Research Article
16 December 2014

Genome-Wide Evaluation of the Interplay between Caenorhabditis elegans and Yersinia pseudotuberculosis during In Vivo Biofilm Formation


The formation of an incapacitating biofilm on Caenorhabditis elegans by Yersinia pseudotuberculosis represents a tractable model for investigating the genetic basis for host-pathogen interplay during the biofilm-mediated infection of a living surface. Previously we established a role for quorum sensing (QS) and the master motility regulator, FlhDC, in biofilm formation by Y. pseudotuberculosis on C. elegans. To obtain further genome-wide insights, we used transcriptomic analysis to obtain comparative information on C. elegans in the presence and absence of biofilm and on wild-type Y. pseudotuberculosis and Y. pseudotuberculosis QS mutants. Infection of C. elegans with the wild-type Y. pseudotuberculosis resulted in the differential regulation of numerous genes, including a distinct subset of nematode C-lectin (clec) and fatty acid desaturase (fat) genes. Evaluation of the corresponding C. elegans clec-49 and fat-3 deletion mutants showed delayed biofilm formation and abolished biofilm formation, respectively. Transcriptomic analysis of Y. pseudotuberculosis revealed that genes located in both of the histidine utilization (hut) operons were upregulated in both QS and flhDC mutants. In addition, mutation of the regulatory gene hutC resulted in the loss of biofilm, increased expression of flhDC, and enhanced swimming motility. These data are consistent with the existence of a regulatory cascade in which the Hut pathway links QS and flhDC. This work also indicates that biofilm formation by Y. pseudotuberculosis on C. elegans is an interactive process during which the initial attachment/recognition of Yersinia to/by C. elegans is followed by bacterial growth and biofilm formation.


Bacterial survival in clinical, environmental, and industrial settings is enhanced by the formation of biofilms. Biofilms are structured populations in which bacterial cells attach to surfaces and/or other bacteria via a self-generated, hydrated extracellular matrix (ECM) incorporating exopolysaccharides, DNA, and proteins (1). Following surface attachment, bacterial cells form microcolonies, which mature before releasing individual cells or groups of cells that seek out new surfaces for further colonization. Bacteria in biofilms are notoriously resistant to antimicrobial agents as well as being resilient and highly refractive to a variety of host and clinical challenges, as exemplified by Pseudomonas aeruginosa infections in individuals with cystic fibrosis (2). Although there is extensive literature on biofilm formation on abiotic (nonliving) surfaces, including studies of their developmental control at transcriptional, translational, and posttranslational levels (39), there is a paucity of information on the genetic control of biofilm formation on biotic (living) surfaces that are frequently important in establishing infection in higher organisms.
A simple model of biofilm development on biotic surfaces is therefore useful if we are to unravel the intricate regulatory interplay that must exist between host and pathogen during biofilm-centered infections so that new methods for eradicating such biofilms can be developed. Yersinia pseudotuberculosis, Yersinia pestis, and Xenorhabdus nematophila form biofilms around the head of the nematode Caenorhabditis elegans (1012). Since C. elegans has been thoroughly studied at the genetic level and orthologous genes have been frequently studied in human health and disease, the C. elegans bacterial biofilm model can be used to identify genetic features of both the pathogen and host that contribute to biofilm-mediated interactions between bacteria and invertebrates. Previous studies have revealed that both host-specific and pathogen-specific factors are important in successful biofilm formation. For example, Y. pseudotuberculosis does not form a biofilm on C. elegans surface antigen mutants (srf-2, srf-3, and srf-5 mutants) (12), and these mutants are also more prone to trapping by nematophagous fungi (13). The srf-3 gene encodes a nucleotide sugar transporter (14), but to date, srf-2 and srf-5 remain undefined. Biofilm formation by Y. pseudotuberculosis is abrogated in three other C. elegans mutants termed bah (named bah for biofilm absent on head) (15). bah-1 codes for a DUF23 protein and is expressed in seam cells which are among the hypodermal cells that synthesize the cuticle (16); the bah-2 and bah-3 genes remain undefined.
When biofilms formed by Y. pseudotuberculosis on C. elegans are examined by fluorescence microscopy, bacterial cells can be observed embedded in a matrix containing both an N-acetylglucosamine GlcNAc-containing exopolysaccharide and DNA (17). Mutation of the hmsHFRS operon which is responsible for the production of a GlcNAc-containing exopolysaccharide in Y. pseudotuberculosis, Y. pestis, and X. nematophila renders all three bacterial species unable to form biofilms on C. elegans, consistent with a requirement for a bacterial determinant(s) in attachment/biofilm formation (11). In addition, while Y. pseudotuberculosis strains with a functional PhoPQ system are unable to form biofilms, mutation of the two-component PhoPQ system restores the ability to form a biofilm (18). Moreover, Y. pseudotuberculosis biofilm development on C. elegans is under quorum sensing (QS) control, i.e., coordinated at the population level through the production and sensing of diffusible signal molecule(s), in this case N-acylhomoserine lactones (AHLs). These signal molecules can be detected in situ in Y. pseudotuberculosis biofilms on C. elegans (17). Y. pseudotuberculosis contains two pairs of LuxRI orthologues termed YpsRI and YtbRI (19, 20) responsible for the production (YpsI and YtbI) and sensing (YpsR and YtbR) of four major AHLs. This QS system is organized hierarchically with YpsR and its cognate AHLs regulating ytbR and ytbI as well as ypsR and ypsI gene expression (19). Biofilm formation on C. elegans can be attenuated either by expressing an AHL-degrading enzyme in Y. pseudotuberculosis or by mutating both ypsI and ytbI (17). Furthermore, although neither flagellum-mediated motility nor flagella are required for biofilm formation, Y. pseudotuberculosis strains carrying mutations in either of the motility regulatory genes, flhDC and fliA or the flagellin export gene flhA are attenuated for biofilm formation. The mechanism involved appears to depend on the Yersinia type III secretion system (T3SS) carried on the virulence plasmid pYV, which is repressed via the QS-dependent control of the motility master regulator FlhDC, since biofilm formation can be restored in the Y. pseudotuberculosis ypsI ytbI double mutant when the strain is cured of pYV (17). These observations are independent of the chromosomally encoded T3SS.
Thus, is it clear that biofilm formation on C. elegans is an interactive process depending on both pathogen and host factors. Tan and Darby (21) postulated that biofilms form and develop on C. elegans by passive attachment of sticky bacterial material as the worm moves through the bacterial lawn. While the motility of C. elegans srf and bah mutants, which move through the bacterial lawn but fail to accumulate biofilms, implies a role for specific worm components, it is not yet clear whether biofilm formation is initiated through the attachment of individual bacterial cells or a larger biomass to C. elegans from the agar surface. Consequently, we chose a time point shortly after exposure to the biofilm-forming bacteria (1 h) when transcriptomic responses are likely to be specific to the infection. We sought to determine the following: (i) whether bacteria or biofilm attaches first to C. elegans; (ii) whether biofilm formation is a dynamic process during which C. elegans exhibits a differential transcriptomic response to Yersinia strains which vary in their ability to form a biofilm; and (iii) the comparative transcriptomic differences between the Y. pseudotuberculosis parent and QS mutants to identify additional genes that may contribute to biofilm development on C. elegans. From selected nematode and bacterial genes identified as differentially expressed in the microarrays, the corresponding mutants were examined for their biofilm phenotype. From the results obtained, we suggest that biofilm formation by Yersinia on C. elegans is a dynamic two-stage process involving attachment of bacteria to the worm surface followed by the aggregation of bacteria via QS and secretion of an extracellular matrix to form a mature biofilm. Furthermore, our transcriptomic experimental results revealed that QS regulates genes within the Yersinia histidine utilization pathway. Mutation of hutC resulted in the upregulation of flhDC, increased motility, and the abrogation of biofilm on C. elegans, indicating that the hut pathway provides a regulatory link between quorum sensing and FlhDC.


Strains and growth conditions.

The wild-type Y. pseudotuberculosis strains 3384 and YPIII together with the isogenic YPIII mutants used in this study were grown in LB (with shaking) or on LB agar with appropriate antibiotics at 28°C; swimming motility assays were performed on semisolid motility agar plates as previously described (19). For sources of Yersinia strains, mutants, and vector used in the study, see Tables S4 and S5 in the supplemental material. The hutC YPIII mutant was cured of the pYV plasmid as described previously (17). C. elegans N2 and mutants were obtained from the Caenorhabditis Genetics Center (CGC; University of Minnesota) and maintained on NGM agar (22).

C. elegans biofilm assays.

For biofilm assays, 400-μl portions of an overnight culture of Y. pseudotuberculosis YPIII were seeded onto NGM agar, allowed to dry, and kept overnight at room temperature. Twenty larval stage four (L4) C. elegans were picked or pipetted onto the bacterial lawn, and biofilm development was examined over 24 h for the onset of biofilm formation, size of biofilm, and changes in movement. After the worm biofilms were washed three times in M9 buffer, they were stained with Texas Red-wheat germ agglutinin (Molecular Probes) at 10 μg · ml−1 for 30 min, washed again as described above, and viewed using a Zeiss Axioplan 2 microscope.
The level of biofilm accumulation on C. elegans was denoted as the biofilm severity incidence and was calculated by the method of Tarr (23) and as previously described (17): biofilm severity incidence = Σ(level X number of samples in this level)/(highest level X total sample numbers) × 100%, where the level of biofilm accumulation ranges from 0 (no biofilm) to 3 (severe biofilm). All assays in which the level of biofilm severity was assessed were carried out with a minimum of 20 to 30 worms performed in a double-blind manner with at least three or four replicates, and each experiment was performed more than once.

Nucleic acid manipulations.

Plasmid minipreparation, PCR, restriction, and ligation were carried out as previously described (20). Chromosomal DNA was extracted from 10-ml overnight cultures of Y. pseudotuberculosis following the manufacturer's instructions (Gentra; Puregene genomic DNA purification kit). RNA was extracted from 3 ml of log-phase culture (optical density at 600 nm [OD600] of ∼0.8) after mixing with 6 ml RNA Protect (Qiagen) using an RNeasy kit (Qiagen). RNA was extracted from C. elegans as described previously (24). Briefly, worms with biofilm (fed on Y. pseudotuberculosis YPIII for 1 h or 24 h) or without biofilm (fed on Y. pseudotuberculosis 3384 for 1 h or 24 h) or on the Y. pseudotuberculosis YPIII ypsI ytbI double mutant (for 1 h) were washed from plates with M9 buffer (22), suspended in 3 volumes of TriReagent (Sigma), frozen in liquid nitrogen, and ground in a pestle and mortar. After thawing, material was disrupted with a Dounce homogenizer; extracted with chloroform, and precipitated with isopropanol. Contaminant DNA was removed from RNA extractions using Turbo DNase (Ambion), and RNAs were analyzed using an Agilent 2100 bioanalyzer (Agilent) and Nanodrop ND 1000 spectrophotometer (Nanodrop Technologies).

Transcriptome analysis.

Yersinia transcriptome analysis was performed using the BμG@S ( Y. pestis microarray (v1.1.0). Total RNA (in triplicate) was isolated from mid-log-phase cultures of Y. pseudotuberculosis YPIII grown at 28°C as well as the isogenic flhDC, ytbI, and ypsR single mutants and ypsI ytbI and ypsR ytbR double mutants. Briefly, an RNA versus DNA protocol was followed. In each case, paired microarrays were hybridized, one with 10 μg wild-type Cy5-labeled RNA plus 1 μg Cy3-labeled Y. pseudotuberculosis YPIII genomic DNA and one with 10 μg mutant Cy5-labeled RNA plus 1 μg Cy3-labeled Y. pseudotuberculosis YPIII DNA. Microarrays were hybridized overnight at 65°C, washed, and scanned using a GMS 418 microarray scanner. Data were quantified using ImaGene 9.0 (Biodiscovery) and analyzed using GeneSpring GX v7.3 (Agilent Technologies). Three biological replicates were performed for each mutant. Wild-type Y. pseudotuberculosis YPIII was hybridized in each paired microarray experiment with each mutant; therefore, a total of 25 microarrays were employed. Results are given showing the relative up- or downregulation for genes with a difference of twofold or more after an analysis of variance (ANOVA) with multiple testing corrections (Benjamini-Hochberg false discovery rate [FDR]). Microarrays comparing gene expression between wild type and ypsI and ytbR mutants showed no statistical differential regulation at this level (P = 0.001 with Benjamini-Hochberg FDR) of statistical stringency.
C. elegans arrays were performed using Affymetrix Genechip genome microarrays (catalog no. 900383), following the manufacturer's instructions, washed using an Affymetrix Genechip 450 fluidics station, scanned with an Affymetrix Genechip scanner and analyzed using GeneSpring GX v7.3. Three biological replicates were performed for C. elegans exposed for 1 h to Y. pseudotuberculosis YPIII for 1 h (with biofilm) YPIII or Y. pseudotuberculosis 3384 (without biofilm). Similarly, C. elegans was exposed to Y. pseudotuberculosis YPIII (with biofilm) or Y. pseudotuberculosis YPIII ypsI ytbI (without biofilm). Results are presented as the relative up- or downregulation for genes with a difference of twofold or more after analyzing expression and using confidence limits with multiple testing corrections (Benjamini-Hochberg FDR). Genes identified in microarrays were specified from Affymetrix identifiers and analyzed as to function using Wormbase version WS238.

Real-time PCR (qPCR).

Quantitative PCR (qPCR) was performed on selected Y. pseudotuberculosis genes using TaqMan gene expression assays in a two-step procedure. cDNAs were made using the same RNA preparations as were used in microarrays. The housekeeping gene used as control was gyrB. Relative quantification was assessed by the ΔΔCT method (2(−ΔΔCT)), and P values were calculated by t test. The wild type was used as the calibrator to assess fold change in mutant gene expression.

Construction of Y. pseudotuberculosis hutC mutant, flhDC′::lux and fliA′::lux promoter fusions.

A Y. pseudotuberculosis hutC deletion insertion mutant was constructed using a modified λ red recombinase one-step inactivation method based on the method of Datsenko and Wanner (25). Briefly, the PCR primers hutCF (CAATCAACGGTTTTACAATTGAGCGCAATGGACGACATTCCTGACGAAAGCCACGTTGTGTCTAA) and hutCR (CACCGTACCTTGTGGGGTAAAACGACCATAGAGCTGATAACGAGGGCCTGGTTAGAAAAACTCATCGAGCAT) incorporated 20 bp of the up- or downstream region of hutC and 50 bp of the kanamycin cassette from pUC4K (Pharmacia). pUC4K was used as a template for PCR, and the resulting fragment was transformed into Y. pseudotuberculosis YPIII containing the helper plasmid pAJD434. Following selection on the relevant antibiotics and the subsequent removal of pAJD434 by growth at 37°C, the hutC mutant carried the kanamycin resistance cassette in place of 648 bp of hutC. The resulting recombinants were confirmed by PCR and Southern blot analysis (data not shown).
The expression of flhDC and fliA was investigated in the Y. pseudotuberculosis hutC mutant background by first introducing a single copy of either pSA200 or pSA208, bioluminescent reporters which carry the flhDC or fliA promoter fused to luxCDABE, respectively, onto the chromosome. The method of construction used was outlined by Atkinson et al. (19). For each strain, bioluminescence was determined as a function of growth using a combined spectrophotometer/luminometer as previously described (19).

Scanning electron microscopy (SEM).

Worms were washed in M9 buffer and transferred to a lawn of Y. pseudotuberculosis YPIII. After 30 min or 24 h, they were fixed in 2.5% paraformaldehyde–2.5% glutaraldehyde–0.1 M sodium cacodylate buffer (pH 7.4), washed in 0.1 M sodium cacodylate buffer (pH 7.4) for 90 min, and postfixed in 1% OsO4–Milli-Q water for 90 min. After the worms were washed, they were stored overnight at 4°C in Milli-Q water prior to dehydration through a series of ethanol solutions (30%, 50%, 70%, 90%, and 100% ethanol [two 100% ethanol solutions]) for 15 min per dilution. The worms were placed on an SEM aluminum stub, air dried from the ethanol solutions, and sputter coated (metal plated) for 1.5 min. Samples were examined on a JEOL JSM25 III scanning electron microscope, and digital images were recorded using the Orion version 6 high-resolution, 32-bit PCI image-grabbing system.

Microarray data accession number.

The complete microarray data set for the experiments studying the impact of Y. pseudotuberculosis on the C. elegans transcriptome in the presence and absence of biofilm is available at ArrayExpress under accession number E-MEXP-2981.


Initial stages of biofilm formation reveal bacterial attachment to C. elegans.

During the early stages of infection, no biofilm is visible on C. elegans migrating over a lawn of Y. pseudotuberculosis strain YPIII via low-power light microscopy even though the worms exhibit the exaggerated body bends characteristic of worms with a biofilm. We therefore used scanning electron microscopy (SEM) to examine the worms early in infection (30 min after exposure to Y. pseudotuberculosis YPIII) to ascertain whether any attached bacterial cells were present and again after 24-h exposure to the bacteria. Under low magnification after 30 min of exposure to the bacteria, no obvious biofilm is apparent, although the worms move with exaggerated body bends (data not shown). However, under higher magnification (Fig. 1A), low numbers of individual bacterial cells (Fig. 1, white arrows) can be seen attached to the anterior end of the worm as well as identifiable bacterial cells that appear to be covered with extracellular matrix (ECM). At this stage of infection, there is insufficient biofilm to mechanically cause aberrant movement, but worms move with exaggerated body bends. After 24 h, there is a mass of biofilm over the anterior of the worm, and a smaller number of bacteria and ECM over the body of the worm (Fig. 1B). In contrast, Y. pseudotuberculosis strain 3384 does not induce exaggerated body bends (data not shown) nor attach to C. elegans and form a biofilm after 30-min exposure (Fig. 1C) or 24-h exposure (Fig. 1D). These data suggest that biofilm-forming Yersinia strains, such as YPIII, attach to the worm prior to the development of the biofilm rather than worms acquiring biofilm from the bacterial lawn.
FIG 1 Attachment of Y. pseudotuberculosis to C. elegans. (A to F) Scanning electron micrographs of C. elegans after 30-min (A, C, and E) and 24-h (B, D, and F) exposure to Y. pseudotuberculosis (bars, 10 μm). (A and B) N2 wild-type worms after exposure to Y. pseudotuberculosis YPIII. After 30-min exposure (A), individual bacteria can be seen attached to the worm (white arrows), as well as extracellular matrix (ECM) After 24-h exposure (B), a mass of biofilm is seen on the anterior of worm. (C and D) No attached bacteria, ECM, or biofilm on wild-type C. elegans N2 when exposed to Y. pseudotuberculosis 3384 (non-biofilm-forming strain) for 30 min (C) and few attached bacteria after 24 h (D). (E and F) No attached bacteria, ECM, or biofilm on the C. elegans Δfat-3 mutant after 30-min exposure (E) and few attached bacteria after 24-h exposure (F).

Impact of Y. pseudotuberculosis on the C. elegans transcriptome in the presence and absence of biofilm.

To obtain genome-wide insights into the response of C. elegans to Y. pseudotuberculosis, the transcriptome of C. elegans after 1-h exposure to Y. pseudotuberculosis YPIII was compared with those of C. elegans fed on either Y. pseudotuberculosis strain 3384 or the Y. pseudotuberculosis YPIII ypsI ytbI double mutant, neither of which forms wild-type biofilms. Significant, differentially regulated genes are shown in Tables S1 and S2 in the supplemental material. Table S1, comparing gene expression in worms with biofilm and worms without biofilm by virtue of incubation with Y. pseudotuberculosis 3384, a strain that does not form a biofilm, shows 198 upregulated genes and 77 downregulated genes. Many of these genes are uncharacterized. There are a number of differentially expressed receptor (C-lectin and nuclear hormone receptor), fatty acid metabolism, rRNA, and cytochrome P450 genes. However, we found no evidence for differential regulation of the C. elegans bah-1, bah-2, and bah-3 genes previously associated with loss of biofilm formation (15) in our transcriptomic analyses.
Comparison of gene expression in worms with biofilm and worms without biofilm by virtue of incubation with a Y. pseudotuberculosis ΔypsI ytbI mutant that does not form a biofilm identified 159 downregulated genes and only 1 upregulated gene. Again, many differentially expressed genes are not characterized. Notably, the list contains five highly downregulated (2- to 23-fold) grl genes, which form cysteine-cross-linked proteins involved in intercellular signaling, and five downregulated flp genes which code for small neuromodularity peptides which regulate several behaviors, including well-coordinated sinusoidal movement (26).

Does biofilm formation on C. elegans influence the expression of genes associated with reduced food intake?

Since Y. pseudotuberculosis forms biofilm around the head of C. elegans blocking the mouth, it is possible that the worm may respond by inducing genes associated with reduced food uptake. Previously, Laing et al. (27) showed that exposure of C. elegans to the nematocidal drug ivermectin results in the induction of many genes associated with starvation. A comparison of the numbers of genes associated with biofilm formation and ivermectin treatment is summarized in Fig. 2. This shows the degree of overlap with 74 out of 276 genes upregulated (or differentially regulated?) in worms with biofilm (compared to worms without biofilm by virtue of growth on Y. pseudotuberculosis 3384) are also upregulated in C. elegans exposed to ivermectin. Furthermore, 8 out of 276 genes downregulated in response to biofilm are also downregulated in C. elegans exposed to ivermectin. In contrast, only 7 genes downregulated in worms with biofilm (compared to worms without biofilm by virtue of growth on the Y. pseudotuberculosis ypsI ytbI double mutant) were also downregulated in C. elegans exposed to ivermectin. Thus, our study, with worms having a mouth blocked with bacteria and biofilm for up to 1 h, may have also identified genes differentially regulated in response to reduced food intake as well as genes responding to the development of biofilm, although the majority of differentially regulated genes (194 and 154, respectively) appear to be associated with the development of a biofilm, rather than reduced food intake. Differentially regulated genes common to worms with biofilm and worms exposed to ivermectin are summarized in Table S3 in the supplemental material. They include genes associated with fatty acid metabolism (fat-3, fat-7, far-7, and acs-17) and cyp-37B1 and scl-2 (thought to be associated with fat storage).
FIG 2 Venn diagram showing overlap of differentially regulated genes in C. elegans with Y. pseudotuberculosis biofilm and C. elegans exposed to ivermectin. (A) Genes upregulated in response to ivermectin; (B) genes downregulated in response to ivermectin; (C) genes differentially regulated in C. elegans with 1-h Y. pseudotuberculosis YPIII biofilm.

Do differentially regulated genes identified by transcriptome analysis influence biofilm development on C. elegans?

To investigate whether C. elegans genes identified as being differentially regulated following exposure to Y. pseudotuberculosis YPIII influenced biofilm development, deletion mutants, where available, for genes identified in Table S1 in the supplemental material were obtained from the Caenorhabditis Genetics Center (CGC) (see Table S4 in the supplemental material). No obvious differences in Yersinia biofilm formation were observed on nematodes with deletions in the following genes: clec-4 [strain RB1660; genotype clec-4(ok2050) chromosome II (II)], flp-9 [RB2067; genotype flp-9(ok2730) IV], flp-13 [NY2037; genotype flp-13(ynIs37) II], snf-11 [RM2710; genotype snf-11(ok156) V], grd-6 [RB2072; grd-6(ok2735) V], grd-10 [RB2192; genotype grd-10(ok2972) IV], acs-17 [RB1377; genotype acs-17(ok1562) X], anc-1 [CB3335; genotype anc-1(e1802) I], and gmn-1 [VC1248; genotype gmn-1 and Y7B82A.18(ok1708) III]. These mutants showed the aberrant motion (data not shown) and normal pattern of biofilm development observed after exposure to Y. pseudotuberculosis YPIII (illustrated in Fig. 3A to C). However, on the Δclec-49 mutant [RB1870; genotype clec-49(ok2416) V], biofilm formation was delayed by 3 h (data not shown). In contrast, deletion of fat-3 (parental strain N2; Jennifer Watts, personal communication) completely abrogated biofilm formation. Y. pseudotuberculosis YPIII is unable to form a biofilm on a C. elegans Δfat-3 mutant [BX30; genotype fat-3(wa22) IV] after 30-min exposure (Fig. 1E) and shows only minimal bacterial attachment with no biofilm development after 24 h (Fig. 1F and Fig. 3D to F).
FIG 3 C. elegans fat-3 is essential for biofilm formation. (A to F) C. elegans showing phase-contrast (A and D), fluorescence (B and E), and overlay (C and F) micrographs of worms exposed to Y. pseudotuberculosis YPIII for 24 h. (A to C) C. elegans N2, showing extensive biofilm over mouth and anterior end; (D to F) C. elegans Δfat-3 mutant, showing a lack of biofilm development after 24-h exposure. Fluorescence in the guts of worms shows the ingested bacteria.

Identification of genes required for biofilm formation that are differentially regulated in Yersinia flhDC and QS mutants.

Since we were unable to recover sufficient Yersinia biofilm from C. elegans for determination of the bacterial transcriptome and as we have previously demonstrated the contribution of QS and FlhDC to biofilm development on C. elegans (17), we compared the transcriptomes of the wild-type Y. pseudotuberculosis YPIII with flhDC, ypsI ytbI, ypsR ytbR, and ytbI mutants, respectively, in an effort to identify additional Yersinia genes required for biofilm formation. The results obtained are shown in Tables S6 to S9 in the supplemental material. (The complete lists are available through BugsBase accession number E-BUGS-147 []) and also ArrayExpress (accession number E-BUGS-147). Table 1 lists a selection of the differentially regulated genes present in more than one mutant. To validate the trends revealed in the microarrays, qPCR was performed comparing gene expression of hutH, mgtB, and adhE in the ΔflhDC mutant and hutH in the ΔypsR ytbR mutant compared with Y. pseudotuberculosis YPIII. In the ΔflhDC mutant, hutH was upregulated 3-fold as shown by microarray and 22-fold as shown by qPCR (P = 0.001), mgtB was upregulated 8-fold by microarray and 70-fold by qPCR (P = 0.02), whereas adhE was downregulated 4-fold by microarray and 5-fold by qPCR (P = 3.2 × 10−6). Similarly, in the ΔypsR ytbR mutant, hutH was upregulated 3-fold as shown by microarray and 14-fold as shown by qPCR (P = 0.002). The microarray results also confirm our previous observations with respect to the interdependence of the ypsRI and ytbRI QS systems (19), given that ypsI expression is reduced in both the Y. pseudotuberculosis ypsI ytbI double mutant and ytbI single mutant (Table 1; see Tables S7 and S9 in the supplemental material). Although few differentially regulated flagellar genes were observed in the transcriptomes of the QS mutants in log phase compared with the wild type under the same growth conditions, we have previously shown that QS regulates flagellum-mediated swimming motility and the motility master regulator FlhDC in a temperature-dependent manner (19). Since flhDC mutants also fail to form a biofilm on C. elegans (17), we determined the flhDC transcriptome (Tables 1 and S6) for comparison with those of the QS mutants to search for common differentially regulated genes potentially involved in biofilm development. From the data obtained, it is clear that FlhDC regulates not only flagellum-mediated motility but also a wide variety of metabolic, regulatory, transport, and virulence genes. This contrasts with the relatively few genes under QS control (Table 1 and compare Tables S6 to S9). Of these, the histidine utilization pathway (hut) genes (hutG, hutH, hutI, and hutU) were upregulated in both flhDC and QS (ypsI ytbI and ypsR ytbR) mutants. Other genes upregulated include the lipoprotein B precursor osmB and a conserved hypothetical protein (YPO2897 in Y. pestis). Downregulated genes include l-ribulose-5PO4 4-epimerase (araD), an ABC transporter periplasmic sugar-binding protein Yptb0802 (YPO3328 in Y. pestis), a putative sugar transporter permease subunit Yptb0800 (YPO3330 in Y. pestis), and a putative exported protein (YPO3050 in Y. pestis).
TABLE 1 Differentially regulated genes in Y. pseudotuberculosis YPIII flhDC and QS mutantsa
GeneProductbGene expression in the following Y. pseudotuberculosis mutantc:
flhDCypsI ytbIypsR ytbRytbI
hutHHistidine ammonia lyase3↑6↑3↑ 
hutUUrocanate hydratase 9↑5↑ 
hutGN-Formylglutamate amidohydrolase 7↑  
araDl-Ribulose-5PO4 4-epimerase5↓  5↓
osmBLipoprotein B precursor5↑4↑  
YPO3328Sugar ABC transporter8↓  6↓
YPO3330Sugar ABC transporter8↓  7↓
YPO2897Conserved hypothetical protein3↑ 4↑ 
YPO3050Putative exported protein4↓  12↓
ytbIAHL synthase YtbI 13↓ 8 ↓
Differentially regulated genes in Y. pseudotuberculosis YPIII flhDC and QS mutants (ypsI ytbI and ypsR ytbR mutants, respectively) compared to Y. pseudotuberculosis YPIII. P values are given in Tables S6 to S9 in the supplemental material. In the flhDC mutant, transcriptional regulation of hutH, mgtB, and adhE (Table S6) were all confirmed by qPCR; in the ypsR ytbR mutant, hutH (Table S8) was confirmed by qPCR.
AHL, acyl-homoserine-lactone.
The fold increase (arrow pointing up) or decrease (arrow pointing down) is shown.

Mutation of hutC impacts reciprocally on biofilm development and motility.

Given our unexpected finding that genes within both hut operons (hutCIG and hutUHT) are controlled by both QS and FlhDC, we constructed a deletion/insertion mutant of the transcriptional regulator of the hut operons, HutC, in order to investigate the relationship between the hut operons and biofilm formation on C. elegans. While we observed no differences in the growth of the parent YPIII strain and the hutC mutant in a minimal medium containing histidine as the sole nitrogen source (data not shown), Fig. 4 shows that compared with the parent Y. pseudotuberculosis YPIII strain, the hutC mutant has greatly reduced biofilm. Figure 4A shows a lack of biofilm on C. elegans at 6 h postinfection, and these data are presented quantitatively by the disease severity incidence (17), which is reduced by ∼6-fold (Fig. 4B). These data are similar to those obtained for the YPIII ypsI ytbI double mutant, which also fails to form a biofilm on C. elegans (17). Since biofilm formation on C. elegans can be restored for the ypsI ytbI double mutant by curing the strain of the pYV virulence plasmid, we also cured the hutC mutant of the plasmid. Figure 4B shows that biofilm formation is restored in the absence of pYV. Since the hut genes are upregulated in the ypsI ytbI, ypsR ytbR, and flhDC mutants compared with the parent strain (Table 1; see Tables S6 to S8 in the supplemental material) and as both QS and FlhDC regulate swimming motility, we examined the motility phenotype of the hutC mutant. Figure 5A shows that the hutC mutant displays a “hypermotile” phenotype comparable to that previously demonstrated for YPIII ypsI and ypsR mutants (20), indicating that HutC represses motility. Attempts to complement the hutC mutant using a range of vectors to express the gene from both constitutive and inducible promoters have not been successful thus far.
FIG 4 Mutation of the Y. pseudotuberculosis hutC gene abrogates biofilm formation on C. elegans. (A) C. elegans showing phase-contrast (A), fluorescence (B), and overlay (C) micrographs of worms fed on the YPIII hutC mutant with no visible biofilm. (B) Biofilm severity incidences for Y. pseudotuberculosis YPIII and the isogenic hutC mutant with and without the pYV virulence plasmid. The error bars represent the standard deviations from the means, and when necessary, independent two-sample t tests were performed with values for P and n given in the text where appropriate.
FIG 5 Mutation of hutC impacts on motility and flhDC expression. (A) Swimming motility of Y. pseudotuberculosis YPIII (wild type [WT]) and the isogenic hutC mutant on semisolid agar after overnight growth. In contrast to the parent strain, the hutC mutant is highly motile, as revealed by the large halo around the inoculation point. (B) Mutation of hutC results in upregulation of the flagellar master regulator flhDC genes rather than the flagellar sigma factor gene fliA. Bioluminescent (luxCDABE) reporter gene fusions for flhDC (top graph) and fliA (bottom graph) were introduced onto the chromosome of wild-type YPIII (open triangles) and the hutC mutant (closed circles). Bioluminescence (in relative light units per optical density [RLU/OD]) was quantified as a function of bacterial growth. The higher expression of flhDC′::luxCDABE in the hutC mutant compared with the wild type suggests that HutC represses the motility master regulator consistent with the motility plate assays. All assays were done in triplicate. The error bars are masked by the symbols.
Since QS regulates expression of both flhDC and the flagellar sigma factor gene fliA (19), we determined their expression as a function of growth in the hutC mutant using lux-based promoter fusions over an 18-h period (Fig. 5B). After 12-h growth, flhDC expression was 5-fold higher in the hutC mutant compared with the parent strain, while fliA expression levels were similar although slightly delayed (Fig. 5B). These data suggest that the Hut pathway is an integral component of a regulatory cascade linking QS and FlhDC to motility and biofilm formation on C. elegans.


The Y. pseudotuberculosis-C. elegans model offers a simple means by which to gain new insights into the development of bacterial biofilm-mediated events in relation to infectious disease. Although Tan and Darby (21) provided evidence to suggest that the formation of Yersinia biofilms on C. elegans is simply a consequence of the worms moving through the bacterial lawn and accumulating bacterial biomass through passive attachment, our recent work indicates that a more dynamic biofilm maturation process is occurring involving QS (17). Here we have shown that individual bacterial cells, some of which appear to be covered with an extracellular matrix rather than a preformed biofilm, can be observed during the early stages of C. elegans infection. We interpret this observation as indicating that bacteria attach to the worm and secrete the extracellular matrix or that biofilm material from the lawn attaches to bacteria rather than to the worm. These data also suggest that the attached bacteria may be responsible for the aberrant motion of C. elegans observed early in infection rather than biomass accumulated from the bacterial lawn hindering normal movement. It is noteworthy, therefore, that flp-1 is downregulated in worms with biofilm (see Table S2 in the supplemental material). Deletion of flp-1 causes aberrant movement, that is exaggerated body bends, identical to the aberrant movement of worms with biofilm; and overexpression of flp-1 causes flattening of the sinusoidal movement (26). flp-1 exaggerated body bends, also termed “loopy”, which is a phenotype also exhibited by goa-1 mutants (28) and when rho-1 is overexpressed (29). Both goa-1 and rho-1 code for small GTP-binding proteins with rho-1 enhancing acetylcholine release at synapses. It is of interest, therefore, to observe the downregulation by a factor of 2 to 23 of a number of C. elegans grl genes (grl-9, grl-17, grl-20, grl-23, and grl-25) when comparing the transcriptomes of worms exposed to Y. pseudotuberculosis YPIII for 1 h, which exhibit exaggerated body bends but have a biofilm detectable only by SEM, and the ΔypsI ytbI mutant with no biofilm (Table S2). Thus, worms exhibiting exaggerated body bends have a set of intercellular signaling (grl) genes highly downregulated alongside downregulated flp-1. In attempting to explain the aberrant movement in the absence of sufficient biofilm to mechanically interfere with movement, we speculate that downregulated intercellular signaling has an analogous effect to deletion of goa-1 or overexpression of rho-1, which results in the same exaggerated-bend phenotype and/or that downregulation of flp-1 imparts the exaggerated-bend phenotype. In either case, the aberrant motion is a consequence of bacterial attachment rather than a mechanical effect caused by a mass of biofilm and bacteria. It is conceivable that the altered C. elegans translocation is a consequence of the release of an as yet unidentified bacterial toxin under QS control, given that Y. pseudotuberculosis QS mutants cause aberrant movement only 3 to 4 h postinfection (17). However, no obvious differentially expressed candidate toxin genes could be identified from our transcriptome experiments comparing the wild-type Y. pseudotuberculosis with the QS mutants.
There is, however, some overlap between worms with biofilm and genes identified as a response to ivermectin exposure (27) which are thought to be a response to reduced food intake (30) as summarized in Fig. 2 and Table S3 in the supplemental material. However, the majority of differentially regulated genes appear to be associated with biofilm development rather than reduced food intake. They include genes associated with fatty acid metabolism (fat-3, fat-7, far-7, and acs-17) and cyp-37B1 and scl-2 (thought to be associated with fat storage). In our study, fat-2 and fat-3 were upregulated and fat-7 downregulated as was the case after exposure to ivermectin (27). However, we have found that a fat-3 deletion mutant is refractory to biofilm formation. The Δ6-desaturase fat-3 gene produces two 18-carbon products, gamma-linolenic acid (GLA) and stearidonic acid (SDA). C. elegans fat-3 mutants exhibit defective cuticle functionality and display pleiotropic defects, including impaired mobility, weakened cuticle, decreased defecation rate, and irregular expulsion. These defects are associated with impaired neurotransmission (31, 32). The fat-3 gene product activity also affects the expression of a subset of genes that are specifically required for immune function (33), and mutation of fat-3 causes increased susceptibility to P. aeruginosa infection in C. elegans (33). However, we speculate that the loss of biofilm formation is due to alteration of surface coat lipids, rather than changes in the innate immune response or effects on neurotransmission, which results in the masking of surface antigens—as has been hypothesized for the prevention of bacterial adherence in srf-2, srf-3, and srf-5 mutants (14, 34) by masking the receptors used in bacterial attachment.
Genome-wide analysis of the transcriptional response of C. elegans to bacterial or fungal infection appears to be determined by the nature of the pathogen, the site of infection and the physiological imbalance provoked by infection (35). However, these studies have mainly focused on bacteria such as Pseudomonas (36), Salmonella (37), Burkholderia (38), and Staphylococcus (39) which cause intestinal infections of C. elegans, but do not form biofilms in or on the worm surface.
Our results show a distinct response from those observed for pathogens that primarily infect the C. elegans intestine. When genome-wide C. elegans gene expression in worms fed on Y. pseudotuberculosis YPIII is compared with that of an isogenic QS-negative mutant (the ypsI ytbI double mutant), several C-type lectin genes (clec-13, clec-17, clec-45, and clec-72) were downregulated after 1 h. This contrasted with the 1-h response to a non-biofilm-forming, nonisogenic Y. pseudotuberculosis strain, Y. pseudotuberculosis 3384 where clec-48, clec-49, and clec-7 were upregulated (and clec-136, clec-137, clec-197, clec-208, and clec-219 were downregulated; see Table S1 in the supplemental material). The only clec deletion mutant available (the upregulated clec-49) showed delayed biofilm formation. Multiple C-lectin genes are similarly differentially regulated in C. elegans infected with Y. pestis (40) and Vibrio cholerae (41), and are upregulated after infection by Serratia marcescens, Enterococcus faecalis, or Photorhabdus luminescens (35). Given that the C. elegans genome contains 278 genes with C-type lectin domains, their differential regulation is likely to reflect the ability of the nematode to mount distinct innate defense responses toward different pathogens (42). It is therefore perhaps not surprising to discover that very different clec gene subsets show altered regulation in response to different pathogens and even to different strains of the same species as noted for Y. pseudotuberculosis strains YPIII and 3384. This raises the possibility of a feedback mechanism whereby engagement of pathogens by specific lectins signals their increased production. Such a mechanism would be beneficial if lectins acted not only in pathogen recognition but also in their detoxification or elimination (43). In the case of Yersinia, their upregulation might be detrimental to C. elegans, promoting rather than reducing biofilm development. Since Yersinia (and also Xenorhabdus) produces hmsHFRS-dependent exopolysaccharides containing GlcNAc and as hmsHFRS mutants fail to form biofilms on C. elegans, biofilm formation is likely to involve a C. elegans lectin(s) anchored or secreted at the nematode surface. However, while the functional significance of these multiple clec gene subsets is not yet clear, it is noteworthy that in a C. elegans clec-49 mutant, biofilm formation was delayed by up to 3 h, suggesting that this specific C-type lectin protein may promote early stage biofilm formation either through direct attachment or as a consequence of downstream signaling events, given that C-type lectins are known to play developmental and structural roles as well as contribute to innate immunity (44).
Comparison of the transcriptomes of the wild-type Y. pseudotuberculosis with the QS and flhDC mutants (which fail to form biofilms on C. elegans) revealed that hut genes from both hutUHT and hutCIG operons are upregulated in both mutant classes. In other bacteria, the hut genes are known to facilitate the utilization of histidine as the sole source of carbon, nitrogen, or both (45). Whether this is the case for Y. pseudotuberculosis remains to be determined, although we have so far observed no difference in the growth of wild type and hutC mutant in minimal medium containing histidine as the sole nitrogen source. Interestingly, l-histidine has been reported to enhance biofilm formation by Acinetobacter baumannii on polystyrene plates, while mutation of hutU results in a statistically significant reduction in biofilm formation compared with the parent strain (46). In a number of bacterial species, the hut genes are regulated by HutC, a member of the GntR family of helix-turn-helix transcriptional regulators (47), which acts a repressor that is derepressed by urocanate, the first intermediate in the hut pathway (45, 48). While the Hut pathway clearly plays a central metabolic role, in P. aeruginosa, upregulation of the hut genes results in the loss of type III secretion-mediated cytotoxicity in the presence of histidine (49). In Brucella abortus, HutC also functions as a coactivator of the virB operon, which encodes a type IV secretion system essential for intracellular replication. This suggests that HutC has been coopted to allow Brucella to coordinate metabolic sensing with mechanisms to overcome intracellular host cell defenses (50). Consequently, we chose to mutate hutC in Y. pseudotuberculosis and discovered that the hutC mutant was unable to form biofilms on C. elegans. Whether this is due to changes in histidine catabolism or because HutC interacts directly with promoters in addition to those of the two hut operons as noted for Brucella and virB (50) remains to be established for Yersinia. Both hut operon promoters contain hut and lux box consensus sequences, and these data taken together with the microarray data would place HutC downstream of QS in the regulatory cascade but potentially upstream of flhDC, given that the latter is repressed by HutC. It is therefore conceivable that the QS-dependent regulation of flhDC occurs at least in part via HutC-mediated repression. However, the flhDC promoter does not contain any obvious hut transcriptional regulatory boxes, and so this effect may be indirect. Interestingly, in Yersinia enterocolitica, which does not form biofilms on C. elegans (12), FlhDC also controls a number of hut genes, including hutU, hutI, hutG, and hutP (51). A simplified diagrammatic representation of our current understanding of the relationships between QS, FlhDC, and FliA, the Hut pathway biofilm formation on C. elegans, and swimming motility is shown in Fig. 6.
FIG 6 Simplified diagrammatic representation of the regulatory links between the YpsRI and YtbRI QS systems, the motility master regulators FlhDC and FliA, and the histidine utilization (Hut) pathway in regulating swimming motility and biofilm formation on C. elegans. Lines ending with arrowheads or horizontal lines indicate activation or repression, respectively. YpsRI positively regulates its own expression and that of ytbRI (red), while YtbRI positively autoregulates ytbI expression (blue). QS positively regulates genes in both hut operons and flhDC while negatively regulating fliA (purple). Overall, QS positively impacts on both swimming motility and biofilm formation, as does flhDC (black). The hut regulatory gene hutC negatively controls flhDC and motility but is required for biofilm formation (green). The model is based on the results of this work but data from previous studies (17, 19, 20) are also included.
Taken together, our findings are consistent with a two-stage process for biofilm formation involving an initial C-type lectin receptor-mediated attachment/recognition of Yersinia to/by C. elegans followed by growth and bacterial aggregation with concomitant secretion of an extracellular biofilm matrix, the process being dependent on a correct cuticular architecture dictated by the fatty acid desaturase content. Our results also provide further insights into a sophisticated regulatory system in Y. pseudotuberculosis in which QS appears to be at the top of a regulatory hierarchy controlling the flagellar master regulator FlhDC that incorporates the Hut pathway and reciprocally links biofilm formation and flagellum-mediated motility.


This work was funded by BBSRC grant BB/D52336X/1.
We thank Maria McCrossan for SEM pictures and acknowledge Lisa Dawson for comments on the manuscript.
Some strains were provided by the CGC, which is funded by NIH Office of Research Infrastructure Programs (P40 OD010440).

Supplemental Material

File (zii999091013so1.pdf)
File (zii999091013so2.pdf)
ASM does not own the copyrights to Supplemental Material that may be linked to, or accessed through, an article. The authors have granted ASM a non-exclusive, world-wide license to publish the Supplemental Material files. Please contact the corresponding author directly for reuse.


Petrova OE, Sauer K. 2012. Sticky situations: key components that control bacterial surface attachment. J Bacteriol 194:2413–2425.
Hassett DJ, Korfhagen TR, Irvin RT, Schurr MJ, Sauer K, Lau GW, Sutton MD, Yu H, Hoiby N. 2010. Pseudomonas aeruginosa biofilm infections in cystic fibrosis: insights into pathogenic processes and treatment strategies. Expert Opin Ther Targets 14:117–130.
Guo Y, Rowe-Magnus DA. 2010. Identification of a c-di-GMP-regulated polysaccharide locus governing stress resistance and biofilm and rugose colony formation in Vibrio vulnificus. Infect Immun 78:1390–1402.
Haussler S, Parsek MR. 2010. Biofilms 2009: new perspectives at the heart of surface-associated microbial communities. J Bacteriol 192:2941–2949.
Lee HS, Gu F, Ching SM, Lam Y, Chua KL. 2010. CdpA is a Burkholderia pseudomallei cyclic di-GMP phosphodiesterase involved in autoaggregation, flagellum synthesis, motility, biofilm formation, cell invasion, and cytotoxicity. Infect Immun 78:1832–1840.
Lory S, Merighi M, Hyodo M. 2009. Multiple activities of c-di-GMP in Pseudomonas aeruginosa. Nucleic Acids Symp Ser (Oxford) 2009:51–52.
Ueda A, Wood TK. 2009. Connecting quorum sensing, c-di-GMP, pel polysaccharide, and biofilm formation in Pseudomonas aeruginosa through tyrosine phosphatase TpbA (PA3885). PLoS Pathog 5:e1000483.
Waters CM, Lu W, Rabinowitz JD, Bassler BL. 2008. Quorum sensing controls biofilm formation in Vibrio cholerae through modulation of cyclic di-GMP levels and repression of vpsT. J Bacteriol 190:2527–2536.
Yan W, Qu T, Zhao H, Su L, Yu Q, Gao J, Wu B. 2010. The effect of c-di-GMP (3′-5′-cyclic diguanylic acid) on the biofilm formation and adherence of Streptococcus mutans. Microbiol Res 165:87–96.
Darby C, Hsu JW, Ghori N, Falkow S. 2002. Caenorhabditis elegans: plague bacteria biofilm blocks food intake. Nature 417:243–244.
Drace K, Darby C. 2008. The hmsHFRS operon of Xenorhabdus nematophila is required for biofilm attachment to Caenorhabditis elegans. Appl Environ Microbiol 74:4509–4515.
Joshua GW, Karlyshev AV, Smith MP, Isherwood KE, Titball RW, Wren BW. 2003. A Caenorhabditis elegans model of Yersinia infection: biofilm formation on a biotic surface. Microbiology 149:3221–3229.
Mendoza De Gives PM, Davies KG, Clark SJ, Behnke JM. 1999. Predatory behaviour of trapping fungi against srf mutants of Caenorhabditis elegans and different plant and animal parasitic nematodes. Parasitology 119:95–104.
Hoflich J, Berninsone P, Gobel C, Gravato-Nobre MJ, Libby BJ, Darby C, Politz SM, Hodgkin J, Hirschberg CB, Baumeister R. 2004. Loss of srf-3-encoded nucleotide sugar transporter activity in Caenorhabditis elegans alters surface antigenicity and prevents bacterial adherence. J Biol Chem 279:30440–30448.
Darby C, Chakraborti A, Politz SM, Daniels CC, Tan L, Drace K. 2007. Caenorhabditis elegans mutants resistant to attachment of Yersinia biofilms. Genetics 176:221–230.
Drace K, McLaughlin S, Darby C. 2009. Caenorhabditis elegans fig-1 is a DUF23 protein expressed in seam cells and required for microbial biofilm binding to the cuticle. PLoS One 4:e6741.
Atkinson S, Goldstone RJ, Joshua GW, Chang CY, Patrick HL, Camara M, Wren BW, Williams P. 2011. Biofilm development on Caenorhabditis elegans by Yersinia is facilitated by quorum sensing-dependent repression of type III secretion. PLoS Pathog 7:e1001250.
Sun YC, Koumoutsi A, Darby C. 2009. The response regulator PhoP negatively regulates Yersinia pseudotuberculosis and Yersinia pestis biofilms. FEMS Microbiol Lett 290:85–90.
Atkinson S, Chang CY, Patrick HL, Buckley CM, Wang Y, Sockett RE, Camara M, Williams P. 2008. Functional interplay between the Yersinia pseudotuberculosis YpsRI and YtbRI quorum sensing systems modulates swimming motility by controlling expression of flhDC and fliA. Mol Microbiol 69:137–151.
Atkinson S, Throup JP, Stewart GS, Williams P. 1999. A hierarchical quorum-sensing system in Yersinia pseudotuberculosis is involved in the regulation of motility and clumping. Mol Microbiol 33:1267–1277.
Tan L, Darby C. 2004. A movable surface: formation of Yersinia sp. biofilms on motile Caenorhabditis elegans. J Bacteriol 186:5087–5092.
Epstein HF, Shakes DC (ed). 1995. Methods in cell biology, vol 48. Caenorhabditis elegans: modern biological analysis of an organism. Academic Press, San Diego, CA.
Tarr SAJ. 1972. The assessment of disease incidence and crop loss, p 632. Macmillan Press, London, United Kingdom.
Joshua GW. 2001. Functional analysis of leucine aminopeptidase in Caenorhabditis elegans. Mol Biochem Parasitol 113:223–232.
Datsenko KA, Wanner BL. 2000. One-step inactivation of chromosomal genes in Escherichia coli K-12 using PCR products. Proc Natl Acad Sci U S A 97:6640–6645.
Nelson LS, Rosoff ML, Li C. 1998. Disruption of a neuropeptide gene, flp-1, causes multiple behavioral defects in Caenorhabditis elegans. Science 281:1686–1690.
Laing ST, Ivens A, Butler V, Ravikumar SP, Laing R, Woods DJ, Gilleard JS. 2012. The transcriptional response of Caenorhabditis elegans to ivermectin exposure identifies novel genes involved in the response to reduced food intake. PLoS One 7:e31367.
Mendel JE, Korswagen HC, Liu KS, Hajdu-Cronin YM, Simon MI, Plasterk RH, Sternberg PW. 1995. Participation of the protein Go in multiple aspects of behavior in C. elegans. Science 267:1652–1655.
McMullan R, Hiley E, Morrison P, Nurrish SJ. 2006. Rho is a presynaptic activator of neurotransmitter release at pre-existing synapses in C. elegans. Genes Dev 20:65–76.
Van Gilst MR, Hadjivassiliou H, Yamamoto KR. 2005. A Caenorhabditis elegans nutrient response system partially dependent on nuclear receptor NHR-49. Proc Natl Acad Sci U S A 102:13496–13501.
Lesa GM, Palfreyman M, Hall DH, Clandinin MT, Rudolph C, Jorgensen EM, Schiavo G. 2003. Long chain polyunsaturated fatty acids are required for efficient neurotransmission in C. elegans. J Cell Sci 116:4965–4975.
Watts JL, Browse J. 2002. Genetic dissection of polyunsaturated fatty acid synthesis in Caenorhabditis elegans. Proc Natl Acad Sci U S A 99:5854–5859.
Nandakumar M, Tan MW. 2008. Gamma-linolenic and stearidonic acids are required for basal immunity in Caenorhabditis elegans through their effects on p38 MAP kinase activity. PLoS Genet. 4:e1000273.
Politz SM, Philipp M, Estevez M, O'Brien PJ, Chin KJ. 1990. Genes that can be mutated to unmask hidden antigenic determinants in the cuticle of the nematode Caenorhabditis elegans. Proc Natl Acad Sci U S A 87:2901–2905.
Engelmann I, Griffon A, Tichit L, Montanana-Sanchis F, Wang G, Reinke V, Waterston RH, Hillier LW, Ewbank JJ. 2011. A comprehensive analysis of gene expression changes provoked by bacterial and fungal infection in C. elegans. PLoS One 6:e19055.
Tan MW, Ausubel FM. 2000. Caenorhabditis elegans: a model genetic host to study Pseudomonas aeruginosa pathogenesis. Curr Opin Microbiol 3:29–34.
Aballay A, Yorgey P, Ausubel FM. 2000. Salmonella typhimurium proliferates and establishes a persistent infection in the intestine of Caenorhabditis elegans. Curr Biol 10:1539–1542.
Kothe M, Antl M, Huber B, Stoecker K, Ebrecht D, Steinmetz I, Eberl L. 2003. Killing of Caenorhabditis elegans by Burkholderia cepacia is controlled by the cep quorum-sensing system. Cell Microbiol 5:343–351.
Sifri CD, Begun J, Ausubel FM, Calderwood SB. 2003. Caenorhabditis elegans as a model host for Staphylococcus aureus pathogenesis. Infect Immun 71:2208–2217.
Bolz DD, Tenor JL, Aballay A. 2010. A conserved PMK-1/p38 MAPK is required in Caenorhabditis elegans tissue-specific immune response to Yersinia pestis infection. J Biol Chem 285:10832–10840.
Sahu SN, Lewis J, Patel I, Bozdag S, Lee JH, LeClerc JE, Cinar HN. 2012. Genomic analysis of immune response against Vibrio cholerae hemolysin in Caenorhabditis elegans. PLoS One 7:e38200.
Schulenburg H, Hoeppner MP, Weiner J, III, Bornberg-Bauer E. 2008. Specificity of the innate immune system and diversity of C-type lectin domain (CTLD) proteins in the nematode Caenorhabditis elegans. Immunobiology 213:237–250.
Nicholas HR, Hodgkin J. 2004. Responses to infection and possible recognition strategies in the innate immune system of Caenorhabditis elegans. Mol Immunol 41:479–493.
Zelensky AN, Gready JE. 2005. The C-type lectin-like domain superfamily. FEBS J 272:6179–6217.
Schwacha A, Bender RA. 1990. Nucleotide sequence of the gene encoding the repressor for the histidine utilization genes of Klebsiella aerogenes. J Bacteriol 172:5477–5481.
Cabral MP, Soares NC, Aranda J, Parreira JR, Rumbo C, Poza M, Valle J, Calamia V, Lasa I, Bou G. 2011. Proteomic and functional analyses reveal a unique lifestyle for Acinetobacter baumannii biofilms and a key role for histidine metabolism. J Proteome Res 10:3399–3417.
Hoskisson PA, Rigali S. 2009. Chapter 1: variation in form and function: the helix-turn-helix regulators of the GntR superfamily. Adv Appl Microbiol 69:1–22.
Zhang XX, Rainey PB. 2007. Genetic analysis of the histidine utilization (hut) genes in Pseudomonas fluorescens SBW25. Genetics 176:2165–2176.
Rietsch A, Wolfgang MC, Mekalanos JJ. 2004. Effect of metabolic imbalance on expression of type III secretion genes in Pseudomonas aeruginosa. Infect Immun 72:1383–1390.
Sieira R, Arocena GM, Bukata L, Comerci DJ, Ugalde RA. 2010. Metabolic control of virulence genes in Brucella abortus: HutC coordinates virB expression and the histidine utilization pathway by direct binding to both promoters. J Bacteriol 192:217–224.
Kapatral V, Campbell JW, Minnich SA, Thomson NR, Matsumura P, Pruss BM. 2004. Gene array analysis of Yersinia enterocolitica FlhD and FlhC: regulation of enzymes affecting synthesis and degradation of carbamoylphosphate. Microbiology 150:2289–2300.

Information & Contributors


Published In

cover image Infection and Immunity
Infection and Immunity
Volume 83Number 1January 2015
Pages: 17 - 27
Editor: B. A. McCormick
PubMed: 25312958


Received: 23 January 2014
Returned for modification: 17 February 2014
Accepted: 26 September 2014
Published online: 16 December 2014


Request permissions for this article.



George W. P. Joshua
Department of Pathogen Molecular Biology, London School of Hygiene and Tropical Medicine, London, United Kingdom
Steve Atkinson
School of Life Sciences, Centre for Biomolecular Sciences, University of Nottingham, Nottingham, United Kingdom
Robert J. Goldstone
School of Life Sciences, Centre for Biomolecular Sciences, University of Nottingham, Nottingham, United Kingdom
Hannah L. Patrick
School of Life Sciences, Centre for Biomolecular Sciences, University of Nottingham, Nottingham, United Kingdom
Richard A. Stabler
Department of Pathogen Molecular Biology, London School of Hygiene and Tropical Medicine, London, United Kingdom
Joanne Purves
School of Life Sciences, Centre for Biomolecular Sciences, University of Nottingham, Nottingham, United Kingdom
Miguel Cámara
School of Life Sciences, Centre for Biomolecular Sciences, University of Nottingham, Nottingham, United Kingdom
Paul Williams
School of Life Sciences, Centre for Biomolecular Sciences, University of Nottingham, Nottingham, United Kingdom
Brendan W. Wren
Department of Pathogen Molecular Biology, London School of Hygiene and Tropical Medicine, London, United Kingdom


B. A. McCormick


Address correspondence to Brendan W. Wren, [email protected].

Metrics & Citations



  • For recently published articles, the TOTAL download count will appear as zero until a new month starts.
  • There is a 3- to 4-day delay in article usage, so article usage will not appear immediately after publication.
  • Citation counts come from the Crossref Cited by service.


If you have the appropriate software installed, you can download article citation data to the citation manager of your choice. For an editable text file, please select Medlars format which will download as a .txt file. Simply select your manager software from the list below and click Download.

View Options

Figures and Media






Share the article link

Share with email

Email a colleague

Share on social media

American Society for Microbiology ("ASM") is committed to maintaining your confidence and trust with respect to the information we collect from you on websites owned and operated by ASM ("ASM Web Sites") and other sources. This Privacy Policy sets forth the information we collect about you, how we use this information and the choices you have about how we use such information.
FIND OUT MORE about the privacy policy