Gene-environment and protein-degradation signatures characterize genomic and phenotypic diversity in wild Caenorhabditis elegans populations

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Abstract

Background

Analyzing and understanding the relationship between genotypes and phenotypes is at the heart of genetics. Research on the nematode Caenorhabditis elegans has been instrumental for unraveling genotype-phenotype relations, and has important implications for understanding the biology of mammals, but almost all studies, including forward and reverse genetic screens, are limited by investigations in only one canonical genotype. This hampers the detection and functional analysis of allelic variants, which play a key role in controlling many complex traits. It is therefore essential to explore the full potential of the natural genetic variation and evolutionary context of the genotype-phenotype map in wild C. elegans populations.

Results

We used multiple wild C. elegans populations freshly isolated from local sites to investigate gene sequence polymorphisms and a multitude of phenotypes including the transcriptome, fitness, and behavioral traits. The genotype, transcriptome, and a number of fitness traits showed a direct link with the original site of the strains. The separation between the isolation sites was prevalent on all chromosomes, but chromosome V was the largest contributor to this variation. These results were supported by a differential food preference of the wild isolates for naturally co-existing bacterial species. Comparing polymorphic genes between the populations with a set of genes extracted from 19 different studies on gene expression in C. elegans exposed to biotic and abiotic factors, such as bacteria, osmotic pressure, and temperature, revealed a significant enrichment for genes involved in gene-environment interactions and protein degradation.

Conclusions

We found that wild C. elegans populations are characterized by gene-environment signatures, and we have unlocked a wealth of genotype-phenotype relations for the first time. Studying natural isolates provides a treasure trove of evidence compared with that unearthed by the current research in C. elegans, which covers only a diminutive part of the myriad of genotype-phenotype relations that are present in the wild.

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Recombinational Landscape and Population Genomics of Caenorhabditis elegans

Matthew Rockman, Leonid Kruglyak (2009)

Introduction The allelic variants that underlie heritable phenotypic variation are distributed along chromosomes. Their distribution is shaped by the machinery of meiosis within individuals and by mutation, selection, and drift among them. To discover the genetic basis of complex traits, and to understand the evolutionary dynamics that shape this genetic architecture, we must characterize empirical patterns of linkage and linkage disequilibrium. We have undertaken this task in the nematode C. elegans. Mapping of thousands of mutants to the genome and molecular studies of meiotic machinery have provided a view of the large-scale landscape of the C. elegans recombination map. The chromosomes exhibit nearly complete crossover interference [1], such that each chromosome experiences one crossover per meiosis and has a genetic length of 50 cM [2]. Accumulated data from thousands of two- and three-point mapping crosses and small-scale SNP-based analyses have demonstrated a general pattern of large, nearly constant-rate domains on the autosomes, with high recombination in chromosome arms and low recombination in chromosome centers. Despite strong global regulation of crossover number, many details remain unclear, including the locations of the domain boundaries, the occurrence of fine-scale variation within domains, and the existence of domain structure on the X chromosome. Moreover, evidence for the genetic control of crossover number and position [1]–[4] leaves open the possibility that segregating variants may influence recombination patterns in experimental crosses of natural isolates. Because recombination patterns have been studied only on broad scales in individual crosses, involving fewer than two dozen markers per chromosome, dense characterization of a massive cross promises to clarify the recombinational landscape. C. elegans is one of the most exhaustively studied of all species with respect to developmental, behavioral, and physiological genomics, but studies of its population biology have lagged. Although natural genetic variation has been a source of alleles for genetic analysis in C. elegans since long before the system became a model [5], the widely accepted notion that worms exhibit little variation has discouraged investigations of their diversity. The difficulty of collecting C. elegans from the wild has compounded the problem. Nevertheless, recent work has revealed abundant heritable phenotypic variation among wild C. elegans strains [6]–[20] and has begun to reveal the ecological context for this species [16], [17], [21]–[25]. C. elegans geneticists have exploited this variation to map quantitative trait loci [26]–[37], and in a handful of cases to identify the causal mutations underlying phenotypic variation (in genes npr-1, mab-23, tra-3, zeel-1, plg-1, and scd-2 [10], [30], [38]–[43]). In parallel, studies of variation at molecular markers have begun to provide an account of the distribution of genetic variation within and among localities and across genomic regions [6], [7], [23], [24], [40], [41], [43]–[60]. These studies have shown that the species exhibits substantially lower levels of polymorphism and higher levels of linkage disequilibrium than other model systems, even those, like Arabidopsis thaliana, that share with C. elegans a primarily selfing mating system. The empirical pattern of linkage disequilibrium may result as much from selection against recombinant genotypes as from attributes of population biology such as population size and outcrossing rate [24],[61]. A genome-wide assessment of linkage disequilibrium is required to determine whether natural isolates of C. elegans will be useful for mapping loci by association. We generated and genetically characterized a recombinant inbred advanced intercross population to gain insights into the recombination map in C. elegans, and we characterized a large panel of wild strains to characterize linkage disequilibrium. The data on recombination in the lab and in the wild reveal the role of population genomic processes in shaping genotypic diversity in C. elegans, and they lay the groundwork for rapid discovery of the genes underlying phenotypic variation. Results Patterns of Recombination in Recombinant Inbred Advanced Intercross Lines We genotyped 1454 nuclear SNP markers in 236 recombinant inbred advanced intercross lines (RIAILs). These lines represent the terminal generation of a 20-generation pedigree founded by reciprocal crosses between the laboratory wild type strain N2 (Bristol) and the Hawaiian isolate CB4856. The pedigree includes ten generations of intercrossing (random pair mating with equal contributions of each pair to succeeding generations [62]) followed by 10 generations of selfing. The SNP markers span 98.6% of the physical length of the chromosomes (Table S1). The median spacing is 61,160 bp, and 80% of intervals are shorter than 100 kb. Only 35 marker intervals (2.4%) are greater than 200 kb. The RIAILs contain 3,629 breakpoints in 772 marker intervals; some breakpoints may be identical by descent because of the shared ancestry during the intercrossing phase of RIAIL construction. An estimate of the mapping resolution of the panel, based on the distances between intervals containing breakpoints, yields a median bin size of 98 kb. Because larger bins contain more of the genome than smaller bins, the expected size of a bin in which a uniformly distributed QTL will fall is 225 kb. The RIAILs exhibit a genetic map length of 1588 cM, a 5.3-fold expansion of the 300 cM F2 genetic map. The realized expansion is 93% of the expected 5.7-fold map expansion, a difference attributable, at least in part, to the action of selection during the construction of the lines. Although selection and drift may alter the relationship between recombination fraction and meiotic recombination rate [63],[64], the observed recombination fractions are qualitatively informative about global patterns of recombination rate variation across C. elegans chromosomes. The genetic maps for the six C. elegans chromosomes are similar to one another and exhibit five distinct domains: two tips with effectively zero recombination, two high recombination arms, and a low recombination center, consistent with the pattern observed in classical two- and three-point mapping crosses [65]. These domains are evident in Marey maps [66], which show genetic position as a function of physical position (Figure 1; Table 1). As the recombination rate within each domain is relatively constant, we used a segmented linear regression to identify the boundaries between the domains. 10.1371/journal.pgen.1000419.g001 Figure 1 Recombination rate domains. Marey maps for each chromosome show genetic position of each marker (black points) as a function of physical position. Genetic position is measured in centiMorgans as defined on the recombinant inbred advanced intercross line population; these are not meiotic distances. Gray lines show the fits of segmented linear regressions, which estimate the boundaries of the recombination domains and their relative recombination rates. The shaded boxes above each plot show the genetically defined positions of the pairing centers [69]. 10.1371/journal.pgen.1000419.t001 Table 1 Chromosomal Domains. Chr left tip left arm center right arm right tip I Size (kb) 527 3331 7182 3835 197 Size (%) 3.5 22.1 47.7 25.4 1.3 Right end (kb) 527 3,858 11,040 14,875 15,072 Ratea (cM/Mb) 0 3.43 1.34 6.78 0 II Size (kb) 306 4573 7141 2589 670 Size (%) 2.0 29.9 46.7 16.9 4.4 Right end (kb) 306 4,879 12,020 14,609 15,279 Ratea (cM/Mb) 0 4.92 1.33 8.47 0 III Size (kb) 494 3228 6618 2877 567 Size (%) 3.6 23.4 48.0 20.9 4.1 Right end (kb) 494 3,722 10,340 13,217 13,784 Ratea (cM/Mb) 0 7.83 1.17 7.24 0 IV Size (kb) 720 3176 9074 3742 782 Size (%) 4.1 18.2 51.9 21.4 4.5 Right end (kb) 720 3,896 12,970 16,712 17,494 Ratea (cM/Mb) 0 7.65 1.05 3.64 0 V Size (kb) 643 5254 10653 3787 583 Size (%) 3.1 25.1 50.9 18.1 2.8 Right end (kb) 643 5,897 16,550 20,337 20,920 Ratea (cM/Mb) 0 3.22 1.32 5.47 0 X Size (kb) 572 5565 6343 3937 1302 Size (%) 3.2 31.4 35.8 22.2 7.3 Right end (kb) 572 6,137 12,480 16,417 17,719 Ratea (cM/Mb) 0 3.81 1.70 5.14 0 ALL Size (kb) 3262 25127 47011 20767 4101 a Rates are derived from the slopes of the segmented linear fits, scaled to yield a total genetic length of 50 cM for each chromosome. The central domain of each autosome occupies roughly half the chromosome's length, despite the very different lengths of the chromosomes (Table 1). For example, the center of chromosome V is 10.7 Mb, 51% of the chromosome length, while the center of chromosome III is 6.6 Mb, 48% of that chromosome's length. Because all the centers have very similar rates of recombination per base pair (Table 1), their different physical lengths mean that the amount of recombination in each center (its genetic length) varies with total chromosome length. The constraint of one breakpoint per chromosome then requires that the amount of recombination in the arms of each chromosome varies inversely with chromosome length; shorter chromosomes have a larger fraction of their recombination events in their arms, and the physical sizes of the arms explain much of the variation among arms in recombination rates (r2 = 0.51, p = 0.009). Nevertheless, the arms are heterogeneous in relative and absolute length and recombination rate, and the central domains are not perfectly centered on the chromosomes, consistent with the finding of Barnes et al. [65]. Most notably, the left arm of chromosome IV has a relative recombination rate more than twice that of the right arm, though they differ in size by only 15% (Figure 1; Table 1). Inspection of the Marey maps suggests that there may be additional rate variation within the defined domains. To determine whether such variation is expected in the case of constant-rate domains, we simulated chromosomes along the RIAIL pedigree with discrete, constant-rate recombination domains, and we recorded the simulated genotypes at the same marker intervals as our actual genotype data. The simulated chromosomes exhibit patterns of variation within the discrete rate domains qualitatively similar to the observed data, preventing us from placing confidence in the fine-scale patterns in the data (Figure 2A). Nevertheless, the fine-scale variation observed in our data is largely concordant with that present in genetic maps derived from independent two- and three-point mapping crosses with classic visible markers (Figure S1), compiled in WormBase [67]. The general concordance between our map, derived from meioses at 25°C, and the WormBase map, which comes from crosses performed at various temperatures but primarily at 20°C, does not support the notion that the distribution of crossovers is strongly temperature dependent [68]. 10.1371/journal.pgen.1000419.g002 Figure 2 Simulated chromosomes. (A) The Marey maps for actual chromosome III data (black) and 10 chromosome III datasets simulated with discrete, constant-rate recombination domains (colors) show that variation within domains and indistinct boundaries between domains are expected. (B) The observed genetic length of chromosome III is smaller than expected. The histogram shows the lengths of 1000 chromosome III datasets simulated assuming one crossover per meiosis. In our data, each chromosome has one very sharp center-arm boundary and one that is less sharp, and boundaries exhibit the identical pattern in the classical maps. In five of the six chromosomes, the less-sharp boundary is on the side of the chromosome that holds the pairing center [69] (Figure 1). The exception is chromosome III. We find two points of disagreement between our results and previous discussion of recombination maps in C. elegans. First, the X chromosome clearly possesses domain structure similar to that of the autosomes (Figure 1), contrary to inferences from sparser data. The major distinguishing feature of the X-chromosome center is its relative size, 36% of the chromosome length, which is substantially less than the 47–52% on the autosomes. Second, we find that the chromosome tips have extremely low recombination rates; the terminal domain of each chromosome end is a region of effectively zero recombination, a pattern observed previously only for the right tip of the X [65] and more recently for chromosome III [68]. Every chromosome terminus contained a series of nonrecombining markers, and these domains ranged in size from 200 kb (IR) to 1300 kb (XR), averaging 600 kb. Selection We previously showed that the allele frequencies in the RIAILs depart from the neutral expectation, implicating selection during the application of the cross design [40].We extend that analysis here, estimating expected allele frequency skew using our simulations that explicitly incorporate marker spacing and recombination domain structure. Chromosome I (p 0.5; fewer than 5% of the 236 RIAILs had confidence scores 0.35. For the 285 SNPs that yielded some confidence scores between 0.35 and 0.5, fluorescence intensities were individually inspected and calls assigned manually when unambiguous. For many of the 1205 RIAIL-confirmed SNPs, one or more wild isolates failed to give any genotyping signal. We identified a threshold of normalized intensities of both fluors ≤0.009 at which 768 wild isolate genotypes gave no signal (0.5018% of all calls) while the RIAILs gave only 8 genotypes at the same level (0.0028%), a 180-fold enrichment for the wild isolates. As these failed wild isolate genotypes exhibit linkage disequilibrium with well-genotyped SNPs, they likely represent mutations that disrupt the hybridization of the Illumina oligos to the genotyping interval. We assigned a third-allele call to these genotypes. The remaining 331 SNP assays were individually examined to assign genotype calls. For 46 assays, N2 and CB4856 yielded the same genotype, implicating false-positive SNPs predictions. An additional 29 SNPs produced uninterpretable fluorescence intensity scatterplots. We were able to assign genotype calls for 196 SNPs which failed to pass the confidence threshold due primarily to low intensity. The remaining 70 SNPs exhibited more than two clusters of genotypes in plots of fluorescence intensities. We found that the extra clusters were due to hybridization of the SNP-assay oligos to additional loci which themselves exhibited segregation. As a result, each cluster could be assigned a homozygous genotype call on the basis of linkage disequilibrium with adjacent SNPs among the RIAILs. The final dataset included 1460 SNPs. We excluded one RIAIL from subsequent analysis because its genotypes included a large proportion of ambiguous calls. The resulting dataset includes 236 RIAILs and 125 wild isolates scored at 1460 SNPs. The 527,061 genotypes include 1450 third allele (putative deletion) calls among the wild isolates, 654 Ns for bad data, and 180 heterozygote calls. Eight of the RIAILs exhibited short tracts of residual heterozygosity. The mitochondrial genotype for each RIAIL was determined by PCR-RFLP, using primers 5′-ctcggcaatttatcgcttgt and 5′- cttactcccctttgggcaat and digesting with PmeI. We estimated a genetic map for the RIAIL cross using r/qtl [74] and found that 6 SNPs had expected physical positions on chromosomes other than those to which they mapped. These may represent errors in the genome assembly or in oligo production; the oligo sequences map uniquely in the genome assembly. The expected and mapped physical positions of these SNPs are in Table S4. Analyses of RIAILs employed the 1454 physically mapped SNPs; the complete dataset is provided in Table S1. We considered the mismapped SNPs in analyses of WI haplotypes but excluded them from analyses that required physical positions. The complete wild isolate dataset is provided in Table S2. In all cases where a RIAIL genotype contained an allele from one strain flanked by alleles from the other parental strain (i.e., a single-marker segment), we re-examined the plots of fluorescence intensities to confirm the genotype call; such a pattern is expected for a genotyping error and can strongly bias estimates of map lengths and breakpoint counts [71]. We estimate bin size as the distance from the end of a chromosome to the midpoint of the first breakpoint-containing interval or as the distance between the midpoints of successive breakpoint-containing intervals. This approach ignores bins created by multiple independent breakpoints within a single interval and uses interval midpoints rather than outside markers to avoid overlapping bins. Expected bin size is the per-base-pair sum of the squares of the bin lengths [106]. Recombination Rate Domain Analysis We estimated genetic distances in r/qtl using the Haldane map function, treating observed recombination fractions as though they had been observed in a backcross. The marker density is sufficiently high that the exact form of map function employed has little effect on estimated genetic distances. We defined the tip domains of each chromosome to include all markers between the chromosome ends and the first recombination breakpoint observed in the RIAILs. The midpoint of this most distal recombinant interval was chosen as the tip-arm domain boundary. The non-tip markers were included in a segmented linear regression analysis, using the segmented package in R [107], to identify arm-center domain boundaries. To estimate confidence intervals for the domain boundaries, we used simulations of the RIAIL chromosomes. We simulated 1000 RIAIL populations for each chromosome, using the known pedigree. Each gamete received a meiotic chromosome with 0 or 1 breakpoints (i.e., complete interference [4]), the position of the breakpoints determined by the relative recombination fractions of the centers and arms estimated from the RIAILs. The tips were specified to be non-recombining and the two arms of each chromosome were assigned equal recombination probabilities per base pair; that is, intra-chromosomal differences in rate between arms were not modeled. Each chromosome was simulated as a sequence of markers with one marker for every kilobase of chromosome. We then sampled markers at spacing defined by the genotyped SNPs, yielding a dataset of RIAIL chromosomes simulated with discrete, constant-rate recombination domains. We estimated domain boundaries for the simulated chromosomes by segmented linear regression. The 95% confidence intervals vary in size depending on the size of the chromosome and the difference in recombination probability between adjacent domains. On average the intervals span 1.1 Mb. The simulated RIAIL chromosomes were also used to estimate expected allele frequency skews and expected genetic lengths for each of the chromosomes. The RIAIL allele frequencies at each marker were estimated using the sim.geno function in r/qtl [74] to infer missing data. WormBase [67] genetic maps are derived from data available on June 7, 2008, for 4542 genes with experimentally determined map positions and known physical positions. As our analyses of these data are qualitative, we made no effort to screen these data for quality, as evident from several obviously mismapped data points in Figure S1. Breakpoint Count QTL Analysis We performed non-parametric interval mapping [76] in r/qtl [74]. The RIAILs differ in their relatedness as a result of the derivation of two selfing lines from each 10th generation intercross hermaphrodite. The paired lines exhibit substantially higher similarity (mean percent bases shared ±standard deviation, 69.6±11.4%) than unpaired lines (52.8±9.5%), so that background similarity could inflate lod scores at markers unlinked to QTLs. Moreover, the significance of the lod scores would be overestimated by conventional permutation, because the RIAILs are not exchangeable; permuted datasets would break the associations between genetically and phenotypically similar RIAILs [75],[108]. Note that the mean similarity among unpaired lines is greater than the expected 50% because of the influence of selection on allele frequencies during RIAIL construction. For this reason we have not used simulated genotypes [108] to assess QTL significance. Instead we used a structured analysis and structured permutations. We split the dataset into two subsets with each RIAIL pair split between the two. We performed linkage scans separately for the two subsets and summed the lod scores. We permuted the two subsets separately 1000 times to derive genome-wide significance estimates for each phenotype. Structure Analysis Estimation of population structure used a dataset of 40 haplotypes (haplotype 21, which differs from haplotype 20 only by a single putative deletion allele, was excluded, as the analysis treats these genotypes as missing data) and 1454 SNPs. We ran structure 2.2 [80] ten times at each of five values of K, the number of ancestral populations. We used the linkage model [79] with a burn-in period of 10,000 replicates followed by 50,000 replicates to collect estimated parameters and likelihoods. The outputs of the repeated runs at each K were aligned using CLUMPP 1.1.1 [109] and Figure 8 generated using distruct 1.1 [110]. Linkage Disequilibrium We computed lower bounds on Rmin for each chromosome using HapBound and upper bounds using SHRUB [81]. We used a dataset with 1318 SNPs, after excluding all sites with missing data or putative deletion alleles. We used Haploview 4.0 [111] to calculate r2 between all pairs of the 1042 sites with minor allele frequencies greater than 0.1 in the 40-haplotype dataset. We used these r2 values to estimate ρ per basepair and its standard error by nonlinear regression using equation 3 of Weir and Hill [112], implemented with the R function nls. This simple method of moments estimator roughly approximates a likelihood estimator. Estimates of the half-length of LD represent the distance at which the expected value of r2 from the nonlinear regression drops below half its initial value. To estimate ρ in sliding windows, we used the r2 values among SNPs within 1 Mb to either side of each focal SNP. These 2 Mb windows are the smallest practicable windows given our marker density. We also estimated ρ for whole arms and centers, using the domain boundaries estimated from the RIAILs and shown in Table 1. We estimated the distribution of r2 among nonsyntenic sites in the absence of association from 100 permutations of chromosomes among the 40 wild isolate haplotypes, preserving allele frequencies and chromosomal haplotype frequencies but breaking correlations among chromosomes. The means of the ranked nonsyntenic r2 values across permutations provides an estimate of the number of false discoveries at each quantile of the r2 distribution. Permutations and calculations were performed in R, and r2 was calculated using the LDmat function in the popgen library (http://www.stats.ox.ac.uk/˜marchini/software.html). The dataset included 784 sites with no missing data and minor allele frequencies greater than 0.1. Association Mapping We excluded singleton SNPs and those with missing data and used the resulting 40×907 matrix to estimate an identity-by-state kinship matrix using EMMA [88]. We did not remove SNPs in perfect linkage disequilibrium with other SNPs because we sought to discern the genomic extent of intervals associated with traits. We estimated the significance of associations in the mixed-model analysis using likelihood ratio tests with the function emma.ML.LRT, incorporating the kinship matrix and in some cases the ancestral population admixture assignments from structure (K = 3) as fixed effects. Supporting Information Figure S1 RIAIL maps recapitulate classical marker mapping results. Chromosomal and regional rate variation patterns observed in the recombinant inbred advanced intercross lines (black points) are similar to those observed from thousands of two- and three-point mapping experiments reported in WormBase (red points). The RIAIL map distances represented here are scaled to yield 50 cM total lengths for each chromosome. The classical mapping data corroborate the great difference in rate between the left and right arms of chromosome IV, with an exceptionally high rate on IVL between roughly 1.0 and 2.4 Mb. At a sub-arm scale, we see corroboration for variation along IIL very clearly and IR, VR, and XL less so. Other regions that show variation in the WormBase map are not evident in the RIAIL map, notably IVR, VL, and XR. Nevertheless, our results support the claim of Barnes et al. [65] that the arms are not truly constant-rate regions. (3.99 MB EPS) Click here for additional data file. Figure S2 Pairwise identity among wild isolate haplotypes. For each chromosome, pairwise allele-sharing between each haplotype is plotted below the diagonal. Above the diagonal we present results of the same analysis excluding all singleton SNPs, all of which are unique to CB4856 (haplotype 41). (1.65 MB EPS) Click here for additional data file. Figure S3 Linkage disequilibrium within chromosomes. Pairwise r 2 values for all sites with minor allele frequencies >0.1 are plotted. The axes represent physical position along each chromosome. Pairs of sites with r 2>0.5 are in black and those with r 2>0.9 are red. (0.06 MB PDF) Click here for additional data file. Figure S4 Decay of linkage disequilibrium. Each point plots r 2 for a pair of sites with minor allele frequencies >0.1, colored by chromosome, as a function of the physical distance between the two sites. The curves plot the nonlinear regression of r 2 on distance using the sample-size-corrected relationship between the variables from Weir and Hill [112]. (0.19 MB PDF) Click here for additional data file. Figure S5 Distributions of p-values for tests of association. The calculated p-value for each SNP marker is plotted under three tests of association as in Figure 10: Fisher's exact test, mixed-model likelihood ratio tests incorporating a genotypic similarity (IBS) matrix, and mixed-model LRT incorporating both genotypic similarity and the results of structure analysis. The straight line represents the expectation for uniformly distributed p-values. Without mixed-model control for genomic similarity, the p-value distribution is profoundly skewed to low values. (4.04 MB PDF) Click here for additional data file. Table S1 SNPs and RIAIL Genotypes. SNP details and genotype data for 236 recombinant inbred advanced intercross lines. (0.94 MB TXT) Click here for additional data file. Table S2 SNPs and Wild Isolate Genotypes. SNP details and genotype data for 125 wild isolates. (0.62 MB TXT) Click here for additional data file. Table S3 Strains and their Haplotypes. Strain, haplotype number, locality, and counts of genotype calls. (0.03 MB XLS) Click here for additional data file. Table S4 Misplaced SNP markers. Illumina oligo sequences, expected positions, and map-based positions. (0.02 MB XLS) Click here for additional data file.

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Chromosome-scale selective sweeps shape Caenorhabditis elegans genomic diversity

Erik C. Andersen, Justin Gerke, Joshua Shapiro … (2012)

The nematode Caenorhabditis elegans is central to research in molecular, cell, and developmental biology, but nearly all of this research has been conducted on a single strain. Comparatively little is known about the population genomic and evolutionary history of this species. We characterized C. elegans genetic variation by high-throughput selective sequencing of a worldwide collection of 200 wild strains, identifying 41,188 single nucleotide polymorphisms. Unexpectedly, C. elegans genome variation is dominated by a set of commonly shared haplotypes on four of the six chromosomes, each spanning many megabases. Population-genetic modeling shows that this pattern was generated by chromosome-scale selective sweeps that have reduced variation worldwide; at least one of these sweeps likely occurred in the past few hundred years. These sweeps, which we hypothesize to be a result of human activity, have dramatically reshaped the global C. elegans population in the recent past.

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Distinct Pathogenesis and Host Responses during Infection of C. elegans by P. aeruginosa and S. aureus

Javier E. Irazoqui, Emily Troemel, Rhonda L. Feinbaum … (2010)

Introduction The study of host-microbe interactions seeks to understand the symbiotic relationships between hosts and microbiota, and their perversion during infectious disease. Essential steps are the identification of bacterial virulence mechanisms and of host defense pathways. In mammalian hosts, Nod-like receptors (NLRs), Toll-like receptors (TLRs), and NF-κB play important roles in the intestinal epithelium, a critical interface of contact between host and microbiota [1], [2], [3]. However, how these signaling pathways function in the context of the whole organism is poorly understood, and potentially novel pathways may yet be uncovered. Likewise, the critical initial stages of infection, before the onset of overt pathogenesis, are poorly defined. Genetically tractable invertebrate model systems have aided efforts to identify evolutionarily conserved components of the innate immune system [4]. For example, studies using Drosophila melanogaster showed the central importance of the Toll and IMD signaling pathways for the regulation of Relish-family (NF-κB) transcription factors [3], [5]. Likewise, studies using Caenorhabditis elegans revealed the involvement of evolutionarily conserved p38 MAPK, insulin, TGF-β, and β-catenin signaling pathways [6], [7], [8]. In addition to being a genetically tractable model system, C. elegans is a transparent bacterivore, which allows the direct, real-time observation of infection and gene expression reporters in vivo. These qualities make it a useful model host for the study of infection and host defense in the context of the whole organism [9]. C. elegans is particularly useful for studying intestinal epithelial innate defenses, because it has only 20 such cells that are not shed (as are mammalian intestinal epithelia) and are non-renewable [9], [10], allowing the study of defense functions in vivo without potentially confounding cell proliferation and tissue repair. Furthermore, the unique biology of C. elegans allows researchers to focus entirely on epithelial innate defense because it lacks a circulatory system, macrophage-like professional immune phagocytes, and antibody-based adaptive immunity [11]. On the bacterial side, it is important to elucidate the virulence mechanisms that defeat host defenses and establish infection. Pathogenic bacteria are thought to have experienced stepwise additions of virulence factors, as they evolved to survive different host antimicrobial responses and to colonize new niches [12]. Our studies using C. elegans as a model host may thus interrogate early steps in the evolution of bacteria as pathogens, and their interactions with prototypical metazoan epithelial cells. Here we focus on two paradigmatic human pathogenic bacteria of great medical importance that represent two broad categories of evolutionarily distant microbes, the Gram-negative Pseudomonas aeruginosa and Gram-positive Staphylococcus aureus. P. aeruginosa causes systemic acute infections in patients with weakened immune systems [13] and establishes chronic infections in the lungs of cystic fibrosis patients [14]. P. aeruginosa can also infect a wide variety of plants, metazoans, and single-celled eukaryotes [15]. S. aureus is a Gram-positive bacterium that can cause severe diseases in many animal species [16], [17]. In recent years, patients lacking classical risk factors have suffered increasing rates of infection by virulent antibiotic-resistant strains [18]. Human colonization by S. aureus is widespread: 30% of the population carries S. aureus in the microflora of epithelia in the nasopharynx, skin, and intestine [19]. S. aureus can cause severe skin infections, osteomyelitis, endocarditis, food poisoning, pneumonia, and flesh-eating disease [20], for which it deploys an impressive armamentarium of virulence factors, including cytolysins that cause the destruction of host immune cells and tissues [21], [22]. Despite great progress in their identification, the exact contribution of each virulence strategy to disease in vivo is poorly understood. The genetic makeup of the host is suspected to determine susceptibility to infection, but the genetic determinants of susceptibility are unknown [20]. Mice lacking adaptive immunity survive intravenous S. aureus infection as well as wild-type animals, suggesting that innate immunity is the main clearing mechanism for S. aureus infection in mammals, but the exact mechanism is unclear [23]. New approaches are needed to understand the molecular basis of innate host defenses against P. aeruginosa and S. aureus infection. To this end, our laboratory has developed C. elegans-P. aeruginosa [24], [25] and C. elegans-S. aureus [17], [26] model systems to facilitate the study of the role of innate host defenses in conferring resistance to bacterial infections and to identify host signaling pathways relevant to defense [7], [27], [28], [29]. These infection models recapitulate key aspects of P. aeruginosa or S. aureus disease in mammals (see below), including the requirement of virulence factors necessary for mammalian infection, and have been used to identify novel P. aeruginosa and S. aureus virulence factors [17], [30], [31], [32]. Despite great progress in the dissection of C. elegans host defense signaling pathways since the initial description of the system in 1999 [9], [24], [33], [34], little information has been available on the cellular basis of bacterial pathogenesis and nematode killing. In this study, we focused on the interactions between C. elegans intestinal cells —as prototypical metazoan epithelial cells, and as the first line of defense against intestinal infection—and P. aeruginosa or S. aureus. We investigated the cytopathologies that occur during infection, which suggest distinct mechanisms of virulence used by each bacterial species in vivo. With P. aeruginosa, we found that initial intestinal distention, putative outer membrane vesicle (OMV) production, and extracellular matrix accumulation on the intestinal cell brush border are followed by host autophagic abnormalities, intracellular invasion, and penetration of the epithelial barrier. Similarly, previous studies found that P. aeruginosa forms biofilms in the lungs of infected patients, where OMV production is also evident [35], [36]. In contrast, faster accumulation of S. aureus in the C. elegans intestine resulted in enterocyte effacement and loss of intestinal cell volume, followed by intestinal epithelial cell lysis and bacterial invasion of the rest of the body, with complete degradation of host internal tissues. Likewise, previous studies showed intestinal colonization by S. aureus, enterocyte effacement in rabbits and neonates, and toxin-mediated cell lysis both in vitro and in vivo [37], [38], [39], [40], [41], [42], [43]. We also evaluated the differential impact of these distinct pathogenic processes on host gene transcription. We previously defined the host transcriptional response to P. aeruginosa infection [44]. To understand if and how the host responds to different virulence mechanisms by employing distinct transcriptional host responses, here we defined the host response to S. aureus. The two responses show minimal overlap; the response to S. aureus apparently involves host damage- and TLR-independent recognition of microbial molecules, potentially pathogen-associated molecular patterns (PAMPs), whereas C. elegans may sense P. aeruginosa-derived heat-labile signals or pathogen-elicited damage. Using functional genomics, we identified host factors critical for host defense against S. aureus, some of which are analogous to human innate defense factors. These observations advance our knowledge of bacterial pathogenesis in C. elegans, and show that the C. elegans infection model illuminates evolutionarily conserved mechanisms of bacterial pathogenesis and epithelial host defense. Results P. aeruginosa infection: intestinal distention, extracellular material accumulation, intracellular invasion, outer membrane vesicles, and abnormal autophagy To determine the cytopathology of P. aeruginosa colonization of the C. elegans intestine, we used transmission electron microscopy (TEM) of the intestinal epithelium (Figure 1A) to evaluate signs of pathogenesis at early times (8 h) or at later times (24 h and 48 h) of infection. At 8 h of infection, we found gross intestinal distention but little bacterial accumulation. Instead, we observed unidentified electron-dense extracellular material accumulating on the apical surface of the brush border (Figure 1C). In addition to coating the brush border, the electron-dense material surrounded bacterial cells that appeared intact and formed clumps in the intestinal lumen. Also surrounding the bacteria and in contact with the extracellular material, we found abundant accumulation of putative outer membrane vesicles (OMVs) (Figure S1A, B). P. aeruginosa OMVs have been shown to act as a virulence factor and toxin delivery mechanism [45]. We did not observe intestinal distention, OMVs, or matrix accumulation in E. coli-fed control animals (Figure 1B). 10.1371/journal.ppat.1000982.g001 Figure 1 P. aeruginosa causes intestinal distention, extracellular material accumulation, intracellular invasion, and abnormal autophagy. A. Schematic representation of C. elegans body plan and plane of section (Left) and midbody transversal section indicating major organs (Right). Square highlights area of focus in TEM micrographs. B–I. TEM micrographs of transversal midbody sections of animals feeding on non-pathogenic E. coli OP50(B, D), virulent P. aeruginosa PA14(C, E, G, H, I), or attenuated gacA P. aeruginosa PA14 (F) for 8 h (B, C), 24 h (D, E), or 48 h (F–I). Scale bar in B–F, 2 µm; in G–I, 0.5 µm. iec, intestinal epithelial cell; mv, microvilli; b, bacterial cell; bwm, body wall muscle; aj, apical junctions; em, extracellular material; au, arrested autophagosomes, ER, expanded endoplasmic reticulum. At 24 h of infection, the intestine became further distended, with noticeably more bacterial cells accumulating in the lumen in the form of clumps of cells surrounded by extracellular matrix (Figure 1E). There was a thick layer of matrix material coating the microvilli, which were present and of approximately normal length (i.e., ∼1 µm). In contrast, E. coli-fed animals lacked these signs of pathogenesis, exhibiting non-distended intestinal lumina and intestinal epithelial cells filled with lipid droplets and other gut granules characteristic of healthy animals (Figure 1D). At 48 h of infection, pathogenesis advanced further, resulting in higher levels of bacterial accumulation in the grossly distended intestinal lumen (Figure 1G). The bacterial cells were mostly not in direct contact with the microvillar surface, but separated from it by a thick layer of extracellular material. At this time, there was widespread shortening of the microvilli and intracellular invasion by the bacteria (Figure 1G). Intracellular invasion was observed in 21% of cross sections (N = 14), only after 48 h infection. In some cases, we found bacterial cells at distal sites beyond the intestine, suggesting that P. aeruginosa can penetrate the intestinal cells and invade other tissues (Figure S1C). In addition to these phenotypes, we observed an increased number of autophagosomes, readily identifiable by their multi-membranous structure (Figure 1H and S2). Indeed, most autophagosomes appeared to be either early autophagosomes (Figure 1H) or aberrant multivesicular autophagosomes (Figure 1I). In contrast, mutant gacA P. aeruginosa, lacking the master virulence regulator GacA and therefore attenuated for C. elegans killing [25], caused much lower levels of autophagosome accumulation (Figure S2), pathogenesis (Figure 1F), and OMVs (Figure S1D) by 48 h. Intracellular PA14 gacA was not observed, even after 72 h (N = 15). We observed less dense matrix accumulation in gacA mutant-infected animals than with wild type P. aeruginosa, and did not observe microvillar shortening, intracellular invasion, or severe luminal distention. At all times during the infection by both wild-type and gacA P. aeruginosa, we only observed what appeared to be live P. aeruginosa cells, in contrast to S. aureus as described below. S. aureus infection: anal deformation, intestinal distention, enterocyte effacement, and cell lysis Interestingly, the cytopathology of S. aureus infection in the C. elegans intestine is markedly different from a P. aeruginosa infection. First, using GFP-labeled S. aureus, we observed rapid accumulation of bacteria in the intestine 4 h after infection initiation, whereas P. aeruginosa did not start accumulating until 8 after initial exposure (Figure S4A, B). At 4 h, S. aureus accumulated in the anterior and posterior ends of the intestine, and the rectum (Figure S3A, C), with less accumulation in the mid section of the intestinal lumen, where the bacteria appeared to be adhering to the apical surface of intestinal cells (Figure S3B). S. aureus accumulated further over the course of the following 4 h (Figure S4B). The infected animals moved slowly, were smaller (Figure S4C, D), and appeared to produce fewer eggs, than healthy animals. In addition to the intestinal distention and accumulation phenotypes, we observed a marked deformation of the anal region with S. aureus (Figure S4C) but not with P. aeruginosa (not shown) or non-pathogenic E. coli (Figure S4D). This deformed anal region (Dar) phenotype [46] appeared 4–8 h after initiation of infection and required live S. aureus (Figure S4E). Interestingly, this Dar phenotype was dependent on bar-1/β-catenin and mpk-1/extracellular signal regulated kinase (ERK) (Figures S4F, G, J, K, and L), which is also required for the Dar response to Microbacterium nematophilum [47]. We previously showed that bar-1/β-catenin and its downstream target gene egl-5/HOX exhibit a defective intestinal response to S. aureus [7]. Unexpectedly, mutants defective in egl-5/HOX exhibited a wild-type Dar response (Figure S4H, L), despite having an altered intestinal host response to S. aureus [7] and a defective anal swelling response to M. nematophilum infection [48]. Similarly, pmk-1/p38 MAPK mutants exhibited a slightly less noticeable, but equally frequent, Dar phenotype following S. aureus infection (Figure S4I, L), consistent with our previous observation that pmk-1 mutants are only subtly more susceptible to S. aureus-mediated killing [7]. These data suggest that the Dar phenotype may be a defensive host swelling response to pathogen-mediated host damage, since it requires an active host response to live bacteria. To investigate the cytopathology of S. aureus infection, we performed TEM of S. aureus-infected animals, focusing on the intestinal epithelium (Figure 2A). After 12 h of infection, we found a striking decrease in the length of the microvilli compared to animals feeding on non-pathogenic E. coli (Figures 2B–E, 3B–C). We also observed significant plasma membrane “blebbing” from the apical surface of intestinal cells (Figures 2D–E, 3C, S6A–B). The intestinal lumina of S. aureus-infected animals were markedly distended, consistent with our previous observations using light microscopy [17]. Distention was apparently a consequence of severe volume loss of the intestinal epithelial cells, with concomitant accumulation of bacterial cells in the enlarged luminal space (Figure 2B, D). In marked contrast to P. aeruginosa, an average 34% of S. aureus cells in the lumen (9–63%, N = 8 cross sections) were lysed at 12 h of infection (Figure 2E); these appeared similar to published TEM micrographs showing S. aureus cells killed with antimicrobial peptides in vitro [49], suggesting that the C. elegans intestine may produce bactericidal factors active against S. aureus. 10.1371/journal.ppat.1000982.g002 Figure 2 Intestinal distention, enterocyte effacement, and bacterial lysis at 12 h of S. aureus infection in C. elegans. A. Schematic representation of C. elegans body plan and plane of section (Left) and midbody transversal section indicating major organs (Right). B. Midbody transversal section of a control animal fed non-pathogenic E. coli. Square indicates area magnified in C. C. Higher magnification showing healthy microvilli (mv), intestinal epithelial cells (iec), apical junctions (aj), bacteria and bacterial debris in the intestinal lumen. D. Midbody transversal section of an animal infected with S. aureus for 12 h. Square indicates area magnified in E. E. Higher magnification showing short or absent microvilli, shrunken intestinal epithelial cells, apical junctions, and live and lysed bacteria in the distented intestinal lumen. Microvilli are false-colored red from the terminal web to the apical membrane, and intestinal epithelial cells are highlighted in yellow. Lysed bacteria in E. are false-colored blue. 10.1371/journal.ppat.1000982.g003 Figure 3 S. aureus causes intestinal cell effacement and lysis in C. elegans. A. Schematic representation of C. elegans body plan and plane of section (Left) and midbody transversal section indicating major organs (Right). Square highlights area of focus in TEM micrographs. B–I. TEM micrographs of transversal midbody sections of animals feeding on heat-killed non-pathogenic E. coli OP50 (B, D, F), virulent S. aureus NCTC8325 (C, E, G, I), or heat-killed S. aureus (H) for 12 h (B, C), 24 h (D, E), or 36 h (F–I). Scale bar, 1 µm. iec, intestinal epithelial cell; mv, microvilli; b, bacterial cell; bwm, body wall muscle; aj, apical junctions, asterisk in C indicates membrane blebbing. In contrast to animals feeding on E. coli, at 24 h of infection the microvilli were almost completely destroyed (Figure 3D, E) and at 36 h were completely absent, in what is termed “enterocyte effacement” (Figure 3F, G, I). Further, at 36 h we observed a reduction of intestinal cell volume (see thin sliver in Figure 3G) and intestinal cell lysis (Figure 3I). Also at 36 h, we observed a few dead animals, the organs of which were completely degraded except for the collagenous cuticle and an unidentified circular internal structure (Figure S5). Heat-killed S. aureus did not cause intestinal distention, microvillar effacement, intestinal cell lysis, or death (Figure 3H). These data show that S. aureus causes membrane and cytoskeletal rearrangements, as well as enterocyte effacement and destruction, possibly by secreted membrane-active bacterial toxins such as cytolysins or other pore-forming toxins (PFTs). Hemolysins α, β, and γ are known S. aureus cytolysins. However, they appeared to not be required for pathogenesis and killing, as a S. aureus strain lacking all three hemolysins exhibited similar kinetics of C. elegans killing as the isogenic wild type (Figure S7). Similarly, the α-hemolysin Δhla mutant was as capable of causing enterocyte effacement, intestinal distention, membrane blebbing, and intestinal cell lysis as wild type (Figure S6C–F). These results indicate that virulence factors other than the hemolysins are responsible for the observed intestinal cell lysis. S. aureus infection triggers an antimicrobial and detoxifying transcriptional host response Because host defense from, and digestion of, ingested bacteria are necessarily linked in bacterivorous animals such as C. elegans, the distinction between innate immune responses and digestive responses is blurred. Previous studies have investigated the long-term effects of ingestion of pathogenic bacteria, defining a common necrotic host response that is triggered by several pathogens after 24 h of infection [50]. To investigate gene expression changes more likely to be elicited directly by S. aureus detection, we evaluated gene expression at an earlier infection time, namely 8 h. We previously reported that P. aeruginosa induces a potent transcriptional host response early during infection of C. elegans, which significantly contributed to our understanding of C. elegans defense from P. aeruginosa infection [44]. To determine whether C. elegans mounts a similar host response to S. aureus infection, we performed whole-genome transcriptional profiling of animals infected with S. aureus for 8 h, relative to animals feeding on non-pathogenic E. coli. We found 186 transcripts that increased at least two-fold in abundance and 198 that decreased at least two-fold after infection (Table S1). Focusing on 46 genes up-regulated 4-fold or higher as a smaller sample, we found that the majority had potential xenobiotic detoxification or antimicrobial activities, consistent with their involvement in a protective host response (Table 1). In this group, a number of genes of unknown function appeared to encode short secreted polypeptides that may possess antimicrobial activities (Table 1). 10.1371/journal.ppat.1000982.t001 Table 1 C. elegans genes induced 4-fold or higher after 8 h infection with S. aureus. Cosmid Name Public name Sequence Description Fold Change Presumptive Function* Expression K08C7.5 fmo-2 Flavin-containing monooxygenase FMO 100.0 Detoxification Intestine, pharynx1,3 Y65B4BR.1 Y65B4BR.1 Phospholipase 14.7 Antimicrobial C45G7.3 ilys-3 Invertebrate lysozyme 14.3 Antimicrobial Intestine, pharynx, vulva3 Y46C8AL.4 clec-71 C-type lectin 9.8 Antimicrobial ZC443.5 ugt-18 UDP-glucoronosyl/UDP-glucosyl transferase 9.4 Detoxification M60.2 M60.2 Placental protein 11 9.2 Intestine2 C48B4.1 C48B4.1 Acyl-CoA oxidase I 8.9 Metabolism Intestine2 K11D12.4 cpt-4 Acetyltransferase 8.7 Metabolism Intestine2 K12G11.3 sodh-1 Zinc-containing alcohol dehydrogenase superfamily 8.5 Detoxification Intestine, pharynx2 F54F3.3 F54F3.3 Lipase 8.4 Antimicrobial Intestine2 T01C3.4 T01C3.4 Lipase 8.4 Antimicrobial ZK666.6 clec-60 C-type lectin, von Willebrand factor, type A 8.3 Antimicrobial Intestine3 B0222.4 tag-38 Glutamate decarboxylase 8.3 Metabolism C23G10.11 C23G10.11 Short, basic 7.4 Antimicrobial C45G7.2 ilys-2 Invertebrate lysozyme 7.3 Antimicrobial F46C5.1 F46C5.1 Short, basic 7.0 Antimicrobial Intestine2 F28G4.1 cyp-37B1 Cytochrome P450 6.9 Detoxification T07G12.5 T07G12.5 Xanthine/uracil/vitamin C permease family 6.9 Detoxification Intestine, vulva2 K01A2.2 far-7 Nematode fatty acid retinoid binding 6.6 Signaling F38E11.1 hsp-12.3 Heat shock protein Hsp20 6.6 Stress ZK507.4 ZK507.4 6.4 T22F3.11 T22F3.11 Permease, major facilitator superfamily transporter 6.3 Detoxification Intestine2 B0213.15 cyp-34A9 Cytochrome P450 6.1 Detoxification Intestine2 F09C8.1 F09C8.1 Phospholipase precursor 6.1 Antimicrobial Intestine2 T09H2.1 cyp-34A4 Cytochrome P450 6.0 Detoxification B0218.8 clec-52 C-type lectin 5.9 Antimicrobial Intestine, rectal gland3,4 F01G10.3 ech-9 Enoyl-CoA hydratase 5.8 Metabolism D1086.5 D1086.5 5.5 hypodermis?2 C34C6.7 C34C6.7 5.4 F36D3.9 cpr-2 Cysteine protease 5.4 Antimicrobial K02G10.7 aqp-8 Major intrinsic protein 5.4 Detoxification Intestine2 Y51H4A.5 Y51H4A.5 Lipase, class 3 5.3 Metabolism F58B3.2 lys-5 Lysozyme 5.2 Antimicrobial C35C5.8 C35C5.8 5.2 F53A9.8 F53A9.8 Short His-rich 5.1 Antimicrobial Intestine, rectal gland2,3 Y46C8AL.5 clec-72 C-type lectin 4.9 Antimicrobial Intestine2 Y5H2B.5 cyp-32B1 Cytochrome P450 4.9 Detoxification Intestine2 C29F7.2 C29F7.2 DUF1679; Predicted small molecule kinase 4.6 Detoxification Intestine2 F43C11.7 F43C11.7 Zn-finger, RING 4.5 Intestine2 T07H3.3 math-38 MATH domain 4.5 Signaling M05D6.7 gbh-2 Gamma-butyrobetaine, 2-oxoglutarate dioxygenase 4.4 Metabolism Hypodermal2 F41C3.1 F41C3.1 Protein of unknown function DUF1265 4.3 Y46C8AL.3 clec-70 C-type lectin 4.3 Antimicrobial Intestine, rectal gland, vulval, uterine, anal depressor muscles3 F46B6.8 F46B6.8 Lipase 4.2 Antimicrobial F58B3.1 lys-4 Lysozyme 4.1 Antimicrobial C50F7.5 C50F7.5 Pro-rich 4.1 Antimicrobial Intestine, rectal epithelial cells?2 References: 1 [99], 2NextDB (http://nematode.lab.nig.ac.jp/db2/index.php); 3This study, 4 [100]. *In the absence of further functional characterization, these predictions should be considered speculative. Bold public names indicate genes selected for qRT-PCR as biomarkers of the response. Functional predictions were based on known homologies and the presence of predicted signal peptides, and should be considered speculative until further functional characterization. To identify potential physiological roles of this host response, we used two complementary methods to study the over-representation of gene ontology (GO) classes (see Materials and Methods). These analyses revealed up-regulation of detoxifying and antimicrobial responses and down-regulation of growth-related metabolic pathways and extracellular structural components (Table S7). Significance of representation analysis revealed that the most significantly induced gene class contains sugar-binding proteins including C-type lectins (CTLs, N = 15, p = 1.2E-5), which could act as signaling receptors, opsonizing agents, or direct antimicrobial effectors [51], [52], [53], [54]. The most significantly repressed GO classes contain genes encoding structural constituents of the cuticle (e.g. collagens; N = 47, p = 4.4E-25), phosphate and inorganic anion transport (N = 47, p = 4.5E-25 and p = 5.8E-21), basement membrane components (N = 4, p = 1E-6), and lipid transporters (N = 5, p = 6E-6). A second approach (see Text S1) expanded these observations to include additional metabolic enzymes and transporters (Table S2). These analyses highlight the potential role of CTLs as a major immune effector strategy used by C. elegans during infection by S. aureus, and the significant metabolic component of the host response. Limited overlap with other stress responses Because we observed cell membrane rearrangements suggestive of the activity of membrane-active toxins (Figures 2E, 3C, E, G, I, S6), we hypothesized that part of the host response to S. aureus may also be triggered by PFTs. Indeed, we found that 22 of 422 probe sets up-regulated during exposure of worms to the Bacillus thuringiensis PFT Cry5B [55] were also induced during infection with S. aureus (Table S3), significantly higher than the 4 probe sets expected by chance alone. These data suggest that the overlapping set of genes shared by the responses to S. aureus and Cry5B may constitute a host response triggered by intestinal cell membrane disruption. Because we observed evidence of intestinal destruction and nutritional deprivation in animals infected with S. aureus, we hypothesized that infected animals might be starving. Previous work identified 18 genes whose expression changed during starvation [56]. In contrast, only one out of 9 previously identified fasting-induced genes (acs-2) and 4 out of 9 fasting-repressed genes (lbp-8, acdh-1, fat-7, and F08A8.2) were affected by S. aureus infection. Furthermore, the fasting-induced gene hacd-1 [56] was repressed during infection. These data suggest that the early transcriptional host response to S. aureus infection is minimally impacted by the starvation response. S. aureus infection triggers at least two waves of transcriptional response in the intestinal epithelium To validate the microarray experiments, we measured transcript levels for ten selected “biomarker” genes by qRT-PCR over a time course of infection. These ten genes, used as models of the larger host response, were chosen to represent different up-regulation levels and functional annotations (Table 2). All ten genes tested were induced in response to infection (Figure 4A, B). A subset of these biomarkers, fmo-2 (FMO), ilys-3 (lysozyme), cpr-2 (protease), Y65B4BR.1 (lipase), exc-5 (GEF), and F53A9.8 (putative antimicrobial peptide), were already induced 10-fold or higher by the first time-point at 4 h (Figure 4A). A second subset, lys-5 (lysozyme), clec-52, clec-60, and clec-71 (CTLs), were only modestly induced by 4 h and exhibited a further increase by 12 h (Figure 4B). Thus, time-resolved gene expression analysis revealed the existence of at least two kinetic groups, defined by their expression levels at 4 h. We also measured transcript levels for 8 genes predicted to be repressed upon infection, confirming reduced expression for 7 of them (Figure 4C). Together, these results confirm the predictive value of the genome-wide profiling. 10.1371/journal.ppat.1000982.g004 Figure 4 The C. elegans host response to S. aureus infection is comprised of two kinetic groups. A, B. qRT-PCR analysis of genes predicted to be up-regulated by microarray analysis. Transcript levels were measured in synchronized young adult wild-type animals feeding on heat-killed E. coli OP50 or infected with S. aureus NCTC8325 for 4, 8, and 12 h. Data are the means of three biological replicates, each replicate measured in duplicate and normalized to a control gene, expressed as the ratio of the corresponding S. aureus-induced levels and the basal E. coli levels. A. Immediate-early-induced genes were highly induced by 4 h of infection. B. Later induction of early response genes. C. qRT-PCR analysis of genes predicted to be repressed by microarray analysis. Transcript levels were measured as in A, after 8 h infection with S. aureus. Data are the means of three biological replicates, each replicate measured in duplicate and normalized to a control gene, expressed as the ratio of the corresponding S. aureus-induced levels and the basal E. coli levels. Error bars are SEM. 10.1371/journal.ppat.1000982.t002 Table 2 Selection of 10 biomarker genes to measure the host response to S. aureus. Cosmid Name Public name Pathogen-specificity (plus S. aureus)* Fold Change Prediction K08C7.5 fmo-2 E. faecalis (UP) 100.0 detoxification enzyme Y65B4BR.1 Y65B4BR.1 S. marcescens (UP) 14.7 antibacterial lipase C45G7.3 ilys-3 P. luminescens (UP) P. aeruginosa (DOWN) 14.3 lysozyme Y46C8AL.4 clec-71 P. aeruginosa (UP) 9.8 C-type lectin ZK666.6 clec-60 M. nematophilum (UP) 8.3 C-type lectin B0218.8 clec-52 P. aeruginosa (DOWN) 5.9 C-type lectin F36D3.9 cpr-2 E. faecalis (UP) 5.4 antibacterial protease F58B3.2 lys-5 E. faecalis (UP) P. aeruginosa (DOWN) 5.2 lysozyme F53A9.8 F53A9.8 E. faecalis, P. aeruginosa, M. nematophilum, P. luminescens, Cry5B (UP) 5.1 antibacterial peptide C33D9.1 exc-5 — N/S CDC-42 GEF *: based on microarray results; UP: up-regulated; DOWN: down-regulated; N/S: not significant by microarray. Genes were selected to represent a variety of induction levels, pathogen-specificities, and molecular functions. To elucidate the spatial pattern of the host response, we used transgenic animals carrying transcriptional reporters in which the promoters for 5 of the 10 biomarker genes (clec-52, clec-60, F53A9.8, fmo-2, and ilys-3), as well as clec-70, an additional CTL gene up-regulated by S. aureus and important for host defense (see below), were fused to GFP. We infected these transgenic animals with S. aureus and compared the intensity and pattern of GFP expression with control animals feeding on E. coli. All the genes tested were expressed at low basal levels in the intestines of the latter (Figure 5, left panels). After infection with S. aureus, all of the GFP reporters were induced in the intestinal epithelial cells (Figure 5, right panels). ilys-3, F53A9.8, clec-52, clec-60, and clec-70 were all expressed more strongly in the posterior end of the intestine than the anterior. ilys-3, fmo-2, and clec-70 were also induced in the pharynx. Although promoter-GFP fusions like these lack potentially important endogenous regulatory 3′ UTR and intronic sequences, these data are consistent with endogenous RNA localization data from ongoing genome-wide in situ hybridization studies of animals feeding on non-pathogenic E. coli (NextDB, http://nematode.lab.nig.ac.jp/db2/index.php, and Table 1). Thus, the C. elegans transcriptional host response to S. aureus is primarily localized in the intestinal epithelial cells and, in some cases, additional sites (Figure S8A–H). 10.1371/journal.ppat.1000982.g005 Figure 5 The C. elegans host response to S. aureus is induced in the intestinal epithelial cells. Transcriptional reporters using upstream sequences to fmo-2, ilys-3, F53A9.8, clec-52, clec-60, and clec-70 fused to GFP were induced in the intestinal epithelial cells after 24 h of S. aureus NCTC8325 infection (left panels) compared to parallel E. coli OP50-fed controls (right panels). Despite strong induction of fmo-2 and ilys-3 as measured by qRT-PCR, the corresponding GFP reporters exhibited low levels of expression; this could be due to their low basal expression on E. coli (not shown). Red, myo-2::NLS::cherry coinjection marker expressed in the pharynx in the head. A fold induction of 1 indicates no induction. a, anterior end; p, posterior end. Arrows indicate pharyngeal expression. ilys-3 was also expressed in the pharynx, in an unidentified cell superimposed on the pharynx, and in unidentified cells, possibly epithelial cells, in the vulva (Fig. S6A,B). clec-70 was also expressed in unknown cells in the pharynx (Fig. S6C) and in the uterine muscle (Fig. S6D), but only in one transgenic line of three. F53A9.8 was also expressed in a group of cells surrounding the rectum, possibly the rectal gland cells that secrete molecules into the rectal lumen (Fig. S6G, H). Distinct modes of detection of S. aureus and P. aeruginosa infection Up-regulation of the 10 biomarker genes could be a result of cell damage caused by S. aureus, or of microbial detection independent of the inflicted damage. To discriminate between these two scenarios, we measured gene induction during exposure to live S. aureus, which causes pathogenesis, or heat-killed S. aureus, which does not (Figures 3G–I and S4C, E). Unexpectedly, all 10 biomarker genes were induced at least equally well on heat-killed S. aureus as on live S. aureus (Figure 6A). This result suggested that the ten biomarker genes form part of a host response against microbe-derived molecules, possibly pathogen-associated molecular patterns (PAMPs) [3], and not of a damage response. 10.1371/journal.ppat.1000982.g006 Figure 6 Potential PAMP-mediated, TLR-independent sensing of S. aureus. A. qRT-PCR analysis of induced genes in animals feeding on heat-killed S. aureus. Transcript levels were measured in synchronized young adult wild-type animals infected with live S. aureus NCTC8325 or feeding on dead S. aureus or E. coli OP50 for 8 h. B. qRT-PCR analysis of induced genes in animals feeding on B. subtilis. Transcript levels were measured in synchronized young adult wild-type animals feeding on B. subtilis PY79 or heat-killed E. coli for 8 h. C. qRT-PCR analysis of induced genes in tol-1 mutant animals. Transcript levels were measured in synchronized young adult animals feeding on heat-killed E. coli or infected with S. aureus for 8 h. D. qRT-PCR analysis of genes induced by live or heat-killed P. aeruginosa. Transcript levels were measured in synchronized young adult animals feeding on heat-killed E. coli or P. aeruginosa, or infected with live P. aeruginosa PA14 for 4 h. In all cases, data are the means of two biological replicates, each replicate measured in duplicate and normalized to a control gene, expressed as the ratio of the corresponding bacteria-induced levels and the basal E. coli levels. Error bars are SEM. Bacterial cell-wall components, such as LPS and flagellin in Gram-negative bacteria and peptidoglycan in both Gram-negatives and Gram-positives, are common PAMPs in many systems [57]. To test whether Gram-positive cell wall components in general were able to trigger the same S. aureus-induced response, we assayed gene induction in animals feeding on non-pathogenic Gram-positive bacterium Bacillus subtilis compared with E. coli-fed controls. Remarkably, of the ten biomarker genes, only fmo-2, ilys-3, and Y65B4BR.1 were induced by B. subtilis, albeit at much lower levels than with S. aureus (Figure 6B). The remaining 7 genes either were not induced or were repressed by B. subtilis. These results suggest that PAMPs other than shared Gram-positive cell wall molecules may be molecular triggers for the S. aureus-induced host response in C. elegans (see Discussion). How is S. aureus detected? In fruit flies and mammals, Toll-like receptors (TLR) are involved in PAMP detection. C. elegans has a single gene encoding a TLR, tol-1, which has been shown to be important for avoidance responses to Serratia marcescens and for full induction of antimicrobial peptide abf-2 in response to Salmonella enterica [58], [59]. However, tol-1 was not required for the induction of any of the 10 S. aureus–induced biomarker genes (Figure 6C). Furthermore, tol-1 mutants were not more susceptible to S. aureus-mediated killing than wild type (Figure S10). One caveat is that the tol-1(nr2033) allele used, a deletion that eliminates the cytoplasmic TIR domain necessary for signaling [60], is viable, thus considered a partial loss of function for viability but a null allele for immune signaling [58]. These results show that tol-1 is most likely not required for the C. elegans host response to S. aureus and suggest that alternative mechanisms may exist for PAMP detection. Since mpk-1/ERK is important for the rectal epithelial cell swelling response to M. nematophilum [47] and for the swelling response to S. aureus (Figure S4K, L), we wondered whether mpk-1 might also be important for the intestinal response to S. aureus. However, mpk-1 animals exhibited no measurable defect in the induction of the biomarker genes relative to E. coli-fed controls (Figure S9). Therefore, mpk-1 is dispensable for at least part of the intestinal transcriptional response to S. aureus, but not the anal swelling response. In contrast to S. aureus, which either alive or dead triggered the induction of 10 biomarker genes (Figure 6A), heat-killed P. aeruginosa did not trigger the induction of 10 P. aeruginosa-induced biomarker genes (Figure 6D). Together, these data are consistent with the idea that C. elegans may recognize S. aureus infection mainly via TLR-independent PAMP detection, whereas it may recognize P. aeruginosa infection via detection of either Damage-Associated Molecular Patterns (DAMPs), or unidentified heat-labile PAMPs. P. aeruginosa and S. aureus trigger partially overlapping host responses Host responses to pathogenic attack in plants and animals are remarkably pathogen-specific. In C. elegans, pathogen-specific gene induction has been observed at late times of infection (i.e., 24 h), when damage and necrosis are apparent [50]. To determine whether pathogen-elicited gene induction at earlier times (i.e., 8 h) is also pathogen-specific, we compared the host response to S. aureus with our previously published study of the early host response to P. aeruginosa [44]. Of 186 genes induced by S. aureus and 259 genes induced by P. aeruginosa, 44 genes were induced by both pathogens, which is more than expected by chance (Figure 7A, Table 3). We validated the results of the microarray comparison using qRT-PCR; four genes predicted to be specifically induced by P. aeruginosa were either unaffected or repressed by S. aureus (Figure 7C), and four genes predicted to be induced by S. aureus were either unaffected or repressed by P. aeruginosa (Figure 7D). In contrast, four of five predicted overlap genes showed increased expression with both S. aureus and P. aeruginosa (Figure 7E). This suggests that the early host response to pathogen attack involves activation of a “pan-pathogen” response against a broad spectrum of pathogens, as well as a more tailored response that is optimized for the defense against the specific class of pathogens that is causing the infection. 10.1371/journal.ppat.1000982.g007 Figure 7 The C. elegans host response is comprised of pathogen-specific and -shared components. A. Comparison of genes induced 2-fold or higher (p≤0.01) upon infection with S. aureus for 8 h, P. aeruginosa for 8 h, and M. nematophilum for 6 h. Gene identities are presented in Table 3. B. Quadrant analysis of genes with expression changes both during S. aureus and P. aeruginosa infection. X axis, Fold Change for each gene during P. aeruginosa infection (log10 values). Y axis, Fold Change for each gene during S. aureus infection (log10 values). C. qRT-PCR analysis of P. aeruginosa–induced genes during S. aureus infection. D. qRT-PCR analysis of S. aureus–induced genes during P. aeruginosa infection. Transcript levels were measured in synchronized young adult wild-type animals feeding on non-pathogenic E. coli OP50 or infected with pathogen for 8 h (S. aureus NCTC8325) or 4 h (P. aeruginosa PA14). Results are the average of three biological replicates, each replicate measured in duplicate and normalized to a control gene, expressed as the ratio of the corresponding P. aeruginosa -induced levels and the basal E. coli levels. E. qRT-PCR analysis of overlap genes during S. aureus or P. aeruginosa infection. Transcript levels were measured in synchronized young adult wild-type animals feeding on non-pathogenic E. coli or infected for 8 h or 4 h, respectively (on P. aeruginosa, these genes were predicted to change by 4 h as well as 8 h). Results are the average of three biological replicates, each replicate measured in duplicate and normalized to a control gene, expressed as the ratio of the corresponding pathogen-induced levels and the basal E. coli levels. Previous microarray analysis wrongly predicted acs-2 to be induced by P. aeruginosa, the only example of this kind that we have found. 10.1371/journal.ppat.1000982.t003 Table 3 Pathogen specificity of the response to infection. a. S. aureus AND P. aeruginosa AND M. nematophilum Name Description Signal Peptide acs-2 Long-chain-fatty-acid-CoA ligase + C15A11.7 Amine oxidase, similar to tetracycline resistance protein TetB − C50F4.9 Contains similarity to Phoneutria reidyi Neurotoxin PRTx3–7 + ech-9 Enoyl-CoA hydratase − F52F10.3 Acyltransferase 3 family − F53A9.8 His-rich short polypeptide − glc-1 Alpha subunit of a glutamate-gated chloride channel + H02F09.3 Contains similarity to Staphylococcus haemolyticus Serine-rich adhesin for platelets precursor.; SW:Q4L9P0 + K05F1.10 Contains similarity to Interpro domains IPR002919 (Protease inhibitor I8, cysteine-rich trypsin inhibitor-like), IPR013032 (EGF-like region) + ZC443.3 Contains similarity to Saccharomyces cerevisiae essential protein involved in intracellular protein transport, coiled-coil protein necessary for transport from ER to Golgi; required for assembly of the ER-to-Golgi SNARE complex; SGD:YDL058W − b. S. aureus AND M. nematophilum NOT P. aeruginosa Name Description Signal Peptide C07A9.8 Putative membrane protein, Bestrophin − ilys-2 Invertebrate lysozyme + clec-60 C-type lectin, von Willebrand factor, type A + clec-61 C-type lectin, von Willebrand factor, type A + clec-62 C-type lectin, von Willebrand factor, type A + clec-70 C-type lectin + F09F7.6 − F54F3.3 Lipase + R07C12.1 − sodh-1 Zinc-containing alcohol dehydrogenase − T22F3.11 Permease, major facilitator superfamily transporter + c. S. aureus AND P. aeruginosa NOT M. nematophilum Name Description Signal Peptide C18A11.1 Short (119 aa) − C23G10.11 Short (63 aa) + C29F7.2 DUF1679; Predicted small molecule kinase − C34C6.7 − C46H11.2 Flavin-containing monooxygenase − C50F7.5 Pro-rich + clec-71 C-type lectin + cpt-4 Carnitine O-acyltransferase CPTI − cyp-34A9 Cytochrome P450 CYP2 subfamily + D1086.5 + dod-22 CUB-like domain + F16C3.2 + F21E9.3 Transthyretin-like family + F21F8.4 Aspartyl protease + F41G4.8 + F45D3.4 + F46B3.1 Metridin-like ShK toxin + F49H6.13 DUF1164 + F53A9.1 Short His/Gly rich (77 aa) − F53A9.2 Short His/Gly rich (83 aa) − F53A9.6 Short His/Gly rich (86 aa) − far-7 Nematode fatty acid retinoid binding protein − gbh-2 Gamma-butyrobetaine,2-oxoglutarate dioxygenase − hsp-12.3 Alpha-B-crystallin − ins-11 Insulin-like peptide of the insulin superfamily of proteins + M60.2 Placental protein 11 + mpk-2 Mitogen-activated protein kinase − srsx-34 Permease, major facilitator superfamily − T01C3.4 Triacylglycerol lipase + T04F8.7 − ugt-18 UDP-glucoronosyl/UDP-glucosyl transferase + Y40B10A.6 O-methyltransferase, family 3 − Y43C5A.3 Short, basic Gly-rich (108 aa); contains similarity to Ixodes scapularis Putative secreted salivary gland peptide.; TR:Q5Q982 + Y60C6A.1 Protein of unknown function CX + d. M. nematophilum AND P. aeruginosa NOT S. aureus Name Description Signal Peptide B0024.4 DUF274 + C17H12.8 CUB-like domain + C31A11.5 Predicted acyltransferase + C49C8.5 Ficolin and related extracellular proteins + clec-67 C-type lectin + clec-69 C-type lectin + clec-86 C-type lectin + F09B9.1 Predicted acyltransferase + F35E12.5 CUB-like domain + F52F10.3 Predicted acyltransferase − M28.8 + Y41D4B.16 DUF274 + Y46C8AL.2 C-type lectin, Proline-rich region + ZK896.5 CUB-like domain + Partially overlapping responses between S. aureus, P. aeruginosa and M. nematophilum (a), S. aureus and M. nematophilum (b), S. aureus and P. aeruginosa (c), or M. nematophilum and P. aeruginosa (d). Genes encoding products with signal peptides suggestive of secretion are indicated. To further test the hypothesis of the existence of pathogen-shared and -specific components of the host response, we compared the genes differentially affected by S. aureus infection with previously published profiling of C. elegans infected with M. nematophilum [61]. The comparison of microarray studies independently performed in these separate laboratories may underestimate overlapping gene sets due to the use of different infection, RNA extraction, and data processing methods. Nonetheless, we found a relatively high degree of overlap between the responses to S. aureus and M. nematophilum (21 genes out of 186); 10 genes were induced by S. aureus, M. nematophilum, and P. aeruginosa (Figure 7A, Table 3). In contrast, 44 (68%) out of 65 genes induced by M. nematophilum were not induced by S. aureus. These data further support the existence of a core, shared response and a specific, tailored response. GO annotations of genes affected by S. aureus or P. aeruginosa also exhibit a degree of specificity. Whereas the S. aureus-induced host response includes many sugar binding proteins, the P. aeruginosa-induced response is not characterized by any particular over-represented GO annotation (Table S7). Additionally, whereas the repressed response to S. aureus consists of many transporters and cuticle components, the repressed response to P. aeruginosa is mostly represented by metabolic pathways (Table S7). Interestingly, both repressed responses included basement membrane genes (N = 5, p = 1.4E-6 for P. aeruginosa), suggesting host growth suppression by both types of infection. To investigate whether there were correlated or anti-correlated components of the responses to S. aureus and P. aeruginosa, we focused on genes whose expression changed both during infection with S. aureus and infection with P. aeruginosa. We broke down the two responses by plotting genes whose expression changed more than 2-fold with S. aureus (Y axis in Figure 7B) and with P. aeruginosa (X axis in Figure 7B). Genes whose expression changed only during infection with one of the two pathogens were thus not included. This method defined four quadrants: I) Genes induced by S. aureus and repressed by P. aeruginosa; II) Genes induced by S. aureus and P. aeruginosa; III) Genes repressed by S. aureus and induced by P. aeruginosa; and IV) Genes repressed by S. aureus and P. aeruginosa (Figure 7B). NextBio biogroup representation analysis (see Materials and Methods) on each quadrant failed to detect any over-represented biogroup in Quadrants I and III. However, in Quadrant II (genes up-regulated by both pathogens) several biogroups were over-represented, e.g. genes involved in detoxification, iron sequestration, and energy generation (Table S2a, b). Likewise, biogroups over-represented in Quadrant IV (down-regulated by both pathogens) included transporters, cuticle components, and fatty acid (FA) β-oxidation (Table S2a, b). The common repression of FA β-oxidation is not consistent with a starvation response, where β-oxidation is induced [56]. Furthermore, only 3 of 9 fasting repressed genes (lbp-8, acdh-1, and F08A8.2) [56] were repressed during P. aeruginosa infection [44]. Similarly to S. aureus, the fasting-repressed gene hacd-1 was induced during P. aeruginosa infection [44]. Together, these observations show the distinct nature of the host response to distinct pathogens, and highlights metabolic components of early host responses to infection. Because overlapping gene expression changes measured by qRT-PCR do not necessarily imply the involvement of the same tissues during infection with different pathogens, we further investigated the expression of clec-60::GFP (up-regulated by S. aureus and M. nematophilum, Figure S11A, B, C, D) and F53A9.8::GFP (up-regulated by S. aureus, M. nematophilum, and P. aeruginosa, Figure S11E, F, G, H). clec-60::GFP was induced by M. nematophilum and by S. aureus in the intestine (Figure S11C, D), and down-regulated by P. aeruginosa (Figure S11B) compared with non-pathogenic E. coli (Figure S11A). Likewise, F53A9.8::GFP was induced by all three pathogens in the intestine (Figure S11F, G, H) compared with E. coli controls (Figure S11E). These data suggest that components of the host response are induced in the intestine by distinct pathogens. Genes induced by S. aureus influence host survival To determine whether S. aureus-induced genes have protective functions in host defense, we performed whole-animal RNAi knockdown of 42 of the 46 most highly up-regulated genes (Table 1), and identified 6 genes [tag-38 (Glutamate decarboxylase/sphingosine phosphate lyase), sodh-1 (sorbitol dehydrogenase), cyp-37B1 (Cytochrome P450), F43C11.7 (F-box containing protein), math-38 (MATH domain-containing signaling protein), and clec-70 (secreted CTL)] whose decreased expression caused enhanced susceptibility to killing by S. aureus (Figure 8A,B, C), but not P. aeruginosa (Figure 8G). We also identified one gene,Y51H4A.5 (a putative intracellular lipase) whose decreased expression caused slightly enhanced resistance to S. aureus killing, suggesting that its expression is detrimental to the host, or that it functions as a repressor of host defense (Figure 8B). Importantly, the lifespan of animals continuously fed dsRNA-expressing, non-pathogenic E. coli was near wild type, except for F43C11.7, which actually caused increased lifespan (Figure 8D). These data show that S. aureus-induced genes have important functions in pathogen-specific host defense, and that the enhanced susceptibility to S. aureus mediated by RNAi knockdown is not due to a non-specific decrease in viability. Although the molecular identities of these genes offer clues to their potential functions, the elucidation of their exact mechanisms of action will require further study. 10.1371/journal.ppat.1000982.g008 Figure 8 Host response genes are biologically relevant to host survival during S. aureus infection. A, B, C. After RNAi knockdown of S. aureus-induced genes, animals were transferred to S. aureus NCTC8325 infection plates. Survival statistics: vector in A (MS = 82 h; N = 125), tag-38 (MS = 66 h, N = 142, p = 0.0002), sodh-1 (MS = 66 h; N = 142; p = 0.009), cyp-37B1 (MS = 66 h; N = 145; p<0.0001), F43C11.7 (MS = 66 h; N = 147; p = 0.0003), vector in B (MS = 71 h; LT50 = 67.6 h, N = 99/8), math-38 (LT50 = 48.6 h; N = 92/7; p = 0.0012), Y51H4A.5 (MS = 91 h; N = 100; p = 0.0122); vector in C (LT50 = 68.1 h; N = 91), clec-70 (LT50 = 67.9 h; N = 98; p = 0.0145). D. Lifespan of E. coli-fed animals. Lifespan statistics: vector (MS = 12.8 d; N = 89), tag-38 (MS = 12.8 d, N = 93, p = 0.1708), sodh-1 (MS = 11.1 d; N = 104; p = 0.1887), cyp-37B1 (MS = 12.8 d; N = 95; p = 0.1473), clec-70 (MS = 12.8 d; N = 99; p = 0.0354), F43C11.7 (MS = 14.08 d; N = 88; p<0.0001). E, F. Overexpression of host response genes enhances host survival during S. aureus infection. E. Animals carrying lys-4,5 cluster transgenes survived longer (LT50 = 54.1 h; N = 60; p<0.0001) during S. aureus infection than control animals bearing transgenes composed of coinjection marker and clec-60::GFP promoter fusion (LT50 = 40.6 h; N = 91/4). F. Animals carrying an integrated array containing a cluster of clec-70,71 (MS = 68 h; N = 94; p<0.0001) were more resistant to S. aureus mediated killing than control animals bearing rol-6 coinjection marker alone (MS = 61 h; N = 90). G. RNAi knockdown of S. aureus induced genes did not result in enhanced susceptibility to P. aeruginosa. Survival statistics: vector (MS = 47 h, N = 80/9); sodh-1 (MS = 47 h, N = 84/8, p = 0.2476); F43C11.7 (MS = 47 h, N = 83/9, p = 0.2592); cyp-37B1 (MS = 47 h, N = 92/11, p = 0.0201); Y51H4A.5 (MS = 47 h, N = 83/5, p = 0.6712); math-38 (MS = 47 h, N = 85/13, p = 0.9919); tag-38 (MS = 47 h, N = 90/10, p = 0.3373); clec-70 (MS = 47 h, N = 84/8, p = 0.0184). H. Overexpression of S. aureus induced C-type lectin genes resulted in mild enhanced susceptibility to P. aeruginosa. clec-60::GFP,myo-2::mCherry control (MS = 72 h; N = 98/20), clec-60,61,myo-2::mCherry (MS = 47 h; N = 94/16; p = 0.0408), clec-70,71, myo-2::mCherry (MS = 47 h; N = 92/12; p = 0.0002). Potential immune effectors identified in our analysis include antimicrobial peptides, lysozymes (enzymes that degrade peptidoglycan in the bacterial cell wall), and CTLs [62]. To test whether lysozymes or CTLs induced by S. aureus can confer resistance to S. aureus-mediated killing when expressed to higher levels than in the wild type, we constructed transgenic C. elegans carrying multiple copies of lys-4 and lys-5, clec-60 and clec-61, or clec-70 and clec-71 (each is a pair of genes that are adjacent to each other in the genome). Transgenic animals carrying the lys-4,5 or clec-60,61 clusters survived significantly longer than transgenic control animals (Figure 8E; Figure S12). Transgenic animals carrying the clec-70,71 cluster extrachromosomally did not exhibit enhanced resistance (not shown); however, three independent spontaneous integrant lines did (Figure 8F). Interestingly, strains with multiple copies of either cluster of C-type lectin genes exhibited enhanced susceptibility to P. aeruginosa-mediated killing (Figure 8H). Collectively, the data suggest that pathogen-induced intestinal expression of epithelial detoxifying and antimicrobial proteins is an important and pathogen-specific mechanism of C. elegans host defense against S. aureus infection. Discussion Among the bacteria that cause intestinal infections in C. elegans, the best-studied is P. aeruginosa, a Gram-negative human pathogen. Despite many advances in understanding P. aeruginosa-C. elegans interactions, little was known about the morphological and cell biological consequences of infection in vivo. This report provides unprecedented high-resolution description of bacterial intestinal infections of clinical relevance in C. elegans, with emphasis on the comparative cytopathology of infection by P. aeruginosa and S. aureus. P. aeruginosa and S. aureus cause markedly different symptoms in C. elegans. During infection with P. aeruginosa, we observed marked intestinal distention, extracellular matrix accumulation in the intestinal lumen, extracellular material accumulation on the apical surface, enlargement of the rough endoplasmic reticulum (RER), and abnormal autophagy in the host intestinal cells. The intestinal distention is likely a result of loss of cytoplasmic volume of the intestinal cells; the identity of the extracellular material is currently unknown. One possibility is that it is a biofilm matrix produced by P. aeruginosa, perhaps as a defensive mechanism. Alternatively, it could be produced by the host in response to P. aeruginosa, but not S. aureus. Further study is required to elucidate the origin of this substance. The cause of abnormal autophagy is also unknown; however, the aberrant structures we observed have also been described in unc-52 mutant animals, which are defective in the early steps of autophagosome assembly [63]. Thus, it is possible that P. aeruginosa infection causes autophagy arrest. It is tempting to speculate that this may represent a virulence mechanism deployed by P. aeruginosa to evade autophagic clearance of intracellular bacteria, an important host defense mechanism in the intestinal cells of C. elegans [64] and humans [65]. The intracellular invasion we observed provides a rationale for the benefit to P. aeruginosa of inhibiting host autophagy. gacA mutant P. aeruginosa was defective in inducing these phenotypes, indicating that GacA orchestrates virulence mechanisms related to host cell disruption in C. elegans. The molecular identity of these virulence mechanisms remains unknown; however, the observed putative OMVs may provide a mechanism for delivery of bacterial virulence factors to C. elegans intestinal cells. Perhaps the reliance on OMVs for virulence may explain why P. aeruginosa defective in type III secretion, an alternative mechanism of virulence factor delivery to host cells, are not defective in nematode killing [66]. OMV production is induced in P. aeruginosa by cellular stress [67]; therefore, the abundant OMV production we observed may indicate that P. aeruginosa perceives the intestinal lumen as a stressful environment, presumably as a result of defensive host factors secreted by the intestinal cells. The molecular identity of such putative intestinal defense factors remains unknown. In previous work, we defined the early transcriptional host response to P. aeruginosa [68]. Among the genes that are up-regulated by P. aeruginosa infection, several encode putative antimicrobial factors such as ShK toxins. Whatever the molecular identity of the antibacterial factors induced during P. aeruginosa infection, heat-killed P. aeruginosa did not induce a set of 10 biomarkers of the response. This observation is consistent with our previous studies suggesting that P. aeruginosa virulence may be required, at least partially, for response induction [44], [69]. These results are consistent with several interpretations. It is possible that detection of virulent P. aeruginosa is mediated by recognition of the damage inflicted on the host cells, e.g., via DAMP perception [70], or by recognition of PAMPs in the context of host damage, in what has come to be known as a pattern of pathogenesis [71]. Alternatively, PAMPs released only by live bacteria (PAMP-per vitae, or PAMP-PV) may be specifically recognized by C. elegans as a trigger for the response [71]. A third option is the release of PAMP-post mortem (PAMP-PM) upon heat inactivation of P. aeruginosa, which in turn could dampen the host response. PAMP-PM detection has been proposed to be a mechanism used by mammals to limit inflammatory damage to the host once the infection has been controlled [71]. Finally, detection of P. aeruginosa could be mediated by heat-labile signals that were destroyed during heat-inactivation in our experiments. Although further work is required to conclusively show which of these scenarios is correct, we have found that P. aeruginosa strains with increasing levels of virulence cause increasing levels of induction of C. elegans host response gene irg-1 [69], suggesting that the extent of host cell damage determines the magnitude of the host response to the hypothetical P. aeruginosa-produced PAMPs or DAMPs that are detected. During S. aureus infection, we observed rapid intestinal colonization, swelling of the anus, and effacement and destruction of intestinal epithelial cells, providing important mechanistic information about S. aureus-mediated pathogenesis of host epithelial cells in vivo. The molecular mechanism of cell destruction remains unknown; here we show that it is hemolysin-independent and is abrogated by heat inactivation of S. aureus. One explanation for this observation is that host tissue damage may be caused by the active pathogen, and not an unbridled host response. Alternatively, unknown heat-labile S. aureus toxins may cause cell lysis, as can occur in human cells [72]; further study is required to distinguish between these possibilities. Previous experiments using fertile animals showed that α-hemolysin Δhla mutant S. aureus were defective in C. elegans killing [17]. To our surprise, the Δhla Δhlb, Δhla Δhlg, or Δhla Δhlb Δhlg hemolysin mutants did not exhibit any killing defect in our assay using sterile animals (Figure S7). Thus, it is possible that α-hemolysin-mediated killing of C. elegans requires internal hatching of eggs retained inside the mother as a result of stress. Eventually, the pathogenic process results in internal tissue degradation and nematode death. These steps recapitulate key features of S. aureus infection in mammals both in vivo and in vitro, e.g. enterocyte effacement during intestinal infection, and cell lysis [37], [38], [39], [40], [41], [42], [43]. Thus, we propose that the C. elegans-S. aureus model has significant relevance to the study of conserved virulence mechanisms used by S. aureus to evade host epithelial defenses and attack host epithelial cells in general. Transcriptional profiling showed that S. aureus infection elicits changes in expression of a minor fraction of the total genome of ∼22,000 genes, indicating high specificity. This early response does not require live bacteria, suggesting that it involves detection of S. aureus per se (perhaps through PAMP perception) as opposed to indirectly through host cell damage (through DAMPs) [70]. Whatever the relevant PAMPs are, it is clear that they are not shared between S. aureus and B. subtilis, also a Gram-positive bacterium. For example, the peptidoglycan differs greatly between these two species [73], and could potentially be differentially sensed by C. elegans. Other possibilities include differential detection of surface-expressed lipoteichoic acids or differentially expressed surface proteins. Further work is required to elucidate the nature of such signal(s). Our results provide insight into the cellular biological effects of pathogenic infection on the epithelial barrier in vivo, as well as the early defense mechanisms deployed by C. elegans to fend off attack. The affected genes represent evolutionarily conserved categories relevant to human innate immunity. To gain insight into evolutionarily conserved effector mechanisms of host defense, we compared the genes up-regulated in C. elegans with previously published data using human neutrophils, which are important effector cells in human innate immunity [74]. When grouped by molecular function, we found some overlapping functional classes during the C. elegans and human neutrophil responses to S. aureus infection (Table S4). These classes included detoxification factors [e.g. transporters, UDP-glucuronosyltransferases (UGTs), cytochrome P450s, GSTs, and flavin-containing monooxygenases (FMOs), [75]], antimicrobial effectors (e.g. CTLs, peptidases, and proteases), galectins [76], and signaling components including EGF-like domain containing proteins, Cdc42 guanyl nucleotide exchange factors (GEFs), F-box proteins, mitogen-activated protein kinases (MAPKs), and Leucine-rich repeat domain (LRR) containing proteins. This observation shows that the C. elegans host response to infection shares important components with the human cellular host response, and suggests that human innate responses have ancient components that are conserved across phylogeny. Indeed, it is thought that modern vertebrate innate immunity represents an accretion of ancient invertebrate innate defenses [77]. Comparative genomics identified pathogen-specific as well as pathogen-shared components of the host response. This observation, consistent with similar ones recently made by others using different approaches [50], [78], [79], illustrates how diverse pathogens affect distinct aspects of host physiology as reflected in the distinct nature of the host responses. We also found overlap in gene expression patterns among the responses to three different pathogens, S. aureus (intestinal infection, Gram-positive), M. nematophilum (cuticular infection, Gram-positive), and P. aeruginosa (intestinal infection, Gram-negative), defining a core induced response that involves intracellular detoxification, iron sequestration, and sugar binding. In addition to a common set of up-regulated genes, we observed a repressed core response common to S. aureus and P. aeruginosa that involves anion transport, growth-related genes, lipid- and alcohol-metabolic genes, and acyl-CoA dehydrogenases. The fact that metabolic regulation is a major component of the C. elegans host response to bacterial pathogens provides a rationale to investigate metabolic changes that occur in higher organisms as a result of infection, particularly in innate defense tissues such as epithelia. Recent reports suggest a significant metabolic component during the host response in mammals as well [80], [81], [82]. The non-overlapping responses to S. aureus and P. aeruginosa may reflect the different virulence strategies of the two pathogens and/or may be a consequence of the distinct molecular composition of Gram-positive and -negative cell walls. Further studies are required to dissect the relative contribution of each factor, including the survey of additional Gram-positive and -negative infections in C. elegans. In a first step along those lines, we found that the Gram-positive non-pathogenic bacterium B. subtilis does not induce the same set of 10 biomarkers as S. aureus. Additionally, of 531 genes up-regulated by the Gram-positive pathogen E. faecalis [50], only 15 were shared with the response to S. aureus (Table S5a). The same report characterized the C. elegans late host response (i.e., 24 h) to four additional pathogens, identifying a set of shared genes that defined a pathogen-shared necrotic response [50]. During S. aureus infection we observed up-regulation of none of 16 up-regulated shared late response genes, and down-regulation of only three of six down-regulated shared response genes (Table S5c), suggesting that the host response evolves significantly over time. Mammalian intestinal epithelial cells directly sense and respond to bacterial stimulation [2], by inducing the expression of antimicrobial genes such as CTLs [53]. Similarly, most of the early C. elegans response to S. aureus occurs in the epithelial cells of the intestine. In Drosophila, mice, and humans, TLRs are important receptors that drive the activation of signaling cascades downstream of microbial stimulation. In C. elegans, however, loss of function of the sole TLR does not result in a defective immune response to S. aureus. Furthermore, C. elegans does not have an NF-kB homolog nor an inflammasome, raising the possibility that in mammals as in C. elegans, at least a portion of the immune response to S. aureus may be regulated independently of the TLR/NF-kB signaling axis [6], [51], [83]. Indeed, we previously reported that β-catenin and HOX genes are required for perception of pathogenic attack by S. aureus to drive the expression of epithelial host response genes [7]. Significantly, we also found that β-catenin and HOX proteins modulated NF-κB signaling in a human epithelial cell line during TLR2 stimulation, illustrating that previously unknown human innate immunity pathways can be identified using C. elegans [7]. We identified 6 host factors out of 42 tested whose lowered expression caused enhanced susceptibility to S. aureus. This corresponds to a 14% hit rate, which was much greater than expected; we had assumed that there would be significant functional redundancy among C. elegans immune effectors. Moreover, RNAi typically exhibits incomplete penetrance and expressivity. In addition to these 6 genes, knockdown of Y51H4A.5 (lipase) caused mild resistance to S. aureus mediated killing, suggesting that Y51H4A.5 acts to limit survival as a negative regulator of the host response or by harming the host instead of protecting it. RNAi of F43C11.7 caused enhanced susceptibility to S. aureus, yet extended lifespan on non-pathogenic E. coli. This is an example of genes that have opposite effects on host defense and lifespan regulation, indicating that these related processes are genetically separable [84]. Increased expression of host defense genes provides a mechanistic explanation of C. elegans defenses, as we found that animals carrying multiple copies of three genomic clusters of lysozymes or CTLs exhibited enhanced resistance to S. aureus. Lysozymes are well-known, evolutionarily ancient antibacterial effector molecules that degrade peptidoglycan and are also produced in human intestinal epithelial cells [1]. Recent studies have shown that some vertebrate CTLs, including human HIP/PAP, have direct bactericidal activity [53]. clec-70 and clec-71 share similar domain architectures with HIP, and clec-60 and clec-61 share similarities with arthropod receptor CTLs, suggesting that they may function respectively as antimicrobials or receptors in the C. elegans host response [85], [86], [87]. Collectively, these observations show that the large number of pathogen-response genes contribute cumulative, incremental defense functions to host survival. It is interesting that elements of the C. elegans immune response enhance host survival during infection with S. aureus (a human pathogen), supporting the notion that pathogen detection and response, as well as mechanisms of bacterial pathogenesis, share conserved features among distantly related hosts or microbes, respectively. It has been proposed that pathogens have experienced stepwise additions of virulence factors, as they evolved to survive different host antimicrobial responses, and to colonize new niches [12]. Our studies of the C. elegans-S. aureus system may thus probe an early step in the evolution of S. aureus as a pathogen and its interaction with prototypical metazoan epithelial cells. In humans, unknown host and bacterial factors determine whether S. aureus will become an innocuous member of the normal microbiota, or whether it will switch to a more virulent state and become a serious pathogen [23]. In this light, studies of the C. elegans intestinal epithelial response to S. aureus provide a unique starting point to identify previously unknown signaling pathways and molecular mechanisms of host immune response to bacterial virulence. Understanding how S. aureus disrupts host defense and causes host damage and death is critical to identifying new therapeutic targets to treat infectious disease. Materials and Methods Strains C. elegans was grown on nematode-growth media (NGM) plates seeded with E. coli OP50-1 at 15–20°C according to standard procedures [88]. C. elegans strains used in this study are detailed in Table S6a. Bacterial strains are detailed in Table S6b. Electron microscopy Wild type N2 Bristol animals were synchronized by hypochlorite treatment and L1 arrest and incubated on NGM plates seeded with E. coli OP50-1. Late L4 animals were collected and plated on 15 cm TSA plates seeded with live S. aureus NCTC8325 or heat-killed NCTC8325, and parallel NGM plates seeded with OP50-1. After 12, 24, and 36 h incubation at 25°C, animals were collected and incubated in fixation buffer (2.5% glutaraldehyde, 1.0% paraformaldehyde in 0.05 M sodium cacodylate buffer, pH 7.4 plus 3.0% sucrose). During the initiation of fixation, animals were cut in half with a surgical blade in a drop of fixative under a dissecting microscope, fixed overnight at 4°C, rinsed in 0.1 M cacodylate buffer, post-fixed in 1.0% osmium tetroxide 0.1 M cacodylate buffer, rinsed in buffer and water, and stained en bloc in 2% aqueous uranyl acetate. After rinsing in water, animals were embedded in 2% agarose in phosphate buffer saline, dehydrated through a graded series of ethanol washes to 100%, then 100% propylene oxide, and finally 1∶1 propylene oxide:EPON overnight. Blocks were infiltrated in 100% EPON and then embedded in fresh EPON overnight at 60°C. Thin sections were cut on a Reichert Ultracut E ultramicrotome and collected on formvar-coated gold grids. Sections were post-stained with uranyl acetate and lead citrate and viewed using a JEOL 1011 transmission electron microscope at 80 kV with an AMT digital imaging system (Advanced Microscopy Techniques, Danvers, MA). For each observation, whenever possible at least 10 cross-sections were evaluated, and representative images were chosen. Microarray analysis C. elegans growth and infection fer-15(b26)ts;fem-1(hc17) animals were synchronized by hypochlorite treatment and L1 arrest [88]. Arrested L1 larvae were placed onto NGM plates seeded with OP50 and grown at the restrictive temperature (25°C) in order to obtain sterile adults. Young adult animals were transferred to slow-killing plates (NGM agar containing 0.35% peptone), seeded with OP50, or tryptic soy agar plates (TSA, see below) seeded with RN6390. Animals were harvested at 8 h after transfer. Three independent replicates of each treatment were isolated. RNA isolation Total RNA was extracted using TRI Reagent (Molecular Research Center, http://www.mrcgene.com) according to the manufacturer's instructions, followed by purification on RNeasy columns (Qiagen, http://www1.qiagen.com). Microarray target preparation and hybridization for Affymetrix GeneChips RNA samples were prepared and hybridized to Affymetrix full-genome GeneChips for C. elegans at the Harvard Medical School Biopolymer Facility, according to instructions from Affymetrix (http://www.affymetrix.com). Briefly, 5 µg of total RNA was reverse transcribed using an oligo dT-T7 primer and Superscript II reverse transcriptase, followed by second-strand cDNA synthesis. The double-stranded cDNA was then purified using a DNA purification kit (Qiagen), and used as the template for in vitro transcription using T7 RNA polymerase and biotinylated nucleotides. The resulting cRNA was fragmented and hybridized onto C. elegans Affymetrix GeneChips as previously described [44]. Microarray analysis for S. aureus studies Affymetrix .cel files were uploaded into the Resolver Gene Expression Data Analysis System, version 5.1 (Rosetta Inpharmatics, http://www.rii.com) at the Harvard Center for Genomic Research for analysis. For each condition, three replicate microarrays were normalized and analyzed using the Resolver intensity error model for single color chips [89]. The two conditions were then compared in Resolver to determine fold change for each probe set and a p-value, using a modified t test. Genes with a 2-fold or greater fold change and a p-value <0.01 were considered differentially expressed. Differentially expressed probe sets were compared for S. aureus infection (this study), P. aeruginosa infection [44], and M. nematophilum infection [90] using Resolver. Significance of over-representation analysis Analysis of over-representation of GO annotation categories was performed using FuncAssociate (http://llama.med.harvard.edu/cgi/func/funcassociate). GO Bioset analysis was performed using NEXTBIO (www.nextbio.com) [91]. For greater inclusivity, gene expression changes greater than 1 were included in the NEXTBIO analysis, as suggested by NEXTBIO. Infection and lifespan assays All assays were conducted at 25°C, 65% relative humidity. Animals were scored as alive or dead by gentle prodding with a platinum wire. Kaplan-Meier statistical analyses were performed using the software Prism (GraphPad, http://www.graphpad.com). Survival data were compared as described using the log-rank test. Data are represented as median survival (MS) or lethal time – 50 (LT50) when MS values were skewed by small number of timepoints, N (number of deaths/censored), and p value. A p-value <0.05 was considered significantly different from control. S. aureus killing assays Assays were performed as described [17]. Briefly, NCTC8325 (or mutant derivatives, as noted) was grown overnight in tryptic soy broth (TSB, BD, Sparks, MD) with 10 µg/ml nalidixic acid (Sigma). 5–10 µl of overnight cultures diluted 1∶5 in fresh TSB were seeded on 35 mm tryptic soy agar (TSA, BD, Sparks, MD) plates with 10 µg/ml nalidixic acid. For accumulation experiments using GFP-S. aureus, plates were supplemented with 10 µg/ml chloramphenicol (Sigma) for plasmid maintenance. A total of 25–35 L4 stage hermaphrodites were transferred to each of three replicate plates per strain. Animals that died as a consequence of a bursting vulva or crawled off the agar were censored. Experiments were performed at least twice. For heat-killing of S. aureus, fresh overnight cultures of NCTC8325 were concentrated 10-fold and incubated at 95°C for 45 min. Following heat treatment, no live cells could be detected by plating undiluted cultures on TSA plates. P. aeruginosa slow-killing assays Briefly, PA14 was cultured in Luria broth (LB), seeded on slow-killing plates and incubated first for 24 h at 37°C and then for 24 h at 25°C. A total of 25–35 L4 stage hermaphrodites were transferred to each of three replicate plates per strain. Experiments were performed at least twice. M. nematophilum infection assays Assays were performed as described [92]. Briefly, OP50 and CBX102 were cultured overnight in LB, mixed at 1∶10 ratio of CBX102:OP50 and plated on NGM. A total of 25–35 L4 stage hermaphrodites were transferred to each of three replicate plates per strain. Assays were conducted at 25°C. For Dar quantification, animals were analyzed directly on infection plates, in triplicate. Lifespan assays Briefly, RNAi plates seeded with dsRNA-expressing E. coli HT115 were used. Approximately 100 synchronized eri-1(mg366) L1 larvae were added to each of 3 plates, and incubated for 24 h at 15°C followed by 24 h at 25°C, which causes animal sterility. 35–50 late L4 stage animals were added to each of 3 fresh RNAi plates seeded with dsRNA-expressing E. coli HT115, and incubated at 25°C. Lifespan is defined as the time elapsed from when animals were put on plates to when they were scored as dead. Experiments were performed at least twice. Animals that died of a bursting vulva or crawled off the agar were censored. Quantitative RT-PCR (qRT-PCR) analysis Animals were treated essentially as described for killing assays described above, with the following modifications. For S. aureus infection assays, infected samples were compared to parallel samples feeding on E. coli OP50, heat-killed by 30 min incubation at 95°C, plated on the same TSA medium. All strains compared were grown in parallel. Total RNA was extracted using TRI Reagent, and reverse transcribed using the Superscript III kit (Invitrogen). cDNA was subjected to qRT-PCR analysis using SYBR green detection (BIO-RAD SYBR Green supermix) on iCycler (Bio-Rad, http://www.bio-rad.com) and RealPlus (Eppendorf, Germany) machines. Primers for qRT-PCR were designed using Primer3Plus (Massachusetts Institute of Technology, http://www.bioinformatics.nl/cgi-bin/primer3plus/primer3plus.cgi), checked for specificity against the C. elegans genome and tested for efficiency with a dilution series of template. Primer sequences are available upon request. All Ct values are normalized against the control gene snb-1, which did not vary under conditions being tested. Fold change was calculated using the Pfaffl method [93]. We found some variability in gene induction levels from experiment to experiment. The source of this variation has not been conclusively ascertained; however, we suspect it may derive from differences between batches of agar plates used for infection assays. Importantly, all experiments were repeated at least twice (biological replicates) and were internally controlled. Additionally, despite numerical variability in fold induction, all results were internally consistent. GFP fusions PCR primers to amplify 1665 bp of sequence upstream of the clec-60 start site, 3505 bp upstream of the clec-70 start site, 1774 bp upstream of the fmo-2 start site, and 913 bp upstream of the ilys-3 start site were designed using the online PCR primer design tool provided by the British Columbia Genome Sciences Center (http://elegans.bcgsc.bc.ca/promoter_primers/index.html). Splicing by overlapping extension PCR (SOE-PCR) was used as described [94] to generate promoter-GFP fusion PCR fragments, which were transformed at 3 ng/ µl into wild type animals by microinjection with 40 ng/ µl of a myo-2::NLS::mCherry construct as coinjection marker used to identify transgenic animals (courtesy of J. Kaplan, Massachusetts General Hospital). Primer sequences are available upon request. GFP fusion induction L4 animals carrying extrachromosomal arrays were transferred from NGM plates seeded with OP50-1 to S. aureus, P. aeruginosa, or M. nematophilum killing plates essentially as described above. After incubation, animals were mounted on glass slides with 2% agarose pads, anesthetized with 30 mM NaN3, and immediately used for imaging. Exposure times were set for the most highly expressed condition and kept constant throughout each experiment. RNAi knockdown RNAi screen Enhanced RNAi eri-1(mg366) mutants were propagated at 15°C. RNAi of selected genes was carried out in triplicate using bacterial feeding RNAi [95]. Synchronized L1 animals were transferred to RNAi plates, incubated at 15°C for 25 h and then 25°C for 24 h to induce sterility, and then transferred to NCTC8325-seeded killing assays. The screen was performed once, and positive clones that exhibited altered survival on S. aureus were tested at least once more. RNAi clones were obtained from the Ahringer laboratory (except clec-70 RNAi, which was made by recombining pDONR201.clec-70 from the ORFeome library [96] with pDEST.L4440 [97]. Sequences of positive RNAi clones were confirmed. cdc-25.1 RNAi To sterilize worms previous to use in killing assays, cdc-25.1 RNAi was carried out by feeding L4 animals for 24 h at 15°C. PMN expression analysis Gene expression microarray data files of S. aureus infection were obtained from Gene Expression Omnibus (GEO accession: GSE2405) and analyzed. The samples were derived from human polymorphonuclear leukocytes (PMNs) from three healthy donors using a separate HU133A GeneChip (Affymetrix) for each donor [74]. We examined the data from PMNs that were either uninfected or infected with live S. aureus for 9 hours. The dataset was MAS5.0-normalized and filtered by excluding probe sets with 100% ‘absent’ calls (MAS5.0 algorithm) across all samples. FDR analysis (q-value<0.005) with 1000 permutations using significance analysis of microarrays [98] was performed to identify genes that were differentially induced in S. aureus-infected PMNs versus uninfected controls. Epifluorescence microscopy Images were acquired using a Zeiss AXIO Imager Z1 microscope with an Zeiss AxioCam HRm camera and Axiovision 4.6 (Zeiss) software. Image cropping and minimal manipulation were performed using Photoshop (Adobe). Gene accession information Gene Public Name, Gene WormBase ID, Source GenBank ID, Gene CGC Name; bar-1, WBGene00000238, U46673, bar-1; col-63, WBGene00000639, Z81143, col-63; col-98, WBGene00000673, Z81503, col-98; cpr-2, WBGene00000782, Z81531, cpr-2; egl-5, WBGene00001174, L15201, egl-5; exc-5, WBGene00001366, Z68159, exc-5; fmo-2, WBGene00001477, Z70286, fmo-2; ins-11, WBGene00002094, U41279, ins-11; lys-2, WBGene00003091, AL021479, lys-2; lys-5, WBGene00003094, Z73427, lys-5; mpk-1, WBGene00003401, Z46937, mpk-1; pmk-1, WBGene00004055, U58752, pmk-1; sod-3, WBGene00004932, U42844, sod-3; tol-1, WBGene00006593, AF348166, tol-1; unc-32, WBGene00006768, Z11115, unc-32; C32H11.1, WBGene00007864, NM_070062.2; C50F4.9, WBGene00008234, Z70750; F01D5.2, WBGene00008493, Z81493; acs-2, WBGene00009221, Z81071, acs-2; F55G11.2, WBGene00010123, Z82272; clec-60, WBGene00014046, Z49132, clec-60; clec-61, WBGene00014047, Z49132, clec-61; clec-52, WBGene00015052, U58752, clec-52; C23G10.6, WBGene00016013, U39851; C30G12.2, WBGene00016274, U21319; ilys-3, WBGene00016670, AF067611, ilys-3; C49G7.5, WBGene00016783, AF016418; acdh-1, WBGene00016943, AC006625, acdh-1; F49F1.6, WBGene00018646, AF100656; F53A9.8, WBGene00018731, U23523; F53E10.4, WBGene00018760, U88177; clec-70, WBGene00021581, AC024785, clec-70; clec-71, WBGene00021582, AC024785, clec-71; Y65B4BR.1, WBGene00022040, AC024847. Supporting Information Figure S1 P. aeruginosa makes putative outer membrane vesicles (OMVs), disrupts the brush border, and penetrates the epithelial barrier. A–D. TEM micrographs of P. aeruginosa-infected animals after 48 h infection. Scale bars, 0.5 µm. A. Detail of intestinal lumen filled with OMVs and bacterial cells (b), and brush border (mv) coated with extracellular material (em). Note disruption of the microvilli (black asterisk) and OMV shedding from the bacterial cells (black arrowheads). At this time point the terminal web (tw) appears whole. B. High magnification TEM showing apparent OMV shedding off bacterial cells (b, indicated with black arrowheads). C. Example of distal dissemination of P. aeruginosa. A bacterial cell (b) is shown between the body-wall muscle (bwm) and the cuticle (cu), which is the exoskeleton of the animal. D. Detail of animal infected with gacA mutant P. aeruginosa. The bacteria (b) appear less rugose than their wild-type counterparts. There is much less microvillar pathology (mv) and extracellular material (em). The terminal web is unaffected (tw). We also find evidence of exocytosis (vesicles labelled with asterisks). (8.19 MB TIF) Click here for additional data file. Figure S2 Wild type, but not gacA mutant, P. aeruginosa causes increased early autophagosomes. fer-15;fem-1 sterile animals were infected with wild type or gacA mutant P. aeruginosa PA14 for 24 h. Autophagosomes (Fig. 1H–I) were counted in TEM transversal sections of both intestinal epithelial cells, and are represented as means of n = 6 different sections each. Error bars are SEM. ***p<0.0001(Two-tailed t test). (1.56 MB TIF) Click here for additional data file. Figure S3 Early intestinal accumulation of S. aureus. A, B, C. High magnification micrographs of a representative animal infected with GFP-expressing S. aureus 4 h after initiation of infection. Accumulation in pharyngeal-intestinal valve and foregut (A), midgut (B), and rectum (C). Arrowheads indicate areas magnified in insets to illustrate the faint green haze likely due to bacterial cell lysis (A, C) and bacterial attachment to the apical surfaces of enterocytes (B). Green, GFP-S. aureus. Red, autofluorescent granules. Distention of the anterior intestinal lumen immediately adjacent to the pharyngeal-intestinal valve was apparent (A). There was less accumulation of bacteria in the mid section of the intestinal lumen. The bacteria appear to attach to the apical surface of the intestinal cells, as well as each other to a thickness of 3–4 bacterial cell diameters, either due to the dumbbell-shaped intestinal lumen or to direct bacteria-enterocyte and bacteria-bacteria interactions (B). (4.07 MB TIF) Click here for additional data file. Figure S4 S. aureus accumulates in the intestine and causes anal swelling in C. elegans. A, B. Timecourse of S. aureus intestinal accumulation. A, representative epifluorescence (left) and Nomarski (right) micrographs of animals infected with GFP-expressing S. aureus, illustrating three categories of intestinal accumulation observed. a, p, indicate anterior and posterior ends respectively. B, quantification of intestinal accumulation of S. aureus (SA) and P. aeruginosa (PA) at different times. At early times, green haze from lysed bacteria could be misinterpreted as P. aeruginosa accumulation if evaluated at low magnification. N≥18 animals for each condition. C, D, E. Deformed anal region (Dar) phenotype during S. aureus infection. a, p, indicate anterior and posterior ends respectively. C. Nomarski micrograph showing a representative animal infected with S. aureus for 12 h. Note smaller size than uninfected animal shown in D. Inset, higher magnification of anal region, highlighting swelling (arrow). D. Micrograph of an uninfected animal. Note smooth tapering of the tail region. Inset, detail of anal region, noting the absence of swelling (arrow). Micrographs in C, D are at same magnification. E. Quantification of the Dar phenotype in S. aureus-infected animals, compared with animals feeding on heat-killed S. aureus. Error bars represent standard deviation. N = 82 (live bacteria), N = 97 (heat killed bacteria). F, G, H, I, J, K. Nomarski micrographs illustrating the Dar phenotype after 12 h of S. aureus infection in wild type (F), egl-5 (H), pmk-1 (I), and unc-32 (J) mutant animals in contrast to non-Dar bar-1 (G) and mpk-1,unc-32 (K) mutants. L. Quantification of Dar phenotype in wild type (N = 93), bar-1 (N = 69), egl-5 (N = 89), pmk-1 (N = 87), unc-32 (107), and mpk-1,unc-32 (N = 83) animals. ***, p<0.001 (two-tailed one-sample t test). (8.78 MB TIF) Click here for additional data file. Figure S5 Complete destruction of internal structure in S. aureus-killed animal. TEM micrograph of a transversal midbody section of a S. aureus-killed animal, after 36 h infection. The only remaining internal structure is an unidentified circular remnant (lower right). Scale bar, 2 µm. (8.72 MB TIF) Click here for additional data file. Figure S6 α-hemolysin-independent membrane blebbing and cell lysis. TEM micrographs of transversal sections of S. aureus-infected animals after 12 h infection. Red false-coloring indicates membrane blebbing and microvillus shortening, by highlighting the apical surface of the intestinal epithelial cells from the underlying terminal web to the end of the microvilli and the membrane blebs in the intestinal lumen. A. Section of intestinal ring 1, showing four intestinal epithelial cells (iec). Two apical junctions are visible (aj). B. Midbody section, showing two intestinal epithelial cells. One apical junction is visible (lower center). Asterisks indicate extensive host cell membrane blebbing. Scale bars, 0.5 µm. C. Cross-section of an animal infected with wild type S. aureus for 24 h. The box indicates section magnified in D. Scale bar, 10 µm. D. Detail of animal infected with wild type S. aureus. Scale bar, 2 µm. E. Cross-section of animal infected with α-hemolysin-defective Δhla mutant S. aureus. The box indicates section magnified in F. Scale bar, 10 µm. F. Detail of animal infected with Δhla mutant S. aureus. Note the upper intestinal cell is not yet lysed, whereas the lower cell is clearly lysed (indicated) and invaded by live bacteria (indicated). Also, microvillar shortening, membrane blebbing, and intestinal cell volume loss were indistinguishable from wild type at this timepoint. Scale bar, 2 µm. iec, intestinal epithelial cell; tw, terminal web; mv, microvilli. (9.89 MB TIF) Click here for additional data file. Figure S7 S. aureus hemolysins are dispensable for C. elegans killing. spe-9;fer-15 sterile animals were infected with triple hemolysin Δhla Δhlb Δhlg mutant RN6390 S. aureus, or with the double mutant combinations. All killed C. elegans with similar kinetics. (7.35 MB TIF) Click here for additional data file. Figure S8 Extraintestinal sites of host response gene expression. ilys-3::GFP expression in pharynx and unidentified cell near terminal bulb (arrow, A) and vulval cells (B). One transgenic line had clec-70::GFP expression in unidentified head cells (arrows, C), vulval and uterine muscles (D), rectal gland cells (arrow, E), and anal depressor muscle (F). F53A9.8::GFP expressed in rectal gland cells in animals feeding on E. coli (arrows, G) and on S. aureus for 24 h (arrows, H). (8.58 MB TIF) Click here for additional data file. Figure S9 mpk-1/ERK is dispensable for the intestinal host response. Transcript levels were measured in synchronized young adult animals feeding on heat-killed E. coli or infected with S. aureus for 8 h. Data are the means of two biological replicates, each replicate measured in duplicate and normalized to a control gene, expressed as the ratio of the corresponding S. aureus-induced levels and the basal E. coli levels. Error bars are SEM. (3.22 MB TIF) Click here for additional data file. Figure S10 tol-1/TLR is dispensable for host survival of S. aureus infection. tol-1(nr2033) mutants exhibit the same susceptibility to S. aureus-mediated killing as wild type. Animals were sterilized with cdc-25 RNAi previous to killing assays (see Experimental Procedures). Wild type (LT50 = 75.6 h; N = 101), tol-1 (LT50 = 75.35 h; N = 117; p = 0.8814). (5.78 MB TIF) Click here for additional data file. Figure S11 Pathogen-specific induction of infection reporters. A, B, C, D. Animals carrying clec-60::gfp arrays were infected with pathogens for 24 h, in parallel with non-pathogenic E. coli control (A). Induction of clec-60::gfp by infection with M. nematophilum (C) and S. aureus (D), and repression by infection with P. aeruginosa (B). Note vulval expression in B and C (arrows). E, F, G, H. Animals carrying F53A9.8::gfp were infected with pathogens for 24 h, in parallel with non-pathogenic E. coli control (E). Induction of F53A9.8::gfp by infection with M. nematophilum (G), S. aureus (H), and P. aeruginosa (F); the levels of induction on S. aureus were highest. On non-pathogenic E. coli, clec-60::GFP was expressed at low levels, mostly in the 9th ring of intestinal epithelial cells (Fig. S7A), and F53A9.8::GFP was weakly expressed mostly in the posterior intestine and the rectal gland cells (Fig. S7E). During infection with M. nematophilum, clec-60::GFP was expressed weakly in the intestine, as well as occasional expression in the vulva, consistent with previous reports (Fig. S7C, [90]). M. nematophilum induced moderate levels of F53A9.8::GFP expression in the intestine (Fig. S7G). During infection with P. aeruginosa, we observed reduced expression of clec-60::GFP in the intestine, below the level observed on E. coli, except for occasional expression in the vulva (Fig. S7B), and induced expression of F53A9.8::GFP in the intestine (Fig. S7F). Finally, during infection with S. aureus we observed highest expression of both reporters, mostly in the posterior half of the intestine (Fig. S7D, H). clec-52::GFP was also downregulated on P. aeruginosa (not shown). (6.29 MB TIF) Click here for additional data file. Figure S12 clec-60,61/CTL overexpression is protective during S. aureus infection. Transgenic animals carrying clec-60,61 cluster extrachromosomal arrays survived longer (LT50 = 54.2 h; N = 79; p = 0.019) during S. aureus infection than control animals bearing arrays composed of coinjection marker and clec-60::GFP promoter fusion (LT50 = 47.3 h; N = 95). (6.37 MB TIF) Click here for additional data file. Table S1 List of genes whose expression changed more than two-fold, after 8 h of infection with S. aureus RN6390. Threshold p≤0.01 for significance. Public names in bold indicate genes selected for qRT-PCR analysis. (0.09 MB XLS) Click here for additional data file. Table S2 Nextbio biogroups, defined by GO annotations, that are over-represented among expression changes 8 h after infection with S. aureus (SA, a) or P. aeruginosa (PA, b). Key in a: green genes have same direction on SA and PA; red genes are SA down, PA up; bold black genes are SA up and PA down (or no change on PA). (0.07 MB XLS) Click here for additional data file. Table S3 List of genes whose expression changed during infection with S. aureus and during exposure to B. thuringiensis PFT Cry5B. (0.03 MB XLS) Click here for additional data file. Table S4 List of shared gene classes upregulated during infection of C. elegans and human neutrophils with S. aureus. (0.10 MB DOC) Click here for additional data file. Table S5 List of genes whose expression changed during infection with S. aureus, and during infection with (a) Enterococcus faecalis, (b) L. chromiireducens, and (c) common late response genes. (0.02 MB XLS) Click here for additional data file. Table S6 A. List of C. elegans strains used in this study. B. List of bacterial strains used in this study. (0.06 MB DOC) Click here for additional data file. Table S7 Over-represented GO annotations among gene expression changes during S. aureus or P. aeruginosa infection for 8 h. “Output is in the form of “Rank, N, M, X, LOD, P, P-adj, GO Attribute”, where Rank: position in the attribute list ranked by significance of association with query; N: number of genes in the most surprising subquery with this attribute; M: size of most surprising sub query; X: number of genes overall with this attribute; LOD: the logarithm (base 10) of the odds ratio; positive values indicate over-representation; P: single hypothesis one-sided P-value of the association between attribute and query (based on Fisher's Exact Test); P-adj: adjusted P-value: fraction (as a %) of 1000 null-hypothesis simulations having attributes with this single-hypothesis P value or smaller.” From FuncAssociate (http://llama.med.harvard.edu/cgi/func/funcassociate). (0.06 MB DOC) Click here for additional data file. Text S1 Supporting text. (0.02 MB DOC) Click here for additional data file.

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Rita JM Volkers

L Basten Snoek

Caspara J van Hellenberg Hubar

Renata Coopman

Wei Chen

Wentao Yang

Mark G Sterken

Hinrich Schulenburg

Bart P Braeckman

Jan E Kammenga

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Journal ID (nlm-ta): BMC Biol

Journal ID (iso-abbrev): BMC Biol

Title: BMC Biology

Publisher: BioMed Central

ISSN (Electronic): 1741-7007

Publication date Collection: 2013

Publication date (Electronic): 19 August 2013

Volume: 11

Page: 93

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[1 ]Laboratory of Nematology, Wageningen University, Droevendaalsesteeg 1, Wageningen 6708PB, The Netherlands

[2 ]Biology Department, Ghent University, Proeftuinstraat 86 N1, B-9000 Gent, Belgium

[3 ]Department of Evolutionary Ecology and Genetics, Zoological Institute, Christian Albrechts-Universitaet zu Kiel, Am Botanischen Garten 1-9, Kiel 24118, Germany

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Publisher ID: 1741-7007-11-93

DOI: 10.1186/1741-7007-11-93

PMC ID: 3846632

PubMed ID: 23957880

SO-VID: 89e1dab2-fe8b-4811-b221-fcb626f68499

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This is an Open Access article distributed under the terms of the Creative Commons Attribution License ( http://creativecommons.org/licenses/by/2.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Gene-environment and protein-degradation signatures characterize genomic and phenotypic diversity in wild Caenorhabditis elegans populations

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