10,12,19,23,54,[59][60][61][62][63][64][65][66][67][68][69][70][71][72][73][74][75][76][77]. For both analyses, a signal-to-noise metric of 1,000 permutations was used with "no collapse" as gene symbols were pre-defined in the ranked gene list. Gene sets that had fewer than 15 genes represented in the RNA-seq data from this study were excluded from statistical analysis. Normalized Enrichment Scores from GSEAPreranked analysis described above were calculated with default parameters. A heatmap of the GSEA analysis was made using GraphPad Prism 9 software. Raw data are shown in S1 Table ; full datasets are shown in S5 Table.
pals-17::gfp and pals-20::wrmScarlet translational reporter constructs were assembled using NEBuilder HiFi kit (New England Biolabs) by fusing PCR products and gene fragments. The pET755 plasmid was used for Q5 polymerase-mediated (New England Biolabs) PCR amplification of the backbones that contain an ampicillin resistance cassette and unc-54 3'utr, and, in the case of pals-20 reporters, a wrmScarlet::3xHA sequence (primers 26-28 in S7 Table ). Promoter regions were PCR amplified using Q5 polymerase (New England Biolabs) from C. elegans genomic DNA using primers 29-32 listed in S7 Table. Amplified promoter regions contain 1,143 base pairs upstream of the pals-17 start codon and 1,000 base pairs upstream of the pals-20 start codon. pals-17(cDNA)::gfp::3xFlag and pals-20 cDNA sequences were synthesized as gene fragments by Twist Biosciences.
pals-17p::gfp and pals-20p::wrmScarlet transcriptional reporter constructs were assembled by PCR amplification of regions of interest from the previously created translational reporter constructs (pals-17 and pals-20 cDNA segments were excluded), which were subsequently circularized using NEBuilder HiFi kit (New England Biolabs). Primers 33-36 were used for creating pals-17 and pals-20 transcriptional reporter constructs (S7 Table ).
The resulting plasmids were verified by restriction digest and sequencing across the insert. Transcriptional reporters were individually injected at concentrations of 50 ng/μl. Translational reporters (c = 13 ng/μl for each) were co-injected with the gcy-8p::gfp neuronal marker (c = 20 ng/μl) into a wild-type genetic background. For functionality testing, pals-5p::GFP expression analysis and Orsay virus infection assay, the pals-17::gfp and pals-20::wrmScarlet construct were separately injected at a higher concentration of c = 95 ng/μl, together with a myo-2p::mCherry pharyngeal marker (c = 5 ng/μl).
The intestinal pals-17::gfp::3xFlag and pals-20::wrmScarlet::3xHA reporter plasmids were created by replacing the endogenous promoters with 1,600 base pairs of vha-6 promoter using PCR amplification (primers 37-42 in S7 Table ) and NEBuilder HiFi kit (New England Biolabs). These intestinal reporters were injected at a concentration of 100 ng/μl.
The pals-5p::7xgfp reporter (allele knuSi886) was created by InVivo Biosystems as a split-GFP system. A construct containing the following elements was generated: pals-5 promoter fused to seven copies of the 11 th segment of the gfp gene, the eft-3 promoter fused to the remaining ten segments of gfp gene, and the unc-119 rescuing cassette. MosSCI transgenesis method was used to insert this construct into the ttTi5605 site on the second chromosome in the unc-119 mutant background.
Orsay viral preps were prepared as previously described [12]. Synchronized L1 animals were exposed to a mixture of OP50-1 bacteria and Orsay virus for 18 h at 20 ˚C (Fig 6A). For infection at the L4 stage (Fig 6B ), synchronized animals were grown on rde-1 RNAi plates (1,000 animals per plate, 2 plates per strain) for 72 h (pals-17(*) mutants) or 48 h (all other strains) at 20 ˚C. rde-1 RNAi increases susceptibility to Orsay virus infection [78,79]. L4 animals were top-plated with a mixture of Orsay virus, OP50-1 and M9, and incubated at 20˚C for 24 h. Following incubation, animals were collected and washed in M9, and subsequently fixed in 4% paraformaldehyde for 30 min. Fixed worms were washed and then stained at 46 ˚C overnight using FISH probes conjugated to the red Cal Fluor 610 or green FAM fluorophore (Biosearch Technologies), targeting Orsay virus RNA1 and RNA2 [80,81]. Analyses were performed visually using a Zeiss AxioImager M1 compound microscope; at least 300 animals were scored for the presence of the FISH fluorescent signal per strain per replicate. Due to relatively high variation in infection levels between replicates, the percentage of infected wild-type animals was normalized to one. Data analysis was performed using GraphPad Prism 9 software. Normalized and raw data are shown in S1 Table.
N. parisii spores were prepared as previously described [82]. Spores (1 million per plate) were mixed with food and L1 (Fig 6C ) or L4 (Fig 6D) synchronized animals. To reach the L4 stage, worms were grown on NGM plates for 72 h (pals-17(*) mutants) or 48 h (all other strains) at 20 ˚C. During infection, animals were incubated at 25 ˚C for 30 h. Animals were collected and fixed in 4% paraformaldehyde for 30 min. Fixed worms were stained at 46 ˚C overnight using MicroB FISH probe conjugated to the red Cal Fluor 610 fluorophore (Biosearch Technologies), targeting microsporidia ribosomal RNA [81,82]. N. parisii pathogen load was measured with the COPAS Biosort machine (Union Biometrica). Data analysis was performed using GraphPad Prism 9. Raw data are shown in S1 Table.
1,200 synchronized L1 worms were mixed with 6 μl fluorescent beads (Fluoresbrite Polychromatic Red Microspheres, Polysciences Inc.), 25 μl 10X concentrated OP50-1 E. coli and M9 (total volume 300 ul) [10,19,54]. This mixture was plated on 6 cm NGM plates, allowed to dry for 5 min and then incubated at 25 ˚C. After 5 min incubation, plates were transferred to ice. Animals were washed off plates using ice-cold PBST and fixed in 4% paraformaldehyde for 30 min. Samples were washed with PBST and animals were imaged using the ImageXpress Nano Automated Imaging System (Molecular Devices, LLC). Fluorescence was analyzed in FIJI program; red fluorescence was analyzed in S8A
PLOS Pathogens | https://doi.org/10.1371/journal.ppat.1011120July 18, 2023