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. 2008 May 30;74(14):4509–4515. doi: 10.1128/AEM.00336-08

The hmsHFRS Operon of Xenorhabdus nematophila Is Required for Biofilm Attachment to Caenorhabditis elegans▿

Kevin Drace 1,2, Creg Darby 1,2,*
PMCID: PMC2493154  PMID: 18515487

Abstract

The bacterium Xenorhabdus nematophila is an insect pathogen and an obligate symbiont of the nematode Steinernema carpocapsae. X. nematophila makes a biofilm that adheres to the head of the model nematode Caenorhabditis elegans, a capability X. nematophila shares with the biofilms made by Yersinia pestis and Yersinia pseudotuberculosis. As in Yersinia spp., the X. nematophila biofilm requires a 4-gene operon, hmsHFRS. Also like its Yersinia counterparts, the X. nematophila biofilm is bound by the lectin wheat germ agglutinin, suggesting that β-linked N-acetyl-d-glucosamine or N-acetylneuraminic acid is a component of the extracellular matrix. C. elegans mutants with aberrant surfaces that do not permit Yersinia biofilm attachment also are resistant to X. nematophila biofilms. An X. nematophila hmsH mutant that failed to make biofilms on C. elegans had no detectable defect in symbiotic association with S. carpocapsae, nor was virulence reduced against the insect Manduca sexta.

The gram-negative enterobacterium Xenorhabdus nematophila is an obligate symbiont of the entomopathogenic nematode Steinernema carpocapsae (24, 25, 33). The only free-living stage of the nematode, the infective juvenile (IJ) stage, carries a microcolony of X. nematophila cells in an intestinal compartment known as the bacterial receptacle (19, 42, 43, 54). Once a suitable insect prey is found, S. carpocapsae invades the host and releases the bacteria. X. nematophila proliferates, suppresses insect immunity, and secretes insecticidal toxins that kill the host, while the nematode completes a few generations of reproduction utilizing the bacteria and host tissue for nutrients. When resources are depleted within the corpse, the nematodes reassociate with the bacteria, and a population of newly colonized IJs exits the host to begin the cycle again (16, 21, 38).

The dual interactions with invertebrates—an intimate symbiosis and a potent pathogenicity—make X. nematophila an excellent model for investigating the mechanisms of mutualistic versus virulent associations (25, 33, 43). Several factors, including some involved in transcriptional regulation or nutrient acquisition, have been demonstrated to play roles in both symbiotic and pathogenic behaviors of X. nematophila and other bacterial species (8, 12, 26, 31, 32).

Diverse bacteria contain homologous 4-gene operons involved in biofilm formation. The genes and their products were first characterized in Staphylococcus epidermidis, in which the icaADBC operon is required for biofilms that contain polysaccharide intercellular adhesin (PIA) (22, 29, 41). The major constituent of PIA is a β-1,6-linked homopolymer of N-acetyl-d-glucosamine (GlcNAc) synthesized by the glycosyltransferase IcaA (22). In Yersinia pestis, the hmsHFRS operon is necessary for biofilm-mediated colonization of fleas (37). Antibodies raised against S. epidermidis PIA react with Y. pestis exopolysaccharide (4), suggesting overlapping compositions. Similar operons are also found in Escherichia coli, Bordetella spp., Actinobacillus spp., Xanthomonas axonopodis, Pseudomonas fluorescens, and other bacteria (36).

Yersinia spp. and X. nematophila produce aggregates on the head of the well-characterized model nematode Caenorhabditis elegans (11, 14). In the case of Yersinia spp., the aggregate has been demonstrated to be an hmsHFRS-dependent biofilm (14). S. epidermidis does not form aggregates on the head, but it infects the C. elegans gut and kills the worm in an icaADBC-dependent manner (3).

In this study, we show that the X. nematophila aggregates on C. elegans are an hmsHFRS-dependent biofilm. We also report the results of experiments to examine the role of hmsHFRS in the life cycle of X. nematophila.

MATERIALS AND METHODS

Bacterial strains, plasmids, and growth conditions.

Strains and plasmids are listed in Table 1. Escherichia coli and X. nematophila were grown at 37°C and 25°C, respectively, in Luria-Bertani (LB) broth. For X. nematophila, the LB broth was supplemented with 0.1% sodium pyruvate (LBP) (62). To distinguish primary and secondary forms, X. nematophila was grown on LB agar containing 0.0025% bromothymol blue and 0.004% triphenyltetrazolium chloride (LBTA) (59). Antibiotics were used at the following concentrations: ampicillin, 10 μg/ml; chloramphenicol, 10 μg/ml; and kanamycin, 30 μg/ml.

TABLE 1.

Strains and plasmids

Strain or plasmid Description Source or reference
E. coli strains
    OP50 C. elegans food source 6
    S17-1(λpir) Donor strain for conjugations 53
X. nematophila strains
    ATCC 19061 Wild type American Type Culture Collection
    AN6/2 ATCC 19061; form variant 59
    HGB340 Green fluorescent protein-expressing derivative of ATCC 19061; Cmr 43
    CDY484 19061 ΔhmsH This study
    CDY486 19061 ΔhmsH(Tn7:hmsH); Cmr This study
Plasmids
    pCVD442 Suicide vector; oriR6K mobRP4 sacB Ampr 15
    pRY109 Source of cat gene 63
    pCBD204 pCVD442 derivative plus cat gene from pRY109 This study
    pCBD206 pCBD204 derivative lacking bla gene This study
    pCBD207 pCBD206 derivative plus 5′ and 3′ regions flanking hmsH; Cmr This study
    pEVSCamKan Transposon delivery vector, Tn7 transposon; oriR6K mobRP4 Cmr Kanr E. Martens, University of Wisconsin—Madison
    pCBD218 pEVSCamKan derivative plus hmsH under native promoter; Cmr Kanr This study
    pUX-BF13 Helper plasmid for transposition, Tn7 transposase; Ampr 2
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C. elegans strains, growth, and maintenance.

C. elegans was maintained on NGM agar seeded with E. coli (OP50) at 20°C as described previously (61). The wild-type strain was N2 Bristol, and the following mutations were used: srf-2(yj26), srf-3(yj10), srf-4(ct109), srf-5(ct115), srf-6(yj5), srf-8(dv38), bus-1(e2678), bus-2(e2687), bus-3(e2696), bus-4(e2693), bus-5(e2688), bus-6(e2691), bus-8(e2698), bus-10(e2702), bus-12(e2740), bus-13(e2710), bus-14(e2779), bus-15(e2709), bus-16(e2802), bus-17(e2800), bus-18(e2795), bah-1(br1), bah-2(br7), and bah-3(br9).

Biofilm formation on C. elegans.

Overnight bacterial cultures were used to seed NGM plates that were then incubated overnight at room temperature. To assess biofilm formation on adult worms, wild-type C. elegans worms were incubated for 24 to 48 h on the bacterial lawns. To analyze larvae, gravid adult nematodes were allowed to lay eggs on bacterial lawns for several hours and then removed; the next day, larvae that had hatched were examined.

Lectin staining of biofilm.

C. elegans worms incubated on X. nematophila overnight were washed three times in M9 buffer (17) and incubated with wheat germ agglutinin (WGA) conjugated to Texas Red (EY Laboratories, San Mateo, CA). Incubation was carried out at 20 μg/ml in 10 mM phosphate, 150 mM NaCl for 30 min at room temperature. Worms were washed four times in buffer and examined by epifluorescence microscopy.

Phylogenetic analysis.

Sequence alignments and trees were constructed with MEGA4 (55). Evolutionary distances were computed by using the maximum composite likelihood method (56). The tree was searched by using the close-neighbor-interchange algorithm at a search level of 3, and the neighbor-joining algorithm was used to generate the initial tree (43, 52).

Construction of hmsH mutant.

Table 2 lists primers used in this study. Briefly, the chloramphenicol acetyltransferase gene (cat) from plasmid pRY109 was isolated with the restriction enzyme XmaI and subsequently ligated into plasmid pCVD442 to make pCBD204. Plasmid pCBD204 was then cut with PstI to remove the β-lactamase gene (bla) and allowed to self-ligate, producing the plasmid pCBD206.

TABLE 2.

Primers used in this study

Primer Sequencea Purpose
5-A CAGCGTCGCCGACAAAACCGCAACTTG ΔhmsH(pCBD484) construction
5-AN GCGCTCTAGACGCAACTTGAAAGACGAAGG ΔhmsH(pCBD484) construction
5-B CCTTTAGAACCGATACATGGGGGTAGTATCCCCA ΔhmsH(pCBD484) construction
3-B TGGGGATACTACCCCCATGTATCGGTTCTAAAGG ΔhmsH(pCBD484) construction
3-A ACGCCGTTATCCACGCCATAGGGCCAG ΔhmsH(pCBD484) construction
3-AN GGATGCATGCGGCAATGATCCGTTGAGTAATC ΔhmsH(pCBD484) construction
HF GCATGGGCCCACTCACATCTTTCCCCATTAAC ΔhmsH(pCBD486) complementation
HR GCATACTAGTATAGCCATTTTCCCGCAAC ΔhmsH(pCBD486) complementation
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a

Introduced restriction sites are underlined.

PCR constructs of approximately 800 base pairs 5′ and 3′ of hmsH were obtained by using the primers 5-A/5-B and 3-A/3-B, respectively. 5-B and 3-B are the reverse complement of each other and allowed for the amplification of a 5′/3′ fusion amplicon using the primers 5-AN/3-AN with the 5′ and 3′ fragments as the template. During this second round of PCR, the primer 5-AN introduced an XbaI site and 3-AN introduced an SphI site into the final product. This construct was subsequently digested with XbaI and SphI and ligated into plasmid pCBD206 to make plasmid pCBD207.

Plasmid pCBD207 was introduced into X. nematophila via conjugation using E. coli S17-1(λpir). The integration of pCBD207 was selected for by chloramphenicol resistance, and allelic-exchange recombinants were selected on LBTA without NaCl and supplemented with 5% sucrose. This created the kanamycin-sensitive X. nematophila strain CDY484, and loss of hmsH was verified by PCR using primers 5-AN and 3-AN. This nonpolar deletion left nine amino acids encoded by hmsH in strain CDY484.

Complementation of hmsH.

Amplification of the hmsHFRS promoter and hmsH was carried out with primers HF and HR (Table 2). HF introduced an ApaI restriction site, while HR introduced an SpeI site into the final product. This was subsequently ligated into plasmid pEVSCamKan, creating the plasmid pCBD218. Donor strains of E. coli S17-1(λpir) carrying pCBD218 or pUX-BF13 were used in a triparental-mating method to introduce hmsH into the attTn7 site of the X. nematophila chromosome and verified by PCR.

Life span analysis.

During the analysis, daily bacterial cultures were grown overnight at 37°C for E. coli (LB) and 25°C for X. nematophila strains (LBP), with appropriate antibiotics. These cultures were used to seed NGM plates that were then incubated overnight at room temperature. Wild-type C. elegans worms, beginning at the fourth larval stage, were transferred daily to freshly seeded plates, and the number of animals that were alive was recorded. Life was scored as the ability to respond to touch or the observation of pharyngeal pumping.

Nucleotide sequence accession numbers.

The following bacterial strains were used for comparison of 16S rRNA sequences (GenBank accession numbers in parentheses): Serratia marcescens subsp. marcescens (AJ550467); Yersinia pseudotuberculosis strain 2883 (AF365934); Y. pestis strain KIM (AF282306); X. nematophila strain riobravis (Z76738); Xenorhabdus bovienii strain DSM 4766 (X82252); Photorhabdus luminescens subsp. luminescens strain Hm (Z76742); Erwinia amylovora strain Ea1/79 (AJ010485); E. coli strain ATCC 11775T (X80725); and Salmonella enterica subsp. enterica serovar Typhimurium strain 65946 (AF227869). The GenBank accession number for the newly deposited X. nematophila hmsHFRS sequence is EU708329.

RESULTS

X. nematophila biofilm adheres to C. elegans worms.

When C. elegans worms are grown on E. coli, the normal laboratory food, no biofilm accumulates on the heads of the animals, but after 24 h of exposure to X. nematophila bacteria, approximately 50% of the worms have biofilms attached (Fig. 1). Previous reports characterizing phenotypic variation of X. nematophila demonstrate that form II cells are defective for multiple characteristics, including antibiotic and crystalline protein production, motility, and binding of the indicator dye bromothymol blue (5, 10, 23, 59). Consistent with these pleiotropies, worms exposed to the form II variant of X. nematophila did not accumulate biofilm (Fig. 1B).

FIG. 1.

FIG. 1.

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X. nematophila biofilm formation on C. elegans. (A) Adult C. elegans worms exposed to E. coli or X. nematophila for 2 days. (B) Percent of larvae with biofilm attachment after 1 day of exposure to indicated bacteria.

X. nematophila biofilms on C. elegans worms are strikingly similar to those made by Yersinia (14, 39, 57). To determine if this similarity extends to the genetic level, we tested whether C. elegans mutants that are resistant to Yersinia biofilm attachment are also resistant to X. nematophila biofilms. Mutations in bah-1, bah-2, bah-3, bus-2, bus-4, bus-12, bus-17, srf-2, srf-3, and srf-5 each confer resistance to Yersinia biofilms (13), and C. elegans worms carrying these mutations are completely resistant to X. nematophila biofilms (Table 3). These mutations are known to alter the C. elegans surface (13, 27, 28, 30, 35, 40, 48-50). Mutations in srf-4, srf-6, and srf-8 conferred partial resistance to biofilm accumulation (Table 3). Conversely, bus mutations not affecting the ability to bind X. nematophila biofilms also do not affect the ability to bind Yersinia biofilms (K. Drace, unpublished results). This concordance suggests strong similarity between the biofilms of the two genera.

TABLE 3.

C. elegans susceptibility to biofilm attachment

Genotype Mean % ± SD of C. elegans worms scored No. of C. elegans worms scored
Wild type 46.4 ± 22.5 469
srf-2(yj262) 0 ± 0 168
srf-3(yj10) 0 ± 0 137
srf-4(ct109) 14.0 ± 24.2 70
srf-5(ct115) 0 ± 0 134
srf-6(yj5) 7.7 ± 7.7 73
srf-8(dv38) 18.9 ± 24.6 40
bus-1(e2678) 47.8 ± 13.7 97
bus-2(e2687) 0 ± 0 115
bus-3(e2696) 33.6 ± 45.8 83
bus-4(e2693) 0 ± 0 136
bus-5(e2688) 81.1 ± 11.5 82
bus-6(e2691) 68.2 ± 34.8 68
bus-8(e2698) 69.4 ± 14.4 72
bus-10(e2702) 70.9 ± 5.1 91
bus-12(e2740) 0 ± 0 70
bus-13(e2710) 50.1 ± 5.2 166
bus-14(e2779) 65.2 ± 3.3 99
bus-15(e2709) 21.3 ± 18.6 68
bus-16(e2802) 55.7 ± 12.7 66
bus-17(e2800) 0 ± 0 94
bus-18(e2795) 56.3 ± 11.1 101
bah-1(br1) 0 ± 0 91
bah-2(br7) 0 ± 0 89
bah-3(br9) 0 ± 0 34
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Lectin binding of X. nematophila biofilm.

The lectins WGA and Limax flavus agglutinin (LFA) bind biofilms on C. elegans worms produced by Y. pseudotuberculosis bacteria (57, 58). WGA and LFA also bind X. nematophila biofilms attached to C. elegans worms (Fig. 2; data not shown). To distinguish bacteria from extracellular matrix, we used X. nematophila cells expressing green fluorescent protein and WGA conjugated to Texas Red (Fig. 2). The images demonstrate that the aggregate on the head of the nematodes is a combination of bacterial cells and extracellular matrix, i.e., a biofilm. WGA has affinity for β-linked GlcNAc and, to a lesser degree, sialic acid (N-acetyl-d-neuraminic acid) (7, 45, 46). In addition, the X. nematophila biofilms on C. elegans worms were bound by LFA (data not shown), which has high affinity for sialic acid (44).

FIG. 2.

FIG. 2.

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Lectin staining of X. nematophila biofilm. (A) Differential interference contrast image of biofilm attached to C. elegans head. (B to D) X. nematophila bacteria expressing green fluorescent protein (B); staining with WGA-Texas Red (C); and merged image of panels B and C (D).

The hmsHFRS operon is required for biofilm attachment to C. elegans.

A comparison of rRNA sequences from several Enterobacteriaceae species (9) indicates the evolutionary relationships between X. nematophila and other species (Fig. 3A). Interestingly, the closely related species X. bovienii and P. luminescens do not possess homologues of the hmsHFRS operon, nor do they produce biofilms that attach to the head of C. elegans worms (data not shown). Because Yersinia biofilms on C. elegans require the hmsHFRS operon (14), we used sequence data from the Xenorhabdus Genome Sequencing Project to locate the homologous operon in X. nematophila. We have named this operon hmsHFRS for consistency with Yersinia spp. (Fig. 3B) (Xenorhabdus Genome Sequencing Project; http://www.xenorhabdus.org). Blast analysis revealed that Yersinia spp. contain the closest homologues of the X. nematophila operon overall, with HmsH, HmsF, HmsR, and HmsS sharing 47%, 65%, 69%, and 48% sequence identity, respectively, with both Yersinia species (1). The next-closest homologues are in the plant pathogen Erwinia carotovora and E. coli. The organization of the operon remains largely unchanged in these species.

FIG. 3.

FIG. 3.

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Relationship of X. nematophila to other Enterobacteriaceae. (A) Unrooted phylogenetic tree of 16S sequences from several related species inferred by the minimum evolution method (51). The bootstrap consensus tree shown is inferred from 10,000 replicates. Evolutionary distances were computed by using the maximum composite likelihood method and are in units of the number of base substitutions per site (18, 56). Scale bar equals 0.5% sequence divergence. (B) The percent amino acid identity of X. nematophila HmsHFRS to Y. pestis and Y. pseudotuberculosis HmsHFRS, to E. coli PgaABCD, and to Bordetella pertussis BpsABCD. NH, no significant homology.

In Y. pestis, nonpolar deletions in any of the four genes within this operon result in a biofilm-defective strain (20). Therefore, to determine whether X. nematophila hmsHFRS is required for biofilms on C. elegans, we constructed a nonpolar deletion mutant of hmsH. In Y. pestis, HmsH is a predicted β-barrel protein with a large periplasmic domain and has been shown to localize to the outer membrane (4), although its function is unknown. HmsH is essential for X. nematophila biofilms on C. elegans (Fig. 4B). This phenotype was complemented by integrating hmsH, under the control of the hmsHFRS promoter, into a heterologous site on the chromosome (Fig. 4C).

FIG. 4.

FIG. 4.

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hmsHFRS is required for biofilm formation but not virulence against C. elegans. (A to C) C. elegans exposed to wild-type X. nematophila (A), the biofilm-defective ΔhmsH mutant (B), and the ΔhmsH mutant complemented with hmsH (C). (D) Life span assay of C. elegans exposed to wild-type X. nematophila, ΔhmsH X. nematophila, ΔhmsH X. nematophila complemented with hmsH (ΔhmsH + hmsH), and E. coli.

Colonization and pathogenesis are not hmsH dependent.

We compared the abilities of wild-type and biofilm-defective X. nematophila strains to colonize the IJ stage of S. carpocapsae. We saw no difference in the initial colonization of nematodes (approximately 200 CFU per IJ at week 1) nor in colonization through week 12 (data not shown). hmsHFRS-dependent biofilms therefore appear unnecessary for IJ colonization under laboratory conditions.

X. nematophila is pathogenic to C. elegans (11), and we compared C. elegans killing by wild-type and that by ΔhmsH X. nematophila in this model. The time to kill 50% of worms was approximately 4 days, which is more rapid than previously reported (11). The greater virulence in our experiments may be due to the use of a different X. nematophila strain or to the fact that we transferred C. elegans worms daily to fresh bacteria. No difference in the rate of killing was detected between wild-type and ΔhmsH strains (Fig. 4D).

Lastly, we analyzed insect pathogenesis by injecting X. nematophila cells into Manduca sexta larvae. No difference in virulence between wild-type and ΔhmsH strains was observed with either exponential- or stationary-phase bacteria (data not shown).

DISCUSSION

We have shown that X. nematophila form I cells make a biofilm that adheres to the head of C. elegans worms (Fig. 1) and that biofilm production is hmsHFRS dependent.

X. nematophila and Yersinia biofilms on C. elegans are similar in several respects. The biofilms are bound by the same lectins, WGA and LFA, suggesting that the attached extracellular matrix contains β-linked GlcNAc and/or sialic acid (57) (Fig. 2). In addition, C. elegans mutants that are resistant to Yersinia biofilms are also resistant to X. nematophila biofilms (13, 35, 39) (Table 3), suggesting that the two exopolysaccharides have the same receptor on the nematode surface.

Biofilm production in Yersinia spp. is dependent on the hmsHFRS operon (14). We show here that X. nematophila hmsH is necessary for biofilm formation, demonstrating functional conservation of hmsHFRS between the two genera (Fig. 4A to C). hmsHFRS-dependent biofilms are made by several species of bacteria (36, 41, 47, 60) and have been demonstrated to be involved in diverse functions, including insect colonization by Y. pestis and the C. elegans pathogenesis of S. epidermidis (3, 14, 37, 39).

The role of biofilms in the life cycle of X. nematophila is unclear. Because the biofilms bind to the exterior of C. elegans, we asked whether the bacteria use a biofilm to colonize the interior of their natural nematode host, S. carpocapsae. We grew IJs on monocultures of bacteria in vitro and quantified colonization through week 12 but found no defect in the hmsH mutant (data not shown). Y. pestis makes an hmsHFRS-dependent biofilm in fleas, its insect vector (34, 37). We therefore asked whether hmsHFRS biofilms are a virulence factor for X. nematophila insect pathogenesis. When X. nematophila cells were injected into M. sexta larvae, there was no difference between killing by the wild type and that by the hmsH mutant. However, in these experiments we tested virulence by injecting bacteria directly into the insect, and it is possible that the biofilm function requires infection by the natural route, i.e., invasion by S. carpocapsae IJs followed by release of X. nematophila cells within the insect. Furthermore, a role for biofilms in insect pathogenesis cannot be ruled out by results with a single experimental prey, M. sexta. It is conceivable that virulence against other insect species requires biofilms.

X. nematophila is a potential food source for bacterivorous nematodes, such as C. elegans, and therefore, the bacteria could have defense mechanisms to protect them from free-living worms that invade the parasitized insect cadaver. Consistent with this notion, X. nematophila produces an unidentified heat-stable product that is toxic to C. elegans (11). We asked whether X. nematophila biofilms contribute to C. elegans killing and found that they do not (Fig. 4D). However, biofilm formation on nematode heads could nevertheless be a defense mechanism that does not require killing, because covering a nematode's head and blocking its feeding protects bacteria from predation. Our results are in contrast to those with S. epidermidis, which does not make biofilms on the head but infects and kills the worms in a biofilm-dependent manner (3).

Although our experiments did not reveal the place of biofilms in the X. nematophila life cycle, this may be because laboratory conditions do not fully model the natural environment. In the laboratory, X. nematophila and its symbiont S. carpocapsae are grown without other organisms present. In the natural environment, many species of bacteria and invertebrates have access to the insect host as S. carpocapsae progresses through its life cycle in the cadaver, and biofilms could protect the nematode and its symbionts from these invaders. It is possible that when other microbes are present, a biofilm contributes to the monospecific colonization of S. carpocapsae by X. nematophila.

Acknowledgments

We thank Heidi Goodrich-Blair and Erin Herbert for the M. sexta experiments, as well as plasmids, strains, and comments on the manuscript. We also thank Barry Goldman and the Xenorhabdus Genome Sequencing Project for unpublished sequence data. We thank Steven Forst for X. nematophila strains and Jonathan Hodgkin and Samuel Politz for C. elegans strains. Additional strains were provided by the Caenorhabditis Genetics Center, which is supported by the NIH National Center for Research Resources.

This work was supported by National Institutes of Health (NIH) research grant AI057512 (C.D.), NIH training grant HL007553 (K.D.), and the Sandler Family Supporting Foundation (C.D.).

Footnotes

▿

Published ahead of print on 30 May 2008.

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