- Research article
- Open Access
Motility is required for the competitive fitness of entomopathogenic Photorhabdus luminescens during insect infection
© Easom and Clarke; licensee BioMed Central Ltd. 2008
- Received: 30 June 2008
- Accepted: 03 October 2008
- Published: 03 October 2008
Photorhabdus are motile members of the family Enterobactericeae that are pathogenic to insect larvae whilst also maintaining a mutualistic interaction with entomophagous nematodes of the family Heterorhabditiae. The interactions between Photorhabdus and its hosts are thought to be an obligate part of the bacteria's life-cycle in the environment. Motility often plays a key role in mediating bacteria-host interactions and, in this study, we were interested in characterising the role of motility in the Photorhabdus-nematode-insect tripartite association.
We constructed deletion mutants of flgG (blocking flagella production) and motAB (blocking flagella rotation) in P. luminescens TT01. Using these mutants we show that both the ΔflgG and ΔmotAB mutants are equally as good as the wild-type (WT) bacteria in killing insects and supporting nematode growth and development suggesting that flagella production and motility are not required for pathogenicity or mutualism. However we show that the production of flagella is associated with a significant metabolic cost during growth on agar plates suggesting that, although not required for pathogenicity or mutualism, there must be a strong selective pressure to retain flagella production (and motility) during the interactions between Photorhabdus and its different hosts. To this end we show that both the ΔflgG and ΔmotAB mutants are out-competed by WT Photorhabdus during prolonged incubation in the insect revealing that motile bacteria do have a fitness advantage during colonisation of the insect larva.
This is the first report of a role for motility in Photorhabdus and we show that, although not required for either pathogenicity or mutualism, motility does contribute to the competitive fitness of Photorhabdus during infection of the insect (and, to a lesser extent, the nematode). This adaptive function is similar to the role ascribed to motility in mammalian pathogens such as uropathogenic Escherichia coli (UPEC). Therefore, in addition to describing a role for motility in Photorhabdus, this study reinforces the relevance and utility of this bacterium as a model for studying bacteria-host interactions.
- Crystal Violet
- Bacterial Attachment
- Infective Juvenile
- Mutant Bacterium
- Competitive Fitness
Photorhabdus are Gram negative bacteria that are highly virulent pathogens of a wide variety of insect larvae whilst also maintaining a mutualistic interaction with nematodes from the family Heterorhabditidieae. Photorhabdus normally colonise the intestinal tract of the infective stage of the nematode, the infective juvenile (IJ). The IJs are a non-feeding stage of the nematode that lives in the soil where they actively seek out potential larval hosts. On finding a suitable host the IJ enters the larva and the bacteria are regurgitated into the insect hemolymph. Photorhabdus actively circumvent the insect innate immune system by inhibiting and adapting to the humoral response whilst also suppressing phagocytosis by circulating haemocytes [1–3]. During infection of the insect the bacteria grow exponentially, producing a wide range of toxins and hydrolytic exoenzymes that are responsible for the death and subsequent bioconversion of the insect larva into a nutrient soup that is ideal for nematode growth and development [4, 5]. The nematodes feed on the bacterial biomass within the insect cadaver and develop through juvenile (J1–J4) stages to form adult males and females. After several rounds of reproduction the J1 stage nematodes receive uncharacterised environmental cues that stimulate the development of IJs. At this point the developing IJs are colonised by Photorhabdus before they emerge from the insect cadaver to find new hosts (for recent reviews see [6, 7]).
Many bacteria are motile through the action of large complex protein assemblages called flagella. The production and function of flagella are best studied in the enteric bacteria Escherichia coli and Salmonella enterica serovar Typhimurium where it has been shown that the expression of genes required for flagella-mediated motility (and chemotaxis) is controlled by a complex regulatory network [8, 9]. Flagellum-mediated motility often plays a key role in mediating different bacteria-host interactions. For example motility is important during the colonisation of the squid by Vibrio fischeri and also during the infection of mammals by both Salmonella and E. coli [10–14]. Photorhabdus are motile through the action of peritrichously arranged flagella and we hypothesised that motility must play some role in the interactions between Photorhabdus and its invertebrate hosts. This was based on the principle that unused or redundant traits in bacteria will be lost over time . Therefore we constructed specific flagellum-minus (Fla-) and non-motile (Mot-) mutants of Photorhabdus and, using these mutants, we show that motility is not required for either pathogenicity or mutualism. However we do show that WT bacteria out-compete non-motile mutants during prolonged incubation in insect cadavers suggesting that motility confers a fitness advantage during colonisation of the insect.
Construction of mutations in flgG and motAB
Motility is required for attachment to surfaces
There is a cost associated with the production of flagella
Motility is required for the competitive fitness of Photorhabdusduring insect colonization
Motility affects the ability of Photorhabdusto colonise the nematode
Photorhabdus are highly virulent to a wide range of insect larvae and, following insect death the bacteria must remain in the insect cadaver at high densities for extended periods of time (up to 2–3 weeks under optimal conditions) to facilitate nematode growth and development. We have shown that, although neither flagella production nor motility is required for pathogenicity, there is a significant advantage to being motile during the normally prolonged incubation in the insect.
Our data show that, on agar plates, the ΔmotAB mutant is as competitive as the WT suggesting that there is no perceived benefit to being motile in this environment. This was not unexpected as the concentration of agar used would prevent cells from swimming or swarming, whether they are motile or not, thus rendering the ΔmotAB mutant neutral in this environment. On the other hand, the ΔflgG mutant has a considerable advantage over WT suggesting that there is a metabolic cost associated with the production of flagella. In a recent study Fontaine et al (2008) showed that the reduced mortality associated with a fliA mutation in E. coli was probably due to decreased internal cell stress due to the absence of physical destabilisation of the membrane . Therefore the cost associated with flagella production in Photorhabdus may be due to the utilisation of resources and energy for the production and/or function of the flagella or it may be due to the stresses associated with assembly of the flagella through the cellular membrane. In the insect this trend is reversed and the ΔflgG cells are now disadvantaged during extended growth (i.e. 21 days). This would suggest that flagella production benefits the bacteria during insect infection. Therefore the costs of flagella production (and presumably motility) appear to be offset during extended incubation in the insect. In support of this, the ΔmotAB mutant, that still bears the cost of producing flagella and yet does not derive any benefit associated with motility in the insect, is present at much lower levels than the ΔflgG mutant in competition assays after 21 days.
The frequency of motile cells observed throughout the growth of Photorhabdus under normal culturing conditions (i.e. shaking at 30°C in LB broth) is very low (<< 1%) (our unpublished data). During growth under static conditions the frequency of motility in the population increased reaching a maximum of 30% at 16 h post-inoculation before rapidly decreasing (data not shown). Therefore motility is restricted, both spatially and temporally, during the growth of Photorhabdus. This limited exposure of motility to any selection pressure in the environment could explain the relatively modest differences in fitness observed in this study.
In uropathogenic E. coli (UPEC) motility has been shown to contribute to the fitness of the bacteria during colonisation of the urinary tract . Indeed recent studies have shown that motility facilitates the movement of UPEC to the upper urinary tract . Moreover motility has been shown to be required for Salmonella to access high-energy nutrients found at sites of inflammation in the mammalian gut . In the same way we expect that motility in Photorhabdus, a closely related bacterium, will facilitate the movement of the bacteria throughout the insect cadaver thus enabling invasion of nutrient-rich niches that facilitate growth of the motile strain. In this regard Photorhabdus would also be expected to undergo chemotaxis and the genome is predicted to encode a complete Che signalling system in addition to 2 methyl-accepting chemotaxis proteins i.e. MCPs (see Fig. 1). However the role of chemotaxis was not investigated in this study.
We have shown that neither flagella production nor motility is required for the mutualistic association between Photorhabdus and the Heterorhabditis nematode. Photorhabdus has 2 roles during the interaction with the nematode: 1) the bacteria must provide nutrients for the nematode and 2) the bacteria must colonise the IJ. The nematodes feed on the Photorhabdus biomass that is present either in the insect or on agar plates and, therefore, the nematodes obtain a substantial part of their required nutrients directly from the bacteria. We did not see any differences in the growth and development of the nematodes growing on agar plates with the WT bacteria compared to the ΔflgG and ΔmotAB mutants. Indeed infecting insects with IJs colonised with either ΔflgG or ΔmotAB mutants resulted in an IJ yield (a quantitative indicator of the ability of the bacteria to support nematode growth and development) very similar to that observed with IJs colonised with WT bacteria (data not shown). Therefore flagella production and motility do not appear to play any role in the nutritional interaction between Photorhabdus and the nematodes either on agar plates or in the insect.
Photorhabdus are maternally transmitted from the hermaphrodite stage nematode (i.e. the mother) to the developing IJ . As the Heterorhabditis nematodes feed on Photorhabdus some viable bacteria enter the lumen of the gut of the hermaphrodite and attach to specific cells in the distal region of the gut (specifically the INT9 cells). The bacteria infect the neighbouring rectal glands cells and replicate within vacuoles. The rectal gland cells rupture, releasing Photorhabdus into the body cavity of the hermaphrodite where the bacteria encounter and colonise the developing IJs . The IJ is initially colonised by 1–2 bacteria that subsequently replicate, resulting in a final population of approximately 100 cfu of WT bacteria per IJ. The proportion of IJs colonised by the ΔflgG and ΔmotAB mutants is the same as the WT (i.e. approx. 80% in all cases) suggesting that attachment to the IJ, and presumably infection of the hermaphrodite, is independent of flagella and/or motility. On the other hand, the final population level of Photorhabdus within the IJ is significantly altered in nematodes that are cultured on either the ΔflgG or the ΔmotAB mutant strain. Therefore IJs that have been grown on the ΔflgG mutant carry, on average, a bacterial population that is 50% greater than the population within IJs cultured on WT bacteria. In contrast the ΔmotAB mutant does not reach population levels within the IJ that are equivalent to the WT suggesting that the production of non-functional flagella negatively influences growth in the nematode. These results might be explained in terms of the metabolic cost associated with the production of flagella. Therefore, in the absence of flagella production, the ΔflgG mutant may be able to put more of the limited resources available within the nematode into biomass production and division. On the other hand the ΔmotAB mutant still bears the cost of producing flagella although these are non-functional. Interestingly the fact that the ΔmotAB mutation is not neutral, in terms of IJ colonisation, suggests that Photorhabdus are likely to be motile at some point during the colonisation of the nematode. The bacteria initially colonise the proximal end of the IJ gut and one possibility is that motility may facilitate exploration of the distal regions of the gut thus allowing the bacteria to make better use of the limited resources available within the nematode.
In this study we show that there is a significant metabolic cost associated with the production of flagella (and motility) in Photorhabdus. Nonetheless Photorhabdus are motile suggesting that motility is an adaptive trait that is under powerful positive selection in the environments where Photorhabdus is normally found i.e. in the insect and nematode. In this study we show that, although motility is not required for either pathogenicity or mutualism, this trait is advantageous during the interactions between Photothabdus and both of its invertebrate hosts. Therefore, in addition to describing a role for motility in Photorhabdus, this work also highlights the functional overlap between pathogenicity and mutualism and reinforces the utility of Photorhabdus as a model for studying these different bacteria-host interactions.
Bacterial strains and culture conditions
A spontaneous rifampicin-resistant mutant of Photorhabdus luminescens subsp. luminescens was used as the wild-type (WT) in all experiments . The bacteria were cultured in LB broth or on LB agar (LB broth plus 1.5% (w/v) agar) at 30°C for P. luminescens. Unless otherwise stated all LB agar plates were supplemented with 0.1% (w/v) pyruvate . Escherichia coli S17-1 (λpir) and E. coli EC100 (Epicentre) were cultured at either 30°C or 37°C as indicated. Swim agar is LB broth plus 0.3% (w/v) added agar. When required antibiotics were added at the following concentrations: ampicillin (Ap), 100 μg ml-1; chloramphenicol (Cm), 20 μg ml-1 and rifampicin (Rif), 50 μg ml-1.
Construction of deletion mutants
Primers used for the construction of the ΔflgG and ΔmotAB mutants
Sequence (5' – 3') a
Cloning of the flgG and motABgenes
The flgG and motAB genes were amplified from P. luminescens TT01 genomic DNA by PCR using KOD polymerase. The primers used for flgG were CAT001 (5'-TAAAACCCATGGTCCGATCATTATGGATTGC-3') and CAT002 (5'-GCTGGATCTAGATTATAACTGAGTCAGTTTTTGTAGC-3') and for motAB CAT003 (5'-GATATCCCATGGTAGTACTTTTAGGATATATC-3') and CAT004 (5'-GCAGTGTCTAGATTACTTTGTCACCTTGGTCGG-3'). The flgG and motAB PCR fragments were digested with NcoI and XbaI and cloned into pTRC99a (Amersham Pharmacia Biotech) resulting in pBMM800 and pBMM802, respectively. The integrity and accuracy of all plasmid clones was verified by DNA sequencing.
The capacity of P. luminescens to form biofilms was assessed by measuring bacterial attachment to a plastic surface . Strains were grown overnight in LB broth, diluted to OD600= 0.05 in fresh LB and 200 μl of the cell suspension was aliquoted in triplicate, into the wells of a Costar® polypropylene (PP) 96-well microtitre plate. The plates were sealed with a gas permeable membrane and incubated, without shaking in a saturated environment to prevent evaporation, at 30°C. At the appropriate time the planktonic cells were removed by aspiration and the wells were washed with 1× phosphate buffered saline (PBS). To observe biofilm formation 250 μl of 0.1% (w/v) crystal violet (CV) was added to each well and the plates were incubated at room temperature for 20 min before rinsing 3 times with 1 × PBS. To quantify biofilm formation the CV was dissolved in 250 μl of 95% ethanol and the CV concentration was determined by measuring the OD595 using a Genios (Tecan) plate reader.
The pathogenicity of P. luminescens was assessed using Galleria mellonella larvae (the Greater Wax Moth) as the model insect host. The G. mellonella larvae were purchased from Livefood (UK). Briefly overnight cultures of P. luminescens TT01 were washed 3 times in 1 × PBS before the OD600 was adjusted to 1.0 as this has been shown to be equivalent to 4 × 108 cfu ml-1 (our unpublished data). The culture was diluted with 1 × PBS to give cell density of 2 × 104 cfu ml-1 and 10 μl (equivalent to 200 cfu) was injected into the hemolymph of a G. mellonella larva using a Hamilton syringe and a BD Microlance™ 3 30G × 1/2" needle. For competition assays the WT and mutant strain were grown overnight in LB at 30°C and equal numbers of cells from each culture were mixed and subsequently diluted so that 200 cfu could be injected into each insect larva. The proportion of WT and mutant bacteria in the injection mixture was assessed by plating an aliquot of the mixture onto LB (Rif) agar followed by patching 100 colonies onto swim agar. To determine the proportion of WT and mutant bacteria in the insect an infected insect larvae was surface sterilised by dipping in ethanol and passing the insect through a Bunsen flame before quickly plunging it into 5 ml sterile 1 × PBS in a universal tube. The insect cadaver was sliced open using a sterile scalpel blade and homogenised by adding 7 (3–4 mm) sterile glass beads to the tube, followed by vortexing for 2 min. The supernatant, containing the bacteria, was plated onto LB (Rif) agar and the proportion of motile/non-motile bacteria was determined by patching 50 colonies onto swim agar.
In vitrosymbiosis assays
An aliquot of 50 μl of an overnight culture diluted to an OD600 = 1.0 of the appropriate bacteria was spread, in a Z pattern, onto the surface of a lipid agar plate using an inoculating loop. The plates were incubated at 30°C for 3 days before adding 50 surface sterilised IJ nematodes to the bacterial biomass. Nematodes were surface-sterilised by washing in a solution (0.4% (w/v)) of hyamine (Sigma). Nematode recovery was assessed 7 days after addition of IJs by counting the number of hermaphrodites on the lipid agar plate. The new generation of IJs migrate to the lid of the Petri dish and, after 21 days, these nematodes were collected, by washing the lid with PBS to a final volume of 50 ml, and the number of IJs present (i.e the IJ yield) was determined. In competition assays the assays were the same except that the lipid agar plates were inoculated with equal numbers of the WT and mutant bacteria. At 3 and 21 days post-inoculation the bacterial biomass was scraped off the plate and the proportion of motile/non-motile bacteria was determined (as before). Colonisation levels in the IJ were determined by crushing single, surface-sterilised IJ nematodes in 100 μl 1 × PBS using a hand-held homogeniser and plating the homogenate onto LB (Rif) agar.
The work outlined in this study was carried out equally in the University of Bath and University College Cork. The research was funded through the Exploiting Genomics initiative of the BBSRC in the UK (86/EGA16183) and Science Foundation Ireland. CAE is supported by a PhD fellowship from the University of Bath. The authors would also like to thank Susan Joyce for technical help and advice during this study.
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