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. 2011 Oct;77(20):7247–7254. doi: 10.1128/AEM.05328-11

Role of Mrx Fimbriae of Xenorhabdus nematophila in Competitive Colonization of the Nematode Host ▿

Holly Snyder 1,2, Hongjun He 2, Heather Owen 1, Chris Hanna 1, Steven Forst 1,*
PMCID: PMC3194880  PMID: 21856828

Abstract

Xenorhabdus nematophila engages in mutualistic associations with the infective juvenile (IJ) stage of specific entomopathogenic nematodes. Mannose-resistant (Mrx) chaperone-usher-type fimbriae are produced when the bacteria are grown on nutrient broth agar (NB agar). The role of Mrx fimbriae in the colonization of the nematode host has remained unresolved. We show that X. nematophila grown on LB agar produced flagella rather than fimbriae. IJs propagated on X. nematophila grown on LB agar were colonized to the same extent as those propagated on NB agar. Further, progeny IJs were normally colonized by mrx mutant strains that lacked fimbriae both when bacteria were grown on NB agar and when coinjected into the insect host with aposymbiotic nematodes. The mrx strains were not competitively defective for colonization when grown in the presence of wild-type cells on NB agar. In addition, a phenotypic variant strain that lacked fimbriae colonized as well as the wild-type strain. In contrast, the mrx strains displayed a competitive colonization defect in vivo. IJ progeny obtained from insects injected with comixtures of nematodes carrying either the wild-type or the mrx strain were colonized almost exclusively with the wild-type strain. Likewise, when insects were coinjected with aposymbiotic IJs together with a comixture of the wild-type and mrx strains, the resulting IJ progeny were predominantly colonized with the wild-type strain. These results revealed that Mrx fimbriae confer a competitive advantage during colonization in vivo and provide new insights into the role of chaperone-usher fimbriae in the life cycle of X. nematophila.

INTRODUCTION

Xenorhabdus nematophila is a host-adapted enteric bacterium that engages in mutualistic associations with specific entomopathogenic nematodes and is also pathogenic toward diverse insects (1, 8, 18, 46). X. nematophila colonizes a specialized region of the anterior intestine (receptacle) of the infective juvenile (IJ) stage of the nematode (8, 41). The IJ enters the intestine of diverse insect hosts and subsequently invades the insect body cavity and releases X. nematophila into the insect blood (hemolymph). During early stages of infection, the bacteria adhere to connective tissue and musculature surrounding the insect midgut (36). X. nematophila proliferates in the hemolymph, where it inactivates the host immune response and produces an array of toxins and exoenzymes that are involved in killing the host. The bacteria also secrete antimicrobial compounds that suppress microbial competitors (16, 22, 33, 35, 48). Proliferation of Xenorhabdus in the insect establishes a nutrient base for nematode growth and reproduction. After several cycles of reproduction the nematodes develop into IJ progeny that are colonized by one or a few X. nematophila cells that multiply in the nematode receptacle, which is formed by 2 specialized intestinal cells (29, 30, 41). Almost all IJ progeny that emerge from the insect cadaver contain X. nematophila, though the level of colonization can vary considerably between individual IJs (41). The number of CFU per IJ was also shown to decline with time (15, 20).

The molecular details of nematode colonization are beginning to be understood. Inactivation of genes encoding cell surface proteins, transcription factors, and metabolic enzymes render X. nematophila defective for colonization (12, 21, 23, 30, 49). The cellular structures and receptors involved in colonization of the nematode by X. nematophila have not yet been elucidated. The role of fimbriae in specific binding to glycoproteins on the cell surface of host tissues by bacterial pathogens has been extensively studied; however, little is known concerning their function in the colonization process of mutualistic bacteria. X. nematophila produces mannose-resistant (Mrx) chaperone-usher-type fimbriae (20). These fimbriae, widely expressed on the surface of many members of the Enterobacteriaceae, are encoded by operons consisting of 7 to 10 structural genes that can be regulated by various mechanisms, including promoter inversion (4). Most enteric bacteria studied contain numerous fimbrial operons. Photorhabdus luminescens, the sister taxon of X. nematophila, possesses 11 different fimbrial operons (13, 42), two of which have been characterized. The maternal-adhesion-defective (mad) operon contains 8 structural genes for fimbriae that are required for binding to rectal gland cells in the maternal nematode (42). The bacteria are subsequently transmitted into the body cavity of the maternal nematode where colonization of the IJ progeny takes place. The mrf fimbrial operon of P. luminescens consists of 9 structural genes (31); however, its role in nematode colonization remains unknown. The expression of both fimbrial operons is controlled by promoter inversion.

The mrx operon of X. nematophila is one of the smallest chaperone-usher-type fimbrial operons so far studied and contains a major subunit (mrxA), an usher protein (mrxC), a chaperone (mrxD), a minor subunit (mrxG), and adhesin (mrxH) (20). The mrx operon is positively controlled by the global regulator Lrp and negatively regulated by the response regulator CpxR and is not controlled by promoter inversion (20, 46). The mrxA operon is the only complete fimbrial operon in the genome of X. nematophila (unpublished data). Mrx fimbriae are produced by cells grown on NB agar but unlike other enteric bacteria are not produced during static growth in broth medium (9, 20, 32). The role of MrxA fimbriae in nematode colonization remains unclear. In one study a ΔmrxA strain was shown to be defective for colonization (11), while in another the ΔmrxA strain colonized as well as wild-type cells (21). In addition, a phenotypic variant (secondary) form of X. nematophila that lacks numerous traits present in wild-type cells has been characterized (2, 39). Secondary cells do not produce fimbriae; however, they colonize nematodes as well as wild-type cells (9, 32, 37, 50). Besides serving as the major subunit for fimbrial assembly, MrxA has been proposed to play a role in insect virulence. MrxA was shown to be associated with outer membrane vesicles (OMVs) that exhibited oral toxicity toward the lepidopteran Helicoverpa armigera (25). Purified MrxA exhibited cytotoxic properties and recombinant MrxA displayed oral toxicity and functioned as a pore-forming toxin on targeted insect hemocytes (5, 26).

In the present study we show that colonization of the nematode host by X. nematophila strains that lacked fimbriae was comparable to colonization by wild-type cells. Competitive colonization studies showed that Mrx fimbriae did not confer a competitive advantage for colonization when IJs were grown on mixed bacterial lawns. However, the mrx strains displayed a competitive colonization defect in vivo, indicating that fimbriae provide a competitive advantage during colonization of the nematode in the insect host. The novel aspects of Mrx fimbrial function are discussed in the context of the life cycle of X. nematophila.

MATERIALS AND METHODS

Bacterial strains, media, and growth conditions.

Table 1 lists strains and plasmids used in this study. X. nematophila cultures were grown in Luria-Bertani (LB) broth, on nutrient broth (NB) agar, and on LB agar at 30°C, and Escherichia coli was grown at 37°C. Final bacterial cultures were normalized based on the optical density at 600 nm (OD600) and used for further analysis. Unless otherwise stated Grace's insect cell culture medium (Gibco) was used for bacterial dilutions. When required, ampicillin and chloramphenicol were added to final concentrations of 50 and 25 μg ml−1, respectively. E. coli S17-1 (λpir) was used for conjugation, and the pKNOCK suicide vector (3) was used for gene disruption as described previously (6, 38).

Table 1.

Bacterial strains and plasmids used in this study

Strain or plasmid Description Source or reference
X. nematophila strains
    AN6 Wild type, phase I variant, Ampr Laboratory stock
    AN6/II (secondary) Variant form of AN6, Ampr 2
    AHA1 AN6 mrxA::Cmr 20
    ASH1 AN6 mrxH::Cmr This study
    HGB151 19061 rpoS::Kanr 48
    ΔmrxA mutant ATCC 19061 mrxA::ΩKan N. Banerjee
E. coli strain
     S17-1 (λpir) Donor strain for conjugation Laboratory stock
Plasmids
    pSTBlue-1 Cloning vector, Ampr Kanr Novagene
    pKNOCK-Cm Broad-host-range suicide vector, Cmr D. Saffarini
    pBH1 pSTBlue-1 with a 354-bp mrxH insert This study
    pKH1 pKNOCK with 354-bp insert from pBHI This study
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Examination of fimbrial production by SDS-PAGE and electron microscopy.

One hundred microliters of an overnight culture of the wild-type strain (AN6) was spread on either NB agar or LB agar and incubated at 30°C for 48 h. Cells were suspended in 1.0 ml of phosphate-buffered saline (PBS) and normalized by OD600. The cell suspension was vortexed vigorously for 2 min and cells were pelleted. Supernatants containing released fimbriae were ultracentrifuged in a TL100 Beckman centrifuge at 90,000 × g for 14 min. The resulting pellet was resuspended in 20 μl of SDS loading buffer and proteins were resolved on a 15% SDS-PAGE gel. For electron microscopic analysis, cells incubated for 48 h were resuspended in 1.0 ml of phosphate-buffered saline and samples were diluted 1:20. Resuspended cells (12 μl), water (12 μl), and 0.8% phosphotungstic acid (PTA) (12 μl) were mixed on Parafilm. Grids were floated on top of the sample for 60 s and transferred to water for 30 s and then to 0.8% PTA for 30 s.

Preparation of polyclonal antiserum and Western blot analysis.

MrxA was obtained from the wild-type strain grown on NB agar for 48 h. Cells washed from the agar surface with phosphate-buffered saline were vortexed for 2 min and pelleted, and the resulting supernatant containing released fimbriae was ultracentrifuged and resolved by SDS-PAGE as described previously. The MrxA protein band was excised and rabbit antisera were produced by the biotechnology company Bio-Synthesis (Lewisville, TX). To prevent nonspecific binding, MrxA antiserum was preabsorbed with the mrxA strain before Western analysis. For Western blot analysis proteins were transferred to a nitrocellulose membrane (Millipore) in 25 mM Tris-192 mM glycine buffer and subsequently immersed in 5% nonfat dry milk for 30 min. The blocked filter was incubated in MrxA antiserum diluted 1:200 for 1 h. After being washed, blots were incubated with a 1:300,000 dilution of the secondary antibody conjugated with horseradish peroxidase and developed with SuperSignal West Dura extended-duration substrate (Thermo Scientific). The specificity of the anti-MrxA antiserum was confirmed by Western blot analysis using cell pellets from wild-type and mrxA strains.

Detection of MrxA in the IJ receptacle.

To first assess the sensitivity of the MrxA antiserum wild-type cells grown on NB agar for 48 h were washed from the agar surface with 2 ml of 1× PBS and serially diluted. One milliliter of the diluted cells was vortexed for 2 min and centrifuged to remove cells, and supernatants were ultracentrifuged as described above. The resulting fimbrial pellets were resuspended in SDS loading buffer and processed for Western blot analysis. The MrxA antiserum was able to detect MrxA derived from 103 cells. To detect MrxA production in the IJ receptacle, fourth-instar Manduca sexta larvae were coinjected with either wild-type cells and aposymbiotic IJs or the rpoS strain and aposymbiotic IJs. Resulting progeny were collected in water traps (20). To confirm that the MrxA antiserum did not cross-react with nematode proteins, 50,000 aposymbiotic IJs were surface sterilized and homogenized. After removing cell debris by centrifugation the resulting supernatants were ultracentrifuged and fimbrial aggregates were resuspended in SDS loading buffer and processed for Western blot analysis. To examine the presence of MrxA in the receptacle, 50,000 IJs colonized with wild-type cells were prepared as described above.

Purification of RNA.

RNA was purified from cells grown for 48 h on either NB agar or LB agar. Cells were harvested by adding 2 ml of 1× PBS to the agar surface and scraping cells into a sterile test tube. RNA was extracted from a pellet derived from 200 μl of cell suspension by using TRIzol reagent (Sigma). RNA was resuspended in 100 μl RNase-free water and concentrations were determined by absorbance at 260 nm. Twenty microliters of 100 μg/μl of total RNA was digested with the RNase-free DNase (Qiagen). This RNA (50 μg/μl) was used for reverse transcription-PCR (RT-PCR) analysis.

RT-PCR.

Reaction mixtures (50 μl) for the AccessQuick RT-PCR system (Promega) contained 300 μg of RNA, 10 pmol each of forward and reverse primers, and 1 unit of reverse transcriptase. cDNA synthesis was conducted at 52°C for 45 min and PCR conditions were 30 s at 94°C, 30 s at 55°C, and 60 s at 72°C. 16S rRNA was used as an internal control.

Construction of the mrxH strain.

A mrxH internal fragment of 354 bp was amplified with primers 5′-CCTGGACAGAGGAAGAAGGG-3′ (forward) and 5′-GACATCAATGGAGCAGGACTGC-3′ (reverse) from the AN6 strain. The fragment was ligated into the EcoRV site of pSTBlue-1 (Novagen) and the resulting plasmid was transformed into supercompetent cells (Novagen) to produce pBH1. The PstI and XhoI fragment of pBH1 was ligated into pKNOCK-Cm and transformed into S17-1 (λpir), resulting in pKH1. After conjugation of pKH1 into AN6 the mrxH strain (ASH1) was obtained by selection on ampicillin and chloramphenicol. The mrxA strain (AHA1) was constructed as previously described (19). Both mrx strains were confirmed by RT-PCR and Western analysis. The mrxA strain did not display reversion under in vitro and in vivo conditions as shown by both RT-PCR and SDS-PAGE.

Phenotypic characterization of mrx strains.

Phenotypic plate analyses were performed for lipase activity (Tween 20 and 60), protease activity, hemolytic activity (sheep red blood cells; Remel Co.), binding of bromothymol blue, and swarming and swimming activity as previously described (34). Briefly, strains were grown for 18 h and normalized, and 6-μl aliquots were spotted onto the respective plates that were incubated overnight at 30°C. Generation times were examined by normalizing overnight cultures and inoculating 5 ml of LB broth with 100 μl. Cell concentration was determined by turbidity (Klett), OD600, and cell counting using a hemacytometer (Reichert).

Injection assay for bacterial pathogenicity.

Manduca sexta (tobacco hornworm) larvae were raised from eggs (North Carolina State University, Raleigh, NC) on artificial diet (gypsy moth wheat germ diet, North Carolina State University) with a photoperiod of 16 h light and 8 h dark. Cultures of bacterial strains were grown in Grace's insect culture medium to mid-exponential stage, normalized to an OD600 of 0.8, and diluted 105-fold. Fifty microliters (∼150 cells) was injected into the last proleg of 4th-instar M. sexta. Twenty insects were injected for each strain of X. nematophila analyzed. The time of death was monitored over 48 h, and the time to 50% lethality (LT50) was determined for each strain. The assay was repeated twice.

Colonization on bacterial lawns and in vivo.

For colonization on bacterial lawns 100 μl of mid-exponential cultures of individual strains was spread on NB oil agar plates as previously described (18). For competition assays, 1:1 ratios both of the wild-type and mrxA strain and of the wild-type and mrxH strain were mixed and spread on NB oil agar plates. Plates were incubated at 30°C for 24 h. One thousand surface-sterilized aposymbiotic IJs were added to the bacterial lawns, which were incubated in the dark. On day 14 plates were placed in water traps to collect IJ progeny. Two hundred surface-sterilized IJs were resuspended in 250 μl of LB broth. Nematodes were homogenized for 2 min, diluted 20-fold in LB broth, and plated on LB agar containing either ampicillin or chloramphenicol. To determine the level of colonization in vivo, 50 μl containing 40 aposymbiotic IJs and ∼100 to 200 CFU of either the wild-type, mrxA, or mrxH strain was coinjected into 4th-instar M. sexta. After the insects died, cadavers were placed in water traps. To assess the level of colonization IJ progeny were surface sterilized, homogenized, and dilutionally plated as described above. Unpaired Student's t test was used to determine the statistical significance of differences in colonization of the IJs with the wild-type and mrx strains. Two approaches were used to analyze the level of colonization under competitive conditions. First, 50 μl of a mixture of 40 IJs colonized either with the wild-type and mrxA strains (1:1) or with the wild-type and mrxH (1:1) strains was injected into the insect. Second, 40 aposymbiotic IJs were mixed with a 1:1 ratio either of the wild-type and mrxA strains or of the wild-type and mrxH strains, and 50 μl was injected into the insect. Insect cadavers were placed in water traps and emerging IJ progeny were surface sterilized, homogenized, and dilutionally plated.

Growth and survival of the mrxA strain in the insect hemocoel.

The wild-type and mrxA strains grown overnight in LB broth under selection were normalized to the same OD600 value and 100 μl of the normalized culture was inoculated into 5 ml of Grace's medium. Cells were grown to exponential phase and normalized to the same OD600 value. The wild-type and mrxA strains were diluted 105-fold and injected individually into insects. For experiments in which the wild-type and mrxA strains were coinjected a 1:1 mixture was made and subsequently diluted 105-fold. Fifty microliters was injected per insect and three insects were used for each time point. Fifty microliters of each culture or coculture was also plated on selective media to obtain the number of CFU injected. At 48 h, 96 h, 144 h, and 192 h hemolymph was obtained from 3 insects and 10 μl of the combined hemolymph was used for serial dilutions. Dilutions of 10−5, 10−6, and 10−7 were used for the 48-h, 96-h, and 144-h time points and 10−4, 10−5, and 10−6 dilutions were used for the 192-h time point. Fifty microliters of each dilution was plated in triplicate on both ampicillin and chloramphenicol plates to obtain CFU/ml. This experiment was repeated 4 times. Unpaired Student's t test was used to determine the statistical significance of differences in survival of wild-type and mrxA strains. To confirm that the mrx strains did not display an intrinsic competitive growth and/or survival defect the wild-type and mrxA strains were coinoculated into Grace's medium. The mrxA/total cell ratio after 168 h of incubation was found to be the same as the preinjection ratio.

RESULTS

Analysis of Mrx fimbrial production on LB agar.

X. nematophila produces Mrx fimbriae when grown on NB agar but not in NB broth or LB broth (20, 32), suggesting that solid surfaces may stimulate fimbrial production. To assess this possibility cells were grown on LB agar, a condition not previously examined. Transmission electron microscopic analysis revealed that cells grown on LB agar produced flagella rather than fimbriae (Fig. 1A). Similar results were obtained with cells grown on LB agar lacking NaCl, indicating that the presence of added salt did not account for differences in fimbrial production between NB agar and LB agar (data not shown).

Fig. 1.

Fig. 1.

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Analysis of fimbria and flagellum production on NB agar and LB agar. (A) Transmission electron micrographs of negatively strained X. nematophila. Fimbriae are produced under NB agar conditions (left), whereas flagella are produced under LB agar conditions (right). Bar, 1 μm. (B) SDS-PAGE gel of fimbriae and flagella from cells grown on NB agar (lane 1) and LB agar (lane 2). MrxA (16 kDa) is produced by cells grown on NB agar, while FliC (33 kDa) is produced by cells grown on LB agar. FliC but not MrxA is produced by the mrxA strain grown on LB agar (lane 3). OpnP is a major outer membrane protein.

Fimbrial production was further analyzed using an SDS-polyacrylamide gel approach. Pellets derived from cells grown on NB agar contained the 16-kDa major fimbrial subunit, MrxA (Fig. 1B, lane 1), whereas pellets derived from cells grown on LB agar contained the 33-kDa flagellin protein, FliC (lanes 2 and 3), which was not present in pellets derived from either an flhC strain (33) or an fliA strain (unpublished data) that lacks flagella. A small amount of protein that migrated at the level of MrxA was also present in the LB agar pellets. This protein was not present in pellets obtained from an mrxA mutant strain (lane 3) that lacked fimbriae, suggesting that a low amount of MrxA was produced under LB agar conditions. OpnP, a major porin protein derived from outer membrane vesicles that copelleted with cell appendages (33), was also present in the samples analyzed.

Nematode colonization in the absence of Mrx fimbriae.

Steinernema carpocapsae IJs are routinely propagated on X. nematophila lawns grown on NB agar (11, 30, 41). Since fimbriae are produced on NB agar but not on LB agar we compared nematode colonization of aposymbiotic IJs propagated on bacterial lawns grown on both NB agar and LB agar. Nematode progeny harvested from water traps were homogenized followed by dilutional plating to determine the number of CFU per IJ (CFU/IJ) (Table 2). IJs derived from bacterial lawns grown on LB agar were colonized at higher levels (125 CFU/IJ) than were IJs grown on NB agar (73 CFU/IJ).

Table 2.

Colonization of progeny IJs on bacterial lawns and during natural infectiona

Strain NB agar CFU/IJ P value LB agar CFU/IJ Natural infection CFU/IJ P value
Wild type 73 (4) 125 (10) 116 (6)
Secondary 54 (7) 0.008 ND 124 (10) 0.792
mrxA mutant 68 (5) 0.789 ND 119 (10) 0.818
mrxH mutant 78 (7) 0.808 ND 90 (7) 0.001
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a

Data are shown as means (SEs). The number of replicates for each strain was at least n = 18. P values compare the wild-type strain to the mutant strain under each colonization condition. ND, not determined.

To further assess the role of fimbriae in nematode colonization aposymbiotic IJs were propagated on lawns of the secondary cells, which do not produce fimbriae. The secondary strain colonized IJs to a slightly lower extent than did the wild-type cells (Table 2). It was shown previously that colonizations of IJs grown in wax worm larvae inoculated with the wild-type and the secondary strains were indistinguishable (37). We also analyzed colonization of IJ progeny in vivo by injecting Manduca sexta with aposymbiotic IJs together with either wild-type or secondary cells (Table 2). The level of colonization with the secondary cells (124 CFU/IJ) was the same as that of the wild-type cells (116 CFU/IJ). Together, these findings suggested that the presence of fimbriae on the cell surface was not required for nematode colonization. Consistent with previous findings the level of colonization of IJs grown in vivo was significantly greater than that seen for IJs grown on NB agar bacterial lawns (18).

mrxA and mrxH mutant strains colonize nematodes.

To establish that fimbriae were not required for colonization we created strains of X. nematophila in which mrx genes were inactivated. We had previously shown that inactivation of the major fimbrial subunit gene, mrxA, eliminated fimbrial production (19). RT-PCR analysis of the mrxA strain showed that the downstream genes, mrxC, mrxD, mrxG, and mrxH, were expressed (40). Mutant strains in which the major fimbrial subunit was inactivated but chaperone and usher proteins were still produced can present adhesin proteins on the cell surface that may promote colonization (24). It was previously shown in Proteus mirabilis that initial binding of adhesin protein to the usher stabilized the assembly process. In mutant strains lacking the adhesin protein fimbriae were not produced (45). For this reason the adhesin gene, mrxH, was inactivated, creating a strain in which fimbrial production was eliminated (Fig. 2). In addition, MrxA was not detected in ultracentrifuged pellets of vortexed mrxH cells (40). Besides the absence of fimbriae, the phenotypic traits and virulence properties of the mrxA and mrxH strains were indistinguishable from those of the wild-type strain. The growth rates of the mrx strains in culture were also identical to that of the wild-type strain.

Fig. 2.

Fig. 2.

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RT-PCR analysis of mrx gene expression in the mrxH strain. (Left) mrx gene expression in the wild-type strain grown on NB agar. rt- represents a control reaction without reverse transcriptase. (Middle) mrx gene expression in the mrxH strain. Note the absence of the mrxH band. (Right) Transmission electron micrograph of the mrxH strain grown on NB agar.

Colonization of nematodes by the mrx mutant strains was analyzed by propagating aposymbiotic IJs on bacterial lawns of the wild-type, mrxA, and mrxH strains. Table 2 shows that the mrxA (68 CFU/IJ) and mrxH (78 CFU/IJ) strains colonized IJs as well as the wild-type strain did (73 CFU/IJ). Aposymbiotic IJs propagated on lawns of a ΔmrxA strain were shown previously to be defective for colonization (11). We used the same ΔmrxA strain (provided by N. Banerjee) and found that nematode colonization of progeny derived from lawns of the ΔmrxA strain (86 CFU/IJ) was indistinguishable from colonization with wild-type cells (91 CFU/IJ). The reason for the discrepancy between the previous study (11) and present study remains unclear (see Discussion).

To determine whether fimbriae were required for in vivo colonization IJs carrying either the wild-type, mrxA, or mrxH strain were injected into M. sexta larvae. Table 2 shows that the level of colonization by the mrxA strain (119 CFU/IJ) was comparable to that of the wild-type strain (116 CFU/IJ), while the mrxH strain colonized to a slightly lower extent (90 CFU/IJ). Taken together with similar findings from previous analysis in our laboratory (19) these data indicate that fimbriae are not required for colonization in vivo.

Analysis of competitive colonization with the mrxA and mrxH strains.

Analysis of mutant strains under competition conditions can reveal more subtle defects not apparent when the mutant strain is studied alone. For this reason we propagated aposymbiotic IJs on comixtures of bacterial lawns containing wild-type and mrx strains. Control bacterial lawns without nematodes were monitored to ensure that the original 50:50 ratio was maintained throughout the experiment. IJ progeny grown on a bacterial lawn containing the wild-type and mrxA strains were colonized at a ratio of 46:54 (wild type to mrxA strain). Similarly, when nematodes were grown on bacterial lawns containing the wild-type and mrxH strains the colonization ratio was 56:44 (wild type to mrxH strain). Thus, the mrx strains grown on agar in the presence of the wild-type strain did not display a competitive colonization defect.

We next addressed the question of whether the mrx strains exhibited a competitive colonization defect in vivo. To first confirm that the growth and survival of the mrxA strain in the insect were comparable to those of the wild-type strain, both strains were individually injected into M. sexta and hemolymph collected over a 192-h time period was plated on both ampicillin, on which both strains grow, and chloramphenicol, on which only the mrxA strain grows (Table 3). Hemolymph collected from insects injected with the wild-type cells did not produce colonies on chloramphenicol, while hemolymph from insects injected with the mrxA strain produced the same number of colonies on ampicillin and chloramphenicol. This experiment was repeated four times. While the levels of wild-type and mrxA strains fluctuated during the early time points the growth and survival levels of the strains at the 144-h and 192-h time points were not significantly different. Together these data indicated that in vivo growth and survival of the mrxA strain were comparable to those of the wild-type strain.

Table 3.

Survival of wild-type and mrxA strains in M. sexta

Time point Survival of indicated straina
 P value
Wild type mrxA mutant
48 h 63.6 (5.2) 46.2 (2.2) 0.003
96 h 11.4 (1.6) 24.3 (3.5) 0.002
144 h 8.9 (1.6) 10.4 (1.2) 0.457
192 h 8.6 (0.8) 9.1 (1.5) 0.793
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a

Data are expressed as 108 CFU/ml and shown as means (SEs). Each value is derived from at least 17 replicate data points.

In vivo competitive colonization experiments were carried out by injecting a comixture of symbiotic IJs (50% colonized with wild type: 50% colonized with mrx strain) into M. sexta. IJ progeny harvested in water traps were homogenized and homogenates were dilutionally plated on selective media to determine the colonization ratio (Table 4). When a comixture of IJs carrying the wild-type and mrxA strains was injected, 2.7% of the IJ progeny were colonized with the mrxA strain. Similarly, 2.3% of the IJ progeny were colonized when a comixture of IJs containing wild-type and mrxH strains was injected into insects. Competitive in vivo colonization experiments were also carried out by coinjecting aposymbiotic IJs together with a 1:1 mixture of the wild-type and mrx strains. When a comixture of wild-type and mrxA strains was injected, 20.7% of the IJ progeny were colonized with the mrxA strain. When a comixture of wild-type and mrxH strains was injected, 1.4% of the IJ progeny were colonized with the mrxH strain. These findings were consistent with previous unpublished studies conducted in our laboratory (19), indicating that fimbriae confer a competitive advantage for colonization in vivo.

Table 4.

In vivo competitive colonization

Coinjection mrxA strain
 mrxH strain
CFU/IJa mrxA mutant/ wtb CFU/IJa mrxH mutant/ wtb
IJs colonized with individual strains 163.8 (3.7) 2.7% 168 (4.3) 2.3%
Aposymbiotic IJs + (wt:mutant strains) 131.4 (9.3) 20.7% 129.9 (6.8) 1.4%
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a

Total CFU on LB agar supplemented with 50 μg ml−1 ampicillin. Data are shown as means (SEs). Each value is derived from at least 20 replicate data points.

b

Data shown are percentages of IJ progeny colonized with the mutant strain when a mixture of the wild-type (wt) strain and the indicated mutant strain was injected.

The in vivo competitive colonization defect of the mrx strains could be due to indirect effects that occurred in the hemocoel rather than during colonization of the nematode. To address this possibility comixtures of wild-type and mrxA cells were injected into M. sexta and the mrxA/total cell ratio was monitored over 192 h. By this time the progeny IJs are being colonized (47). This experiment was repeated four times (Table 5). The mean of the mrxA/total cell ratio at 192 h for the four experiments was 28.5%. In contrast, the proportion of IJs colonized by the mrx strains during three of the four in vivo competitive colonization experiments (Table 4) was on average 10-fold lower, supporting the idea that fimbriae confer a competitive advantage for colonization of the IJs in vivo.

Table 5.

In vivo competition of the wild-type and mrxA strains in the hemocoel of M. sextaa

Time point Exp 1
 Exp 2
 Exp 3
 Exp 4
CFU/ml mrxA/total CFU/ml mrxA/total CFU/ml mrxA/total CFU/ml mrxA/totalc
48 h 51.9 (1.7) 29.8% 101 (2.5) 74.9% 61.8 (7.2) 85.9% 20.4 (1.9) 85.3%
96 h 9.5 (2.2) 28.4% 9.4 (0.5) 32.9% 38.7 (3.5) 54.5% 28.4 (3.0) 76.4%
144 h 5.2 (0.5) 18.3% 71.0 (13.2) 43.4% 9.9 (0.4) 69.7% 7.1 (0.4) ND
192 h 3.8 (0.3) 35.5% 10.3 (0.4) 35.0% 4.4 (0.3) 20.9% 12.0 (1.3) 22.7%
% changeb 120% 47% 24% 27%
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a

Data are expressed as 108 CFU/ml and shown as means (SEs). Each value is derived from at least 6 replicate data points.

b

% change at 192 h relative to value at 48 h.

c

% of mrxA strain present at each time point when a mixture of wild-type and mrxA strains was injected.

MrxA production inside the nematode receptacle.

The question of whether X. nematophila produces fimbriae inside the IJ receptacle remained unclear. To address this question a Western blot approach was taken to determine if MrxA was produced in the intestinal receptacle (Fig. 3). We first determined that MrxA derived from a pellet of 105 wild-type cells (Fig. 3, lane 1) was readily detectable and was still detectable with a pellet derived from 103 cells (lane 2). The antiserum was specific for MrxA since a signal was not detected with a pellet derived from 107 mrxA cells (unpublished data). Since IJs are colonized with on average 100 to 200 cells it was estimated that 5 × 104 IJs would be sufficient to detect the presence of MrxA. Surface-sterilized IJs (5 × 104) colonized with wild-type cells were macerated and vigorously vortexed to release fimbrial structures from the cell surface. After removing cell debris by centrifugation the resulting supernatant was subjected to ultracentrifugation and pellets were processed for Western blot analysis. The same procedure was carried out with aposymbiotic IJs. MrxA was detected in IJs carrying the wild-type strain (lane 3), while no cross-reactive protein was detected in aposymbiotic IJs (lane 4). These findings show that MrxA was present in the nematode receptacle of colonized IJs.

Fig. 3.

Fig. 3.

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Western blot analysis of MrxA production in the nematode receptacle. The sensitivity of the MrxA antiserum was assessed by using fimbriae obtained from 105 (lane 1) and 103 (lane 2) wild-type cells. MrxA was detected in the nematode by grinding 50,000 IJs colonized with wild-type cells (lane 3). In a control experiment, the specificity of the MrxA antiserum was confirmed by grinding 50,000 aposymbiotic IJs.

DISCUSSION

In this study we show that Mrx fimbriae of X. nematophila are not required for colonization of S. carpocapsae. The mrxA, mrxH, ΔmrxA, and secondary strains colonized IJs to the same extent as wild-type cells did. In addition, in vivo colonization with the mrxA and mrxH strains was comparable to that with the wild-type strain. Furthermore, mrx strains did not display a competitive colonization defect when cocultured with the wild-type strain. Thus, during growth on bacterial lawns and under noncompetitive conditions in the insect other cell envelope molecules such as NilB and NilC (12) appear to be primary determinants for colonization. A previous study found that the ΔmrxA strain was defective for colonization (11). The reason for the different results in the respective studies remains unclear. In the previous study the ΔmrxA strain was grown under selection (kanamycin), while in the present study cells were grown without selection. It is also possible that different strains of S. carpocapsae were used in the respective studies. It has been shown previously that fimbriae may be involved in functions other than direct colonization of the symbiotic partner. In the sister taxon Photorhabdus luminescens, Mad fimbriae were required for adhesion to specific rectal gland cells in the intestine of the maternal nematode and transmission to IJs developing inside the maternal body cavity (42). Thus, Mad fimbriae are apparently necessary for bacterial transmission rather than being directly involved in IJ colonization.

Subtle defects in the colonization ability of symbiotic bacteria have been revealed by in vivo competition experiments. For example, a strain of Vibrio fischeri in which the major type IV pilus subunit gene (pilA) was inactivated colonized the squid host normally but displayed a competitive defect in the presence of the wild-type strain (43). We found that in three of four in vivo competitive colonization experiments, <3% of the IJ progeny were colonized by mrx strains. The in vivo competitive colonization defect was not due to an intrinsic growth and/or survival disadvantage since the mrx strains grew and survived in the insect as well as the wild-type strain. When the wild-type and mrxA strains were coinoculated into M. sexta the mrxA/total cell ratio varied from an increase of 20% to a 4-fold reduction, with an overall tendency to decrease over time. The variability may be due to nonuniform distribution of the respective strains, resulting in the collection of different subpopulations during hemolymph sampling. These findings suggest that a component of the in vivo competitive colonization defect may be due to events that occur in the hemolymph, resulting in a greater proportion of wild-type cells being available for colonization. For example, a portion of the nonfimbriated strains may self-aggregate (27), nonspecifically adhere to cells and tissues to a greater extent than fimbriated cells, and/or be more susceptible to clearance by the insect immune system. However, the defect in competitive colonization was significantly greater than could be accounted for by the reduction in the mrxA/total cell ratio in the hemolymph, suggesting that fimbriae confer a competitive colonization advantage in vivo. The finding that MrxA is produced in the IJ receptacle supports this idea.

While X. nematophila genes involved in colonization have been identified the molecular details of how cell surface proteins and structures, transcription regulators, and metabolic enzymes function in the colonization process in vivo remain to be elucidated. Colonization of the IJ in the insect occurs under conditions distinctly different from those on bacterial lawns. The hemocoelic environment consists of a fluid component (hemolymph), a complex mixture of macromolecules and solutes, and a variety of cells and tissues. We speculate that in vivo conditions present a more stringent environment than bacterial lawns for colonization of the nematode receptacle. During colonization, adherence of the bacteria to receptor sites in the receptacle may represent a low-probability event. Initial adherence is apparently mediated by nonfimbrial factors. Fimbriae may favor transition to a stable association such that wild-type cells colonize the receptacle more efficiently. In uropathogenic E. coli, P pili mediate binding to glycolipids on epithelial cell surfaces. When the bacterium is exposed to external forces, such as urine flow, the P pili can elongate from a helical to a linear conformation. This property is important for the ability of a bacterium to withstand shear forces caused by urine flow (14). Mrx fimbriae may also elongate helping X. nematophila to withstand shear forces that reduce adherence to cell surfaces. The adherence of strains lacking fimbriae would be less stable, resulting in a decrease in colonization efficiency.

It is not presently known whether fimbriae are produced in the hemolymph before X. nematophila colonizes the IJs or when the bacteria reside within the receptacle. During early stages of infection, flagella are produced and most cells are motile (34). X. nematophila also produces flagella when cultured in broth and grown on LB agar. In contrast, cells grown on NB agar produce fimbriae. The primary difference between NB and LB media is that the former contains beef extract and peptone, while the latter contains casein digest (tryptone) and yeast extract. We have found that X. nematophila still produces fimbriae and not flagella on NB agar containing yeast extract (unpublished data). It is conceivable that NB agar mimics in vivo conditions that control the production of fimbriae production. Interestingly, mrxA was found to be expressed in cells growing in LB broth, suggesting that posttranscriptional events may be involved in controlling fimbrial production (reference 19 and H. Snyder, unpublished data). On the transcriptional level, expression of the mrx operon is positively regulated by Lrp (20), which in E. coli controls a large number of genes involved in amino acid biosynthesis and catabolism, transport, and production of pili (10, 44), while it more specifically controls the biosynthesis of branched-chain amino acids in other bacteria (7, 17, 28). Strains of X. nematophila in which lrp was inactivated were defective for colonization and attenuated for virulence (23). Whether Lrp plays a role in controlling mrx expression by sensing environmental signals within the insect cadaver remains to be determined. Future studies on the expression of mrx genes under different laboratory conditions and in vivo will help to further our understanding of the unique role of Mrx fimbriae in the life cycle of X. nematophila.

ACKNOWLEDGMENTS

We are grateful to N. Banerjee for providing the ΔmrxA strain and J. Witten for providing the Manduca sexta larvae.

This work was supported in part by the Shaw Award from the Milwaukee Foundation.

Footnotes

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Published ahead of print on 19 August 2011.

REFERENCES

  • 1. Akhurst R. J. 1983. Neoaplectana species: specificity of association with bacteria of the genus Xenorhabdus. Exp. Parasitol. 55:258–263 [DOI] [PubMed] [Google Scholar]
  • 2. Akhurst R. J., Boemare N. E. 1988. A numerical taxonomic study of the genus Xenorhabdus (Enterobacteriaceae) and proposed elevation of the subspecies of X. nematophilus to species. J. Gen. Microbiol. 7:1835–1845 [DOI] [PubMed] [Google Scholar]
  • 3. Alexeyev M. F. 1999. The pKNOCK series of broad-host-range mobilization suicide vectors for gene knockout and targeted DNA insertion into the chromosome of Gram-negative bacteria. Biotechniques 26:824–828 [DOI] [PubMed] [Google Scholar]
  • 4. Bahrani F. K., Mobley H. L. 1994. Proteus mirabilis MR/P fimbrial operon: genetic organization, nucleotide sequence, and conditions for expression. J. Bacteriol. 11:3412–3419 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 5. Banerjee J., Singh J., Joshi M. C., Ghosh S., Banerjee N. 2006. The cytotoxic fimbrial structural subunit of Xenorhabdus nematophila is a pore-forming toxin. J. Bacteriol. 188:7957–7962 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 6. Bao Y., Lies D. P., Fu H., Roberts G. P. 1991. An improved Tn7-based system for the single-copy insertion of cloned genes into chromosomes of Gram-negative bacteria. Gene 109:167–168 [DOI] [PubMed] [Google Scholar]
  • 7. Belitsky B. R., Gustafsson M. C., Sonenshein A. L., Von Wachenfeldt C. 1997. An lrp-like gene of Bacillus subtilis involved in branched-chain amino acid transport. J. Bacteriol. 179:5448–5457 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 8. Bird A. F., Akhurst R. J. 1983. The nature of the intestinal vesicle in nematodes of the family Steinernematidae. Int. J. Parasitol. 13:599–606 [Google Scholar]
  • 9. Brehelin M., et al. 1993. Ultrastructural study of surface components of Xenorhabdus sp. in different cell phases and culture conditions. J. Invertebr. Pathol. 61:188–191 [Google Scholar]
  • 10. Calvo J. M., Mathews R. G. 1994. The leucine-responsive regulatory protein (Lrp) a global regulator of metabolism in Escherichia coli. Microbiol. Rev. 58:466–490 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 11. Chandra H., Khandelwal P., Khattri A., Banerjee N. 2008. Type 1 fimbriae of insecticidal bacterium Xenorhabdus nematophila is necessary for growth and colonization of its symbiotic host nematode Steinernema carpocapsiae. Appl. Environ. Microbiol. 10:1285–1295 [DOI] [PubMed] [Google Scholar]
  • 12. Cowles C. E., Goodrich-Blair H. 2008. The Xenorhabdus nematophila nilABC genes confer the ability of Xenorhabdus spp. to colonize Steinernema carpocapsae nematodes. J. Bacteriol. 190:4121–4128 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 13. Duchaud E., et al. 2003. The genome sequence of the entomopathogenic bacterium Photorhabdus luminescens. Nat. Biotechnol. 21:1307–1313 [DOI] [PubMed] [Google Scholar]
  • 14. Fallman E., Schedin S., Jass J., Uhlin B., Axner O. 2005. The unfolding of the P pili quaternary structure by stretching is reversible, not plastic. EMBO Rep. 6:52–56 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 15. Flores-Lara Y., Renneckar D., Forst S., Goodrich-Blair H., Stock P. 2007. Influences of nematode age and culture conditions on morphological and physiological parameters in the bacterial vesicle of Steinernema carpocapsae. J. Invertebr. Pathol. 95:110–118 [DOI] [PubMed] [Google Scholar]
  • 16. Forst S., Clarke D. 2002. Bacteria-nematode symbiosis, p. 57–77 In Gaugler R. (ed.), Entomopathogenic nematology. CABI Publishing, Wallingford, United Kingdom [Google Scholar]
  • 17. Friedberg D., Midkiff M., Calvo J. M. 2001. Global versus local regulatory roles for Lrp-related proteins: Haemophilus influenzae as a case study. J. Bacteriol. 183:4004–4011 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 18. Goetsch M., Owen H., Goldman B., Forst S. 2006. Analysis of the PixA inclusion body protein of Xenorhabdus nematophila. J. Bacteriol. 188:2706–2710 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 19. He H. 2002. Functional analysis of the mannose fimbrial operon, mrx, in Xenorhabdus nematophila Ph.D. dissertation. University of Wisconsin—Milwaukee, Milwaukee, WI.
  • 20. He H., Snyder H. A., Forst S. 2004. Unique organization and regulation of the mrx fimbrial operon in Xenorhabdus nematophila. Microbiology 150:1439–1446 [DOI] [PubMed] [Google Scholar]
  • 21. Herbert E. E., Cowles K. N., Goodrich-Blair H. 2007. CpxRA regulates mutualism and pathogenesis in Xenorhabdus nematophila. Appl. Environ. Microbiol. 73:7826–7836 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 22. Herbert E. E., Goodrich-Blair H. 2007. Friend and foe: two faces of Xenorhabdus nematophila. Nat. Rev. Microbiol. 8:634–646 [DOI] [PubMed] [Google Scholar]
  • 23. Heungens K., Cowles C. E., Goodrich-Blair H. 2002. Identification of Xenorhabdus nematophila genes required for mutualistic colonization of Steinernema carpocapsae nematodes. Mol. Microbiol. 45:1337–1353 [DOI] [PubMed] [Google Scholar]
  • 24. Hultgren S. J., Duncan J. L., Schaeffer A. J., Amundsen S. K. 1990. Mannose sensitive haemagglutination in the absence of piliation in Escherichia coli. Mol. Microbiol. 4:1311–1318 [DOI] [PubMed] [Google Scholar]
  • 25. Khandelwal P., Banerjee-Bhatnagar N. 2003. Insecticidal activity associated with the outer membrane vesicles of Xenorhabdus nematophilus. Appl. Environ. Microbiol. 4:2032–2037 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 26. Khandelwal P., et al. 2004. Insecticidal pilin subunit from the insect pathogen Xenorhabdus nematophila. J. Bacteriol. 186:6465–6476 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 27. Li X., Johnson D. E., Mobley H. L. 1999. Requirement of MrpH for mannose-resistant Proteus-like fimbria-mediated hemagglutination by Proteus mirabilis. Infect. Immun. 67:2822–2833 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 28. Madhusudhan K. T., Lorenz D., Sokatch J. R. 1993. The bkdR gene of Pseudomonas putida is required for expression of the bkd operon and encodes a protein related to Lrp of Escherichia coli. J. Bacteriol. 175:3934–3940 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 29. Martens E. C., Goodrich-Blair H. 2005. The Steinernema carpocapase intestinal vesicle contains a subcellular structure with which Xenorhabdus nematophila associates during colonization initiation. Cell. Microbiol. 12:1723–1735 [DOI] [PubMed] [Google Scholar]
  • 30. Martens E. C., Heungens K., Goodrich-Blair H. 2003. Early colonization events in the mutualistic association between Steinernema carpocapsae nematodes and Xenorhabdus nematophila bacteria. J. Bacteriol. 10:3147–3154 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 31. Meslet-Cladiere L. M., Pimenta A., Duchaud E., Holland I. B., Blight M. A. 2004. In vivo expression of mannose-resistant fimbriae of Photorhabdus temperata K122 during insect infection. J. Bacteriol. 3:611–622 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 32. Moureaux N., Karjalainen T., Givaudan A., Bourlioux P., Boemare N. 1995. Biochemical characterization and agglutinating properties of Xenorhabdus nematophilus F1 fimbriae. Appl. Environ. Microbiol. 7:2707–2712 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 33. Park D., et al. 2009. Genetic analysis of xenocoumacin antibiotic production in the mutualistic bacterium Xenorhabdus nematophila. Mol. Microbiol. 5:938–949 [DOI] [PubMed] [Google Scholar]
  • 34. Park D., Forst S. 2006. Co-regulation of motility, exoenzyme and antibiotic production by the EnvZ-OmpR-FlhDC-FliA pathway in Xenorhabdus nematophila. Mol. Microbiol. 6:1397–1412 [DOI] [PubMed] [Google Scholar]
  • 35. Park Y., et al. 2007. Clonal variation in Xenorhabdus nematophila virulence and suppression of Manduca sexta immunity. Cell. Microbiol. 9:645–656 [DOI] [PubMed] [Google Scholar]
  • 36. Sicard M., et al. 2004. Stages of infection during the tripartite interaction between Xenorhabdus nematophila, its nematode vector, and insect hosts. Appl. Environ. Microbiol. 11:6473–6480 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 37. Sicard M., Tabart J., Boemare N. E., Thaler O., Moulia C. 2005. Effect of phenotypic variation in Xenorhabdus nematophila on its mutualistic relationship with the entomopathogenic nematode Steinernema carpocapsae. Parasitology 5:687–694 [DOI] [PubMed] [Google Scholar]
  • 38. Simon R., Priefer U., Puhler A. 1983. A broad host range mobilization system for in vivo genetic engineering: transposon mutagenesis in Gram negative bacteria. Biotechnology (NY) 1:784–791 [Google Scholar]
  • 39. Smigielski A. J., Akhurst R. J., Boemare N. E. 1994. Phase variation in Xenorhabdus nematophilus and Photorhabdus luminescens: differences in respiratory activity and membrane energization. Appl. Environ. Microbiol. 60:120–125 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 40. Snyder H. 2010. Analysis of the regulation and role of fimbriae of Xenorhabdus nematophila and release process from the intestinal receptacle of Steinernema carpocapsae Ph.D. dissertation. University of Wisconsin—Milwaukee, Milwaukee, WI.
  • 41. Snyder H., Stock S. P., Kim S. K., Flores-Lara Y., Forst S. 2007. New insights into the colonization and release process of Xenorhabdus nematophila and the morphology and ultrastructure of the bacterial receptacle of its nematode host, Steinernema carpocapsae. Appl. Environ. Microbiol. 16:5338–5346 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 42. Somvanshi V. S., Kaufmann-Daszczuk B., Kim K.-S., Mallon S., Ciche T. A. 2010. Photorhabdus phase variants express a novel fimbrial locus, mad, essential for symbiosis. Mol. Microbiol. 77:1021–1038 [DOI] [PubMed] [Google Scholar]
  • 43. Stabb E. V., Ruby E. G. 2003. Contribution of pilA to competitive colonization of the squid Euprymna scolopes by Vibrio fischeri. Appl. Environ. Microbiol. 2:820–826 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 44. Tani T. H., Khodursky A., Blumenthal R. M., Brown P. O., Matthews R. G. 2002. Adaptation to famine: a family of stationary-phase genes revealed by microarray analysis. Proc. Natl. Acad. Sci. U. S. A. 99:13471–13476 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 45. Thanassi D. G., et al. 1998. The PapC usher forms an oligomeric channel: implications for pilus biogenesis across the outer membrane. Proc. Natl. Acad. Sci. U. S. A. 95:3146–3151 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 46. Tran E. E. H., Andersen A. W., Goodrich-Blair H. 2009. CpxRA influences Xenorhabdus nematophila colonization initiation and outgrowth in Steinernema carpocapsae nematodes through regulation of the nil locus. Appl. Environ. Microbiol. 75:4007–40014 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 47. van der Hoeven R., Betrabet G., Forst S. 2008. Characterization of the gut bacterial community in Manduca sexta and effect of antibiotics on bacterial diversity and nematode reproduction. FEMS Microbiol. Lett. 286:249–256 [DOI] [PubMed] [Google Scholar]
  • 48. Vigneux F., et al. 2007. The xaxAB genes encoding a new apoptotic toxin from the insect pathogen Xenorhabdus nematophila are present in plant and human pathogens. J. Biol. Chem. 282:9571–9580 [DOI] [PubMed] [Google Scholar]
  • 49. Vivas E. I., Goodrich-Blair H. 2001. Xenorhabdus nematophilus as a model for host-bacterium interactions: rpoS is necessary for mutualism with nematodes. J. Bacteriol. 183:4687–4693 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 50. Volgyi A., Fodor A., Forst S. 2000. Inactivation of a novel gene produces a phenotypic variant cell and affects the symbiotic behavior of Xenorhabdus nematophila. Appl. Environ. Microbiol. 66:1622–1628 [DOI] [PMC free article] [PubMed] [Google Scholar]

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