Contaminant biodegradation in soil is frequently limited by hindered physical access of bacteria to the contaminants. In the frame of the development of novel bioremediation approaches based on ecological principles, we tested the hypothesis that fungal networks facilitate the movement of bacteria by providing continuous liquid films in which gradients of chemoattractants can form and chemotactic swimming can take place. Unlike bacteria, filamentous fungi spread with ease in water-unsaturated soil. In a simple laboratory model of a water-unsaturated environment, we studied the movement of polycyclic aromatic hydrocarbon-degrading Pseudomonas putida PpG7 (NAH7) along a mycelium of Pythium ultimum. Some undirected dispersal was observed in the absence of a chemoattractant or when the non-chemotactic derivative strain P. putida G7.C1 (pHG100) was used. The bacterial movement became fourfold more effective and clearly directed when the chemotactic wild type was used and salicylate was present as a chemoattractant. No dispersal of bacteria was found in the absence of the fungus. These findings point at a role of mycelia for the translocation of chemicals and microorganisms. The results suggest that fungi improve the accessibility of contaminants in water-unsaturated environments.
The capacity of microorganisms to rapidly sense and adapt to environmental changes is an important factor for the stability of microbial ecosystem services, such as contaminant biodegradation in soil (de Lorenzo, 2008; Miller et al., 2009). Marx and Aitken (2000), for instance, have shown that bacterial chemotaxis, i.e. the ability to sense and move along chemical concentration gradients, enhances the biodegradation of naphthalene, a polycyclic aromatic hydrocarbon (PAH), in bioavailability-limited heterogeneous systems (for a review: Harms and Wick, 2006; Miller et al., 2009). Bacterial motility and chemotaxis in porous media, however, are still a field full of unknowns and experimental evidence for its importance is limited (Ford and Harvey, 2007). The efficiency of bacterial chemotaxis in bioremediation strongly depends on the effective mobility of bacteria within the system. As bacteria require high soil matric potentials or at least continuous liquid films for swarming and swimming (Or et al., 2007), directed chemotactic dispersal is restricted in water-unsaturated environments. This may affect the bioaccessibility of patchy, hydrophobic organic compounds (Semple et al., 2007), as average distances between bacterial microcolonies in soil are supposed to be in the range of 10−4 m (Bosma et al., 1997). Microbial dispersal and substrate mobilization are hence needed to overcome the distance between substrates and organisms. The strategy of filamentous fungi is to enlarge their external surface and to develop mycelia of high fractal dimension that optimally exploit the three-dimensional space containing the substrate (Nakagaki et al., 2004). Contrary to bacteria, the habitat of fungi is not restricted to water films. Fungal hyphae easily breach through air–water interfaces and form dense networks of up to 20 000 km length per cubic meter of soil (Pennisi, 2004). Importantly, by doing so they connect saturated and unsaturated soil pores (Wessels, 1997). Several reports on the role of fungal mycelia on ‘underground networking’ for nutrient translocation and provision to bacteria in the hyphosphere (Bending et al., 2006) and shaping of communities above and below the earth's surface (Whitfield, 2007) have been published. As both bacteria and fungi are important degraders of (anthropogenic) organic substances in soil, fundamental knowledge of bacteria–fungus interactions is also essential for the development of novel bioremediation approaches based on ecological principles. For instance, it has been shown that liquid films developing around hydrophilic fungi can be used by PAH-degrading bacteria to enhance their mobility in such a way that PAH-biodegradation in unsaturated soil is enhanced (Kohlmeier et al., 2005; Wick et al., 2007).
In the frame of our attempts to develop novel bioremediation approaches based on ecological principles we tested the hypothesis that fungal networks facilitate the movement of bacteria by providing continuous liquid films in which gradients of chemoattractants can form and chemotactic swimming can take place.
Chemotaxis and motility tests
Chemotactic response was tested in capillaries containing sodium salicylate. The chemotactic strain PpG7 exhibited a statistically significant (twofold) attraction to the chemoattractant relative to the salicylate-free control. The non-chemotactic derivative G7.C1 showed no chemotactic response to salicylate (P = 0.01). Both strains showed equal (P < 0.05) swimming and swarming motility on soft agar plates with colony diameters of 2.3 ± 0.3 and 2.8 ± 0.1 cm obtained after 24 and 48 h on swimming agar plates and 5.3 ± 0.5 and 6.3 ± 1.1 cm after 24 and 44 h on swarming agar plates respectively.
Effect of salicylate on fungus-mediated dispersal of Pseudomonas putida
Chemotactic movement of bacteria along fungal networks was determined in linear arrangements of agar-patches and gaps (air-filled) separating them (Fig. 1). The ‘test track’ included a central and two terminal patches of potato dextrose agar (PDA) and could thus be completely overgrown by fungal mycelia. These circular patches framed rectangular patches on which bacterial distributions were determined. High-purity agarose free of nutrients prevented bacteria from growing there. Chemoeffectors could be placed on the L3 and R3 positions of the agarose patches and bacterial inocula on the middle patch. Depending on both the presence and the placement of the chemical (one side, both sides, control without chemical) this allowed observing and quantifying random movement and positive or negative chemotaxis of the bacteria respectively. Three configurations were tested for each of the strains: 5 mg sodium salicylate was either placed at position R3 (set-up: Sal −/+), at positions L3 and R3 (set-up: Sal+/+), or it was omitted (set-up: Sal −/−; Fig. 1).
In the absence of a fungal mycelium no dispersal of chemotactic and non-chemotactic bacteria beyond the gap surrounding, the middle patch was observed (data not shown). By contrast, about 0.5–1% [ca. 106−107 colony-forming units (cfu)] of the inoculum moved over the gaps when a mycelium of Pythium ultimum crossed them. Figure 2 summarizes the distribution of those bacteria that passed over the gap next to the middle batch within 48 h. In salicylate-free controls, a quasi uniform distribution of strain PpG7 (P < 0.05) in the L3 and R3 positions was observed (Fig. 2A). Similar results, but with a significant increase in the more distal parts of the agarose patches, were obtained when salicylate was put on both sides (Fig. 2B). Salicylate placed in the R3 position clearly attracted strain PpG7 to the right side (> 70% of the recovered cells, P = 0.0034) with a maximum accumulation in position R1 (Fig. 2C). Phenanthrene, a substrate of strain PpG7 not acting as chemoattractant, placed in the same position did not attract strain PpG7 (not shown). Considerable dispersal, yet no directed movement was observed for the chemotaxis-deficient strain G7.C1 when salicylate was put on the R3 position (Fig. 2D), on either sides or without salicylate (not shown). Quantitative balances, including the cells remaining on the middle patch, revealed recovery rates of 51 ± 16% and 42 ± 20% of the inoculated PpG7 in presence (n = 9) and absence (n = 6) of salicylate (Fig. 3). This indicates that no growth-related bias on the mobilization had to be expected in presence of the chemoeffector. There was no significant influence (P < 0.05) of the chemoeffector on the overall mobilization rate of both strains tested (Fig. 3).
Visualization of fungus–bacteria interactions
Confocal laser scanning microscopy (CLSM) was employed for visualizing the hypothesized continuous liquid films forming along the mycelia of P. ultimum growing through air-filled space. For this purpose P. ultimum was inoculated on a PDA patch placed on a microscope slide in order to let the mycelia overgrow the dry and clean glass surface. Subsequent staining of the fungal cell surface proteins with fluorescein isothiocyanate (FITC) allowed analysing the gross fungal morphology, whereas an overlay of the reflection and the FITC signals of the hyphae allowed visualization and assessment of the thickness (3–4 μm) of the synaeretic liquid film forming along the filaments of P. ultimum (Fig. 4A). CLSM was further used to visualize the dispersal of P. putida PpG7 along the filaments P. ultimum. For this purpose the bacteria were stained with the fluorescent dye (Syto 24), subsequently, spot inoculated on the fungal filaments and the bacterial dispersal continuously analysed for 2 h by simultaneous recording of the reflection signal of the filamentous network and the fluorescent signal of the bacteria. Analysis of the time series recorded demonstrated that the synaeretic films allowed for bacterial dispersal along the mycelia of P. ultimum, as depicted in the CLSM maximum intensity projection in Fig. 4B. Due to experimental hardware/software limitations, however, the directed dispersal of the bacterial cells along the fungal filaments could not be quantified.
Translocation of sodium salicylate along the mycelium of P. ultimum
Chemotaxis requires the development of a chemical gradient. The spatial distribution of sodium salicylate 48 h after addition to position R3 was analysed. In the presence of P. ultimum, salicylate was found along the entire network, also beyond both gaps around the middle patch (Fig. 5). In the absence of a mycelium, sodium salicylate was detected solely in positions R3 to R1. Degradation studies showed that P. ultimum was unable to metabolize sodium salicylate. No decrease of salicylate concentration was detected and no growth was observable when P. ultimum was incubated > 7 days with salicylate as sole carbon and energy source in shaken liquid cultures (data not shown).
Chemotactic navigation along mycelia of P. ultimum
Recent studies revealed that fungal hyphae can promote microbial dispersal and accelerate biotransformation in soil (Kohlmeier et al., 2005; Wick et al., 2007). Using this effect for improving microbial contaminant degradation, however, would benefit from a comprehensive, mechanistic understanding of the dispersal process. In this study, we hence tested the hypothesis that continuous liquid films along fungal mycelia passing through air-filled space may allow for the build-up of chemical gradients and promote the chemotactic dispersal of otherwise immobilized bacteria. Sodium salicylate was selected as known chemoattractant of PAH-degrading P. putida PpG7 (NAH7) (Velasco-Casal et al., 2008). Criteria for the choice of salicylate were its environmental relevance as metabolite and inducer of microbial PAH-degradation, its high water solubility and the absence of degradation by P. ultimum. The observed directed dispersal of strain PpG7 (yet not of its non-chemotactic derivative G7.C1) in the presence of mycelia of P. ultimum thus suggests that liquid water films supporting chemotactic swimming formed along the mycelia. This was unequivocally confirmed by CLSM visualization clearly showing the liquid film along the fungal filaments and bacterial motility within such liquid films respectively. At this point it remains open if the chemical gradient required for chemotaxis developed only in extra-hyphal liquid films or if its build-up was also promoted by the hyphae themselves. The accumulation of strain PpG7 near the chemoattractant cannot be explained by advective transport along the hyphae as: (i) bacteria have to be motile to be transported along P. ultimum (Wick et al., 2007), (ii) the similarly motile but non-chemotactic derivative G7.C1 showed no preferred movement to salicylate and (iii) no preferred accumulation of strain PpG7 near growth substrates not acting as chemoattractant (e.g. phenanthrene) was observed. Directed navigation to sodium salicylate is in good agreement with results from classical capillary experiments showing two to fourfold enhanced transport (this study, Velasco-Casal et al., 2008). No influence of salicylate on overall bacterial mobility (0.5–1%) was found (Fig. 3). This may indicate that under non-growth conditions chemical gradients influence the direction but not the extent of bacterial swimming. It may also be that the fraction of transported bacteria represented basically all those on the middle patch that could get access to the liquid films in fungal mycelium and make use of the chemical gradient developing therein. As a further possibility the capacity of water films to facilitate active movement of bacteria may simply be limited and/or further restricted by mechanisms such as bacterial attachment to hyphae and swimming into dead ends, etc., although the nearly homogeneous distribution of bacteria in the Sal +/+ and Sal −/− set-up seems to contradict an importance of the latter mechanisms.
Relevance for the ecology of contaminant biodegradation
In terrestrial environments, fungi are of fundamental importance as decomposer organisms and plant symbionts, playing pivotal roles in the carbon, nitrogen and phosphorous cycles. In most natural soil habitats, the spatio-temporal distribution of nutrients and minerals is heterogeneous due to their distinct and complex physical structures (Boswell et al., 2003). It is thought that many fungi are able to grow in low-nutrient or polluted habitats by translocating resources available in other parts of the mycelium. There is currently evidence of passive translocation (diffusion-driven) and active translocation (metabolically driven) of nutrients (Boswell et al., 2002) by mycorrhizal fungi. Recent studies have shown that the underground fungal networks link plants together by transferring nutrients from plant to plant (Whitfield, 2007) or provide nutrients to bacteria in their hyphosphere and hence shape soil microbial communities. As not much is known about the translocation of substances other than nutrients in non-mycorrhizal fungi, we compared the observed translocation of sodium salicylate in the presence of P. ultimum within 48 h with the calculated efficiency of aqueous diffusion. During this time diffusion of salicylate should be restricted to less than 2 cm. However, we found significant concentrations of salicylate behind the gap on the middle patch and minor concentrations of salicylate on positions L1–3, which is between 1.6 cm and more than 4 cm away from the source at position R3. As salicylate is non-volatile, the transport over relatively long distances may indeed point at a mechanism of intrahyphal transport.
The observed directed transport of bacteria through air-filled space suggests that soluble chemicals diffusing along liquid layers on fungal hyphae can be sensed by chemotactic bacteria and allow them to rapidly navigate towards substrate-rich microhabitats. This may be of particular importance for the establishment of plant–microbe interactions in the rhizosphere or for the spreading of bacteria in soil pores (Alexandre et al., 2004). Whereas dispersal of bacteria is generally constrained to matric potentials above −50 kPa (Paul and Clark, 1996), fungal subsurface movement is not restricted to fully water-filled pathways (Ritz, 1995). As fungi are predominantly aerobic microorganisms that tend to oxidize hydrophobic organics and break-down polymers by the use of extracellular enzymes (Ekschmitt et al., 2008), one may speculate that they do not only render substrates more soluble but also transfer them to bacterial commensals in the hyphosphere (Johnsen et al., 2005; Wick et al., 2009).
In defined systems, bacterial chemotaxis has been shown to improve rates of biodegradation (Marx and Aitken, 2000), given that the rate-limiting factor is the slow effective mass transfer of the pollutant. Chemotactic dispersal coefficients are two- to three orders of magnitude greater than random motility coefficients of bacteria (Ford and Harvey, 2007). From our observations we can conclude that fungal mycelia transform water-unsaturated porous media into microhabitats, in which bacterial chemotaxis can function and thus contribute to improved accessibility of chemicals, such as environmental contaminants. The likely consequence is increased contaminant degradation and easier build-up and maintenance of contaminant-degrading bacterial communities.
Organism and culture conditions
Pseudomonas putida PpG7 (NAH7) (Dunn and Gunsalus, 1973) and its derivative P. putida G7.C1 (pHG100) (Grimm and Harwood, 1997) were used throughout this study. Whereas the parent strain PpG7 is known to be chemotactic towards naphthalene and salicylate, strain P. putida G7.C1 (pHG100) is non-chemotactic (Velasco-Casal et al., 2008). Further criteria for the choice of these organisms were insignificant differences in motility, as well as in physico-chemical surface properties and deposition efficiencies in sand-filled columns (Velasco-Casal et al., 2008). Both strains are motile by means of polar flagella and able to use naphthalene and salicylate as the sole carbon and energy source. Naphthalene degradation kinetics of both strains in well-mixed conditions are comparable (Marx and Aitken, 2000). The strains were grown at 25°C on a gyratory shaker (130 r.p.m.) in 300 ml Erlenmeyer flasks containing 100 ml of liquid minimal medium (Wick et al., 2001) in the presence of 1.5 g of solid naphthalene (> 98%, Fluka; crystals taken as obtained by the provider). Cells used in mobilization experiments were harvested in the late exponential phase after about 48 h of growth, washed three times in cold 10 mM potassium phosphate buffer (PB) at pH = 7.2, and suspended in 10 mM PB to obtain a bacterial suspension containing ca. 5 × 109−1 × 1010 cells ml−1. In mobilization experiments, bacteria were quantified as cfu on Luria broth (LB) agar (2% w/v) containing 200 mg l−1 of actidione (cycloheximide). Fungus-associated bacteria were detached by sequential vortexing (60 s at 40 Hz) and ultrasonication (2 × 30 s) in PB prior to cultivation. The fast-growing, hydrophilic (Smits et al., 2003; Wick et al., 2007) oomycete P. ultimum was cultivated at room temperature on solid medium containing 2% (w/v) PDA (Difco) in the dark.
Swimming and swarming tests
The motility of bacterial strains was tested using a standard procedure (Rashid and Kornberg, 2000) based on soft agar plates: swimming agar consisted of (l−1): 10 g of tryptone (Oxoid), 10 g of yeast extract (Oxoid), 12.5 g of NaCl (Merck) and 3 g of agar (Difco). Swarming agar was prepared from (l−1): 8 g of nutrient broth (Oxoid), 5 g of glucose and 5 g of agar. Freshly poured agar plates were dried for 1 h (swimming agar) and 14 h (swarming agar) at ambient temperature prior to spot-inoculation with a sterilized toothpick. Motility was examined by measuring the mean diameter of the bacterial colonies after 24–48 h of incubation.
Degradation of salicylate by P. ultimum
Salicylate biodegradation by P. ultimum was studied by adding 500 μl of a dense mycelium suspension to amber 250 ml bottles containing 1 g l−1 of sodium salicylate (Merck) dissolved in 100 ml of GASnM medium (Crowe and Olsson, 2001). The fungal suspension was prepared by transferring PDA-grown mycelia of a 3-day-old P. ultimum to 10 ml GASnM medium and subsequent vortexing for 60 s at 40 Hz. Samples without fungal mycelium and/or GASnM medium without glucose and asparagine, respectively, were used as controls. Biodegradation of sodium salicylate was quantified for 7 days by a standard procedure (Trinder, 1954). Briefly, 1 ml of the analyte solution was mixed with the Tinder's reagent in the absence of mercuric chloride and the concentration of sodium salicylate in iron–salicylate complex was quantified spectrophotometrically at 540 nm. Sodium salicylate standards with known concentrations were used for calibration.
Capillary chemotaxis experiments
A modified version of the capillary tests described by Adler (1973) was used to quantify chemotactic response of strain G7.C1 to sodium salicylate. In short, late exponential cells were harvested, centrifuged and re-suspended in minimal medium to an OD578 of ca. 0.020 (corresponding to 106 cells ml−1). About 0.1 ml of this suspension was placed in a small chamber formed between two capillary tubes placed parallel to each other on a microscope slide. Another capillary tube (1 μl volume), heat-sealed at one end, containing the chemoeffector solution, was immersed in the cell suspension with its open end. The system was then closed with a glass coverslip, avoiding any formation of air bubbles in the chamber. The chemoeffector solution in the capillary contained 10 mM PB supplemented with 100 mM of salicylate. Minimal medium lacking any chemoeffectors was used as a control. The chambers were incubated for 1 h at room temperature and the numbers of bacterial cells accumulated in the test capillaries quantified as cfu on LB agar.
Chemotaxis experiments on fungal networks
Set-up and fungal growth. Chemotactic response of strains PpG7 and G7.C1 moving along fungal networks towards chemoeffector gradients was determined in a controlled laboratory system mimicking a combination of water-saturated (agar surface) and non-saturated (air gap between the agar) microhabitats (Fig. 1): a patch of PDA (diameter: 1.4 cm, height: 0.4 cm) inoculated with P. ultimum was placed in the centre of a sterile plastic Petri dish. Two agarose rectangles [(l) × (w) × (h): 1.5 × 1 × 0.4 (cm)] containing 10 mM of PB were placed next to it allowing for a 1 mm air-filled gap between the agarose and the middle patch. This gap prevented the bacteria from leaving the middle patch in absence of fungi. For the rectangles, nutrient- and carbon-deficient agarose (Sigma) was used to limit bacterial growth during the experiment. At the outer end of each agarose rectangle, a piece of PDA (diameter: 1.4 cm, height: 0.4 cm) was placed to promote the growth of fungal mycelium over the entire arrangement including the agarose rectangles, where no substrate was present. The Petri dishes were closed and incubated at the room temperature in the dark. After the fungus had fully overgrown the system (ca. 5 d), the outer PDA patches were removed and 5 mg of solid sodium salicylate was added unilaterally at the outer end of one of the agarose rectangles (position R3 in Fig. 1). Two control experiments were performed by either placing no sodium salicylate or by placing salicylate at positions L3 and R3 (Fig. 1). Forty-eight hours after the addition of sodium salicylate, 5 × 109 cells m l−1 (as determined as cfu on LB agar) suspended in 5 μl 10 mM PB were applied onto the fungal inoculum on the middle patch with a sterilized 25 μl microsyringe. Experiments without P. ultimum served as controls to exclude bacterial dispersal to the agarose rectangles in the absence of fungal hyphae.
Quantification of bacterial transport efficiency. Forty-eight hours after the addition of the bacteria, their spatial distribution was assessed. The time point was chosen as the time needed for salicylate to diffuse through the entire rectangle on which it was placed. The calculation was: with td, L and D representing the diffusion time (s), the diffusion length (cm) and the diffusion constant (Dsalicylate = 1.2 × 10−5 cm2 s−1) (Schwarzenbach et al., 2003). The agarose rectangles were subsequently cut into three individual sections of 0.5 cm length. Each of them, as well as the middle patch, were placed in sterile glass test tubes containing 2 ml 10 mM PB. Surface-associated bacteria were subsequently detached from the fungal mycelia and the agar as described above and quantified as cfu on LB agar containing 200 mg l−1 of actidione (cycloheximid) to suppress fungal growth. Distribution of salicylate on the agarose after 48 h was measured using the Trinder method as described above.
Confocal laser scanning microscopy
The CLSM observations were performed with a TCS SP1 (Leica) attached to an upright microscope equipped with a 63× NA 0.7 air and 63× NA 0.9 water immersion objectives. The instrument was controlled by the Leica Confocal Software Version 2.61 Build 1537. For excitation the laser lines at 488, 561 and 633 nm were available. The microscope was used for imaging the transmission, reflection, autofluorescence and fluorescence signals. The samples were screened for protein, polysaccharides, nucleic acids, lipids and lectin binding (Staudt et al.) stains. In Fig. 4A the fungal filament surfaces were stained with FITC (Research Organics) and in Fig. 4B the bacterial nucleic acids with Syto 24 (Invitrogen). The sample was excitated at 488 nm, and emission signals were detected at 480–500 nm (reflection) and 500–550 nm (FITC and Syto 24). Image data were projected using the microscope software and Imaris 6.3 (Bitplane). In order to improve signal to noise ratio, the data set in Fig. 4A was subjected to blind deconvolution using the classic MLE algorithm in Huygens 3.0.0 (SVI). Images were finally printed from Photoshop (Adobe) without any further adjustments.
Unless otherwise stated, all P-values are < 0.05 as determined based either on Student's t-test or one-way anova.
This study was supported by the European MC-EST 20984 grant (RAISEBIO). It contributes to the CITE research programme of the Helmholtz Association. Technical help by R. Remer, B. Würz, J. Reichenbach, M. Kolbe, K. Lübke and U. Kuhlicke is greatly acknowledged. We further thank Prof. C. S. Harwood (University of Washington) for the provision of P. putida G7.C1 (pHG100).