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/lp/ou_press/bacterial-biofilm-formation-on-the-hyphae-of-ectomycorrhizal-fungi-a-g0SAixxpB2
- Publisher
- Oxford University Press
- Copyright
- © FEMS 2018.
- ISSN
- 0168-6496
- eISSN
- 1574-6941
- DOI
- 10.1093/femsec/fiy093
- Publisher site
- See Article on Publisher Site
Abstract
ABSTRACT Ectomycorrhizal (ECM) fungi establish symbiosis with roots of most trees of boreal and temperate ecosystems and are major drivers of nutrient fluxes between trees and the soil. ECM fungi constantly interact with bacteria all along their life cycle and the extended networks of hyphae provide a habitat for complex bacterial communities. Despite the important effects these bacteria can have on the growth and activities of ECM fungi, little is known about the mechanisms by which these microorganisms interact. Here we investigated the ability of bacteria to form biofilm on the hyphae of the ECM fungus Laccaria bicolor. We showed that the ability to form biofilms on the hyphae of the ECM fungus is widely shared among soil bacteria. Conversely, some fungi, belonging to the Ascomycete class, did not allow for the formation of bacterial biofilms on their surfaces. The formation of biofilms was also modulated by the presence of tree roots and ectomycorrhizae, suggesting that biofilm formation does not occur randomly in soil but that it is regulated by several biotic factors. In addition, our study demonstrated that the formation of bacterial biofilm on fungal hyphae relies on the production of networks of filaments made of extracellular DNA. biofilms, eDNA, ectomycorrhizal symbiosis, fungal/bacterial interactions INTRODUCTION Among the myriad of organisms that live in forest soils, bacteria and fungi largely exceed their counterparts in terms of abundance and diversity (Nazir et al. 2010). Both highly contribute to the decomposition of soil organic matter and to the nutrient cycling and thus have a key role in the modulation of soil fertility and productivity (Rousk and Bengtson 2014; Lindahl and Tunlid 2015). In addition, some mutualistic fungi called ectomycorrhizal fungi (ECM), act as providers of carbon sources to the soil and of nutrients to the trees through the symbiosis they establish with roots (Heijden, Martin and Selosse 2015). ECM fungi provide a habitat for specific and complex bacterial communities that physically and metabolically interact with the fungi (Frey Klett, Garbaye, Tarkka 2007; Warmink, Nazir and van Elsas 2009; Marupakula, Mahmood and Finlay 2016). Bacteria are thought to gain two main benefits from this association. First, the hyphosphere—i.e. the area surrounding hyphae and under their metabolic influence—provides a nutritional source for bacteria that either consume nutrients released directly or indirectly by hyphae, or directly prey on fungi (Nazir et al. 2010; Ballhausen, Vandamme and de Boer 2016). Second, fungal hyphae can serve as vectors for bacteria to travel across the soil and to reach otherwise inaccessible nutrient sources (Nazir et al. 2010). These so called 'hyphal highways', can be followed by bacteria that swim along the water film that covers the hyphae, or by bacteria that settle at the tip of the growing hyphae (Warmink and van Elsas 2009; Otto et al. 2016). Conversely, some fungi can benefit from the metabolic activity of their associated bacteria (Li et al. 2016), gain protection against stresses (Nazir, Tazetdinova and van Elsas 2014) or even 'farm' bacteria to later use them as a source of nutrients (Pion et al. 2013). However, this close interaction between fungi and bacteria can also be detrimental to the fungi and a number of them produces defensins to prevent the bacterial colonization of their hyphae (Essig et al. 2014). Bacteria can establish in the hyphosphere in four states: as free-living cells, as attached single cells, as endohyphal cells or as organized biofilms. Depending on the nature of the physical interaction established between bacteria and fungi, the mechanisms of the interaction as well as their outputs differ (Frey-Klett et al. 2011; Deveau et al. 2018). Biofilms arise through the aggregation of bacterial cells and their embedding into a self-produced matrix of extracellular polymeric substances (Flemming et al. 2016). Life as a biofilm has the double advantage to increase the bacterial resistance against biotic and abiotic stresses, and to permit the organization of cells into functional sub-communities. As a consequence, a large number of bacterial species have developed the ability to build biofilms on hydrated abiotic surfaces (e.g. water pipes, medical devices) but also on living tissues (e.g. epithelial cells, root surfaces). Fungal hyphae can also support bacterial biofilms, and in vitro formation of bacterial biofilms on the hyphae of soil Ascomycetes, Basidiomycetes and Zygomycetes has been reported (Scheublin et al. 2010; Burmølle, Kjøller and Sørensen 2012; Nazir, Tazetdinova and van Elsas 2014; Hover et al. 2016). These studies mainly focused on saprophytic or arbuscular mycorrhizal fungi and little is known regarding biofilm formation of hyphae of ECM fungi (Warmink and van Elsas 2009; Ul Haq et al. 2014). Yet ECM fungi could provide large surfaces for bacteria to establish as biofilms thanks to their extended networks of hyphae that colonize large volumes of soil. In the present work, we focused on a specific group of ECM associated bacteria, so called Mycorrhiza Helper Bacteria (MHB) for their ability to promote the establishment of ECM symbiosis (Frey-Klett, Garbaye and Tarkka 2007; Deveau and Labbé 2017). We have shown in previous studies that the MHB strain Pseudomonas fluorescens BBc6 is chemoattracted by the hyphae of the ECM fungus Laccaria bicolor S238N on which it forms biofilm-like structures (Deveau et al. 2010; Miquel Guennoc et al. 2017). Whether such biofilm formation is specific to the interaction between the ECM fungus L. bicolor S238N and its MHB P. fluorescens BBc6 and whether it is influenced by the mycorrhizal symbiosis with trees were unknown. In this context, we investigated in depth the process of biofilm formation by P. fluorescens BBc6 on the hyphae the ECM L. bicolor and we evaluated the degree of specificity of the interaction by comparing the behaviour of various bacteria and of various ECM and saprophytic fungi. Lastly, we tested the impact of the root system and of the ectomycorrhizal symbiosis on the fate of the interaction. Our data suggest that biofilm formation on hyphae does not occur randomly but then it is regulated by several biotic factors. In addition, the detailed study of the structure of the bacterial biofilms revealed a feature that seems to be surprisingly widespread in fungal-bacterial interactions: the use of extracellular DNA filaments to build the skeleton of biofilms. MATERIALS AND METHODS Microbial strains and culture conditions All the strains used in this study are listed in Table 1. GFP-tagged versions of P. fluorescens BBc6 (Deveau et al. 2010), Burkholderia ginsengisoli E3BF7_7 and Dyella sp. E3BF9_7 were used to facilitate imaging. None of the other microbial strains used in this study constitutively expressed a fluorescent protein. Bacterial strains were maintained at −80°C in Luria-Bertani (LB) medium with 30% glycerol and were first grown on 10% tryptic soy agar (TSA)-plates for 24 h (3 g l−1 tryptic soy broth from Difco and 15 g l−1 agar) at 28°C except Lactococcus lactis MG1363 that was grown on M17-plates medium (5 g l−1 pancreate digest casein, 5 g l−1 soy peptone, 5 g l−1 beef extract, 2.5 g l−1 yeast extract, 0.5 g l−1 ascorbic acid, 0.25 g l−1 magnesium sulfate, 19 g l−1, 1% glucose and 15 g l−1 agar). Then, for each strain except L. lactis MG1363, 2– 3 individual bacterial colonies were collected from culture plates to inoculate 25 ml of liquid LB medium and incubated at 28°C and 150 rpm until late exponential growth before their use for biofilms formation. Lactococcus lactis MG1363 was inoculated in 25 ml of M17 medium. Fungal cultures were maintained on P5 medium then transferred to P20 agar plates covered with EDTA pre-treated cellophane membranes as described by Miquel Guennoc et al. (2017), except for Tuber melanosporum that require low carbon levels to grow in vitro that was always kept on P20 medium. Table 1. List and characteristics of the microbial strains. Microbial strains Taxonomic classification Gram type Ecological trait References Bacterial strains Pseudomonas fluorescens BBc6 Gamma-proteobacteria, Pseudomonadaceae − Mycorrhiza helper bacterium Duponnois and Garbaye (1991) Pseudomonas protegens Pf5 Gamma-proteobacteria, Pseudomonadaceae − Biocontrol Howell and Stipanovic (1979) Pseudomonas fluorescens SBW25 Gamma-proteobacteria, Pseudomonadaceae − Plant growth promoting and biocontrol Bailey et al. (1995) Pseudomonas fluorescens Pf29A Gamma-proteobacteria, Pseudomonadaceae − Biocontrol Chapon et al. (2002) Pseudomonas sp. GM18 Gamma-proteobacteria, Pseudomonadaceae − Mycorrhiza helper bacterium Labbé et al. (2014) Pseudomonas syringae pv. tomato DC3000 Gamma-proteobacteria, Pseudomonadaceae − Plant pathogen Cuppels (1986) Dyella sp. E3BF9_7 Gamma-proteobacteria, Rhodonobacteraceae − White rot associated bacterium Hervé et al. (2016) Collimonas fungivorans Ter331 Beta-proteobacteria, Oxalobacteraceae − Mycophagous bacterium de Boer et al. (2004) Burkholderia ginsengisoli E3BF7_7 Beta-proteobacteria, Burkholderiaceae − White rot associated bacterium Hervé et al. (2016) Sinorhizobium meliloti 1021 Alpha-proteobacteria, Rhizobiaceae − Nitrogen fixing plant symbiont Meade et al. (1982) Pedobacter sp. D3AIN17A Bacteroidetes, Sphingobacteriaceae − Black truffle associated bacterium Deveau et al. (unpublished) Paenibacillus sp. F2001-L Firmicutes, Paenibacillaceae + Endohyphal strain of L. bicolor Bertaux et al. (2003) Bacillus subtilis MB3 Firmicutes, Bacillaceae + Mycorrhiza helper bacterium Duponnois and Garbaye (1991) Bacillus sp. EJP109 Firmicutes, Bacillaceae + Mycorrhiza helper bacterium Poole et al. (2001) Lactococcus lactis MG1363 Firmicutes, Streptococcaceae + Lactic acid bacteria – dairy industry Chuzeville et al. (2015) Fungal strains Laccaria bicolor S238N Basidiomycota, Tricholomataceae N/A ECM Martin et al. (2008) Lactarius quietus Basidiomycota, Russulaceae N/A ECM INRA Nancy Hebeloma cylindrosporum Basidiomycota, Cortinariaceae N/A ECM Kohler et al. (2015) Piloderma croceum Basidiomycota, Atheliaceae N/A ECM Kohler et al. (2015) Phanerochaete chrysosporium RP78 Basidiomycota, Phanerochaeteceae N/A Saprotroph (white rot) Hervé et al. 2016 Thelephora terrestris Basidiomycota, Thelephoraceae N/A ECM INRA Nancy Tuber melanosporum Mel28 Ascomycota, Tuberaceae N/A ECM Martin et al. (2010) Penicillium funiculosum PF01 Ascomycota, Aspergillaceae N/A Saprotroph and plant pathogen INRA Nancy Aspergillus ustus AU01 Ascomycota, Aspergillaceae N/A Saprotroph INRA Nancy Microbial strains Taxonomic classification Gram type Ecological trait References Bacterial strains Pseudomonas fluorescens BBc6 Gamma-proteobacteria, Pseudomonadaceae − Mycorrhiza helper bacterium Duponnois and Garbaye (1991) Pseudomonas protegens Pf5 Gamma-proteobacteria, Pseudomonadaceae − Biocontrol Howell and Stipanovic (1979) Pseudomonas fluorescens SBW25 Gamma-proteobacteria, Pseudomonadaceae − Plant growth promoting and biocontrol Bailey et al. (1995) Pseudomonas fluorescens Pf29A Gamma-proteobacteria, Pseudomonadaceae − Biocontrol Chapon et al. (2002) Pseudomonas sp. GM18 Gamma-proteobacteria, Pseudomonadaceae − Mycorrhiza helper bacterium Labbé et al. (2014) Pseudomonas syringae pv. tomato DC3000 Gamma-proteobacteria, Pseudomonadaceae − Plant pathogen Cuppels (1986) Dyella sp. E3BF9_7 Gamma-proteobacteria, Rhodonobacteraceae − White rot associated bacterium Hervé et al. (2016) Collimonas fungivorans Ter331 Beta-proteobacteria, Oxalobacteraceae − Mycophagous bacterium de Boer et al. (2004) Burkholderia ginsengisoli E3BF7_7 Beta-proteobacteria, Burkholderiaceae − White rot associated bacterium Hervé et al. (2016) Sinorhizobium meliloti 1021 Alpha-proteobacteria, Rhizobiaceae − Nitrogen fixing plant symbiont Meade et al. (1982) Pedobacter sp. D3AIN17A Bacteroidetes, Sphingobacteriaceae − Black truffle associated bacterium Deveau et al. (unpublished) Paenibacillus sp. F2001-L Firmicutes, Paenibacillaceae + Endohyphal strain of L. bicolor Bertaux et al. (2003) Bacillus subtilis MB3 Firmicutes, Bacillaceae + Mycorrhiza helper bacterium Duponnois and Garbaye (1991) Bacillus sp. EJP109 Firmicutes, Bacillaceae + Mycorrhiza helper bacterium Poole et al. (2001) Lactococcus lactis MG1363 Firmicutes, Streptococcaceae + Lactic acid bacteria – dairy industry Chuzeville et al. (2015) Fungal strains Laccaria bicolor S238N Basidiomycota, Tricholomataceae N/A ECM Martin et al. (2008) Lactarius quietus Basidiomycota, Russulaceae N/A ECM INRA Nancy Hebeloma cylindrosporum Basidiomycota, Cortinariaceae N/A ECM Kohler et al. (2015) Piloderma croceum Basidiomycota, Atheliaceae N/A ECM Kohler et al. (2015) Phanerochaete chrysosporium RP78 Basidiomycota, Phanerochaeteceae N/A Saprotroph (white rot) Hervé et al. 2016 Thelephora terrestris Basidiomycota, Thelephoraceae N/A ECM INRA Nancy Tuber melanosporum Mel28 Ascomycota, Tuberaceae N/A ECM Martin et al. (2010) Penicillium funiculosum PF01 Ascomycota, Aspergillaceae N/A Saprotroph and plant pathogen INRA Nancy Aspergillus ustus AU01 Ascomycota, Aspergillaceae N/A Saprotroph INRA Nancy View Large Table 1. List and characteristics of the microbial strains. Microbial strains Taxonomic classification Gram type Ecological trait References Bacterial strains Pseudomonas fluorescens BBc6 Gamma-proteobacteria, Pseudomonadaceae − Mycorrhiza helper bacterium Duponnois and Garbaye (1991) Pseudomonas protegens Pf5 Gamma-proteobacteria, Pseudomonadaceae − Biocontrol Howell and Stipanovic (1979) Pseudomonas fluorescens SBW25 Gamma-proteobacteria, Pseudomonadaceae − Plant growth promoting and biocontrol Bailey et al. (1995) Pseudomonas fluorescens Pf29A Gamma-proteobacteria, Pseudomonadaceae − Biocontrol Chapon et al. (2002) Pseudomonas sp. GM18 Gamma-proteobacteria, Pseudomonadaceae − Mycorrhiza helper bacterium Labbé et al. (2014) Pseudomonas syringae pv. tomato DC3000 Gamma-proteobacteria, Pseudomonadaceae − Plant pathogen Cuppels (1986) Dyella sp. E3BF9_7 Gamma-proteobacteria, Rhodonobacteraceae − White rot associated bacterium Hervé et al. (2016) Collimonas fungivorans Ter331 Beta-proteobacteria, Oxalobacteraceae − Mycophagous bacterium de Boer et al. (2004) Burkholderia ginsengisoli E3BF7_7 Beta-proteobacteria, Burkholderiaceae − White rot associated bacterium Hervé et al. (2016) Sinorhizobium meliloti 1021 Alpha-proteobacteria, Rhizobiaceae − Nitrogen fixing plant symbiont Meade et al. (1982) Pedobacter sp. D3AIN17A Bacteroidetes, Sphingobacteriaceae − Black truffle associated bacterium Deveau et al. (unpublished) Paenibacillus sp. F2001-L Firmicutes, Paenibacillaceae + Endohyphal strain of L. bicolor Bertaux et al. (2003) Bacillus subtilis MB3 Firmicutes, Bacillaceae + Mycorrhiza helper bacterium Duponnois and Garbaye (1991) Bacillus sp. EJP109 Firmicutes, Bacillaceae + Mycorrhiza helper bacterium Poole et al. (2001) Lactococcus lactis MG1363 Firmicutes, Streptococcaceae + Lactic acid bacteria – dairy industry Chuzeville et al. (2015) Fungal strains Laccaria bicolor S238N Basidiomycota, Tricholomataceae N/A ECM Martin et al. (2008) Lactarius quietus Basidiomycota, Russulaceae N/A ECM INRA Nancy Hebeloma cylindrosporum Basidiomycota, Cortinariaceae N/A ECM Kohler et al. (2015) Piloderma croceum Basidiomycota, Atheliaceae N/A ECM Kohler et al. (2015) Phanerochaete chrysosporium RP78 Basidiomycota, Phanerochaeteceae N/A Saprotroph (white rot) Hervé et al. 2016 Thelephora terrestris Basidiomycota, Thelephoraceae N/A ECM INRA Nancy Tuber melanosporum Mel28 Ascomycota, Tuberaceae N/A ECM Martin et al. (2010) Penicillium funiculosum PF01 Ascomycota, Aspergillaceae N/A Saprotroph and plant pathogen INRA Nancy Aspergillus ustus AU01 Ascomycota, Aspergillaceae N/A Saprotroph INRA Nancy Microbial strains Taxonomic classification Gram type Ecological trait References Bacterial strains Pseudomonas fluorescens BBc6 Gamma-proteobacteria, Pseudomonadaceae − Mycorrhiza helper bacterium Duponnois and Garbaye (1991) Pseudomonas protegens Pf5 Gamma-proteobacteria, Pseudomonadaceae − Biocontrol Howell and Stipanovic (1979) Pseudomonas fluorescens SBW25 Gamma-proteobacteria, Pseudomonadaceae − Plant growth promoting and biocontrol Bailey et al. (1995) Pseudomonas fluorescens Pf29A Gamma-proteobacteria, Pseudomonadaceae − Biocontrol Chapon et al. (2002) Pseudomonas sp. GM18 Gamma-proteobacteria, Pseudomonadaceae − Mycorrhiza helper bacterium Labbé et al. (2014) Pseudomonas syringae pv. tomato DC3000 Gamma-proteobacteria, Pseudomonadaceae − Plant pathogen Cuppels (1986) Dyella sp. E3BF9_7 Gamma-proteobacteria, Rhodonobacteraceae − White rot associated bacterium Hervé et al. (2016) Collimonas fungivorans Ter331 Beta-proteobacteria, Oxalobacteraceae − Mycophagous bacterium de Boer et al. (2004) Burkholderia ginsengisoli E3BF7_7 Beta-proteobacteria, Burkholderiaceae − White rot associated bacterium Hervé et al. (2016) Sinorhizobium meliloti 1021 Alpha-proteobacteria, Rhizobiaceae − Nitrogen fixing plant symbiont Meade et al. (1982) Pedobacter sp. D3AIN17A Bacteroidetes, Sphingobacteriaceae − Black truffle associated bacterium Deveau et al. (unpublished) Paenibacillus sp. F2001-L Firmicutes, Paenibacillaceae + Endohyphal strain of L. bicolor Bertaux et al. (2003) Bacillus subtilis MB3 Firmicutes, Bacillaceae + Mycorrhiza helper bacterium Duponnois and Garbaye (1991) Bacillus sp. EJP109 Firmicutes, Bacillaceae + Mycorrhiza helper bacterium Poole et al. (2001) Lactococcus lactis MG1363 Firmicutes, Streptococcaceae + Lactic acid bacteria – dairy industry Chuzeville et al. (2015) Fungal strains Laccaria bicolor S238N Basidiomycota, Tricholomataceae N/A ECM Martin et al. (2008) Lactarius quietus Basidiomycota, Russulaceae N/A ECM INRA Nancy Hebeloma cylindrosporum Basidiomycota, Cortinariaceae N/A ECM Kohler et al. (2015) Piloderma croceum Basidiomycota, Atheliaceae N/A ECM Kohler et al. (2015) Phanerochaete chrysosporium RP78 Basidiomycota, Phanerochaeteceae N/A Saprotroph (white rot) Hervé et al. 2016 Thelephora terrestris Basidiomycota, Thelephoraceae N/A ECM INRA Nancy Tuber melanosporum Mel28 Ascomycota, Tuberaceae N/A ECM Martin et al. (2010) Penicillium funiculosum PF01 Ascomycota, Aspergillaceae N/A Saprotroph and plant pathogen INRA Nancy Aspergillus ustus AU01 Ascomycota, Aspergillaceae N/A Saprotroph INRA Nancy View Large In vitro biofilm formation on fungal hyphae and glass fibers The method described step-by-step in Miquel Guennoc et al. (2017) was used. Briefly, bacterial cultures in late exponential growth phases were spin down and washed once in potassium phosphate buffer (PPB; KH2PO4 25 g l−1, K2HPO4 2.78 g l−1, pH 5.8). Bacterial pellets were suspended in PPB and the cell density was adjusted to 109 cfu ml−1 to prepare the bacterial inoculum. Five milliliters of this bacterial suspension were added to each well of 6 well-micro plates. Then, using sterile tweezers, 1-cm diameter fungal colonies grown on P20-agar plates were added to each well of the micro-plates. The cellophane membranes on which they were growing were kept to avoid harming the fungal hyphae during the transfer. Plates were gently shaken for 1 min to allow the fungus to unstick from the cellophane sheets and cellophane sheets were removed from each well. These resulting micro-plates containing the bacterial and fungal inocula were incubated at 20°C and 60 rpm for 16 h, except if otherwise stated. To test for biofilm formation on dead fungal colonies, 1-cm diameter growing fungal colonies were dipped into either 3% paraformaldehyde solution for an hour or progressive ethanol baths for 3 min each (20%, 50%, 70%, 100%), then washed three times with PPB to remove traces of paraformaldehyde or ethanol. Dead fungal colonies were then immediately used to test for biofilm formation using same protocol as above. To test the formation of bacterial biofilm on abiotic surface that mimic fungal hyphae, fungal colonies were replaced by 10-µm diameter sterile glass fibres. For each treatment, at least two independent assays in triplicate were analyzed. Sample preparation for confocal and electron microscopy imaging Samples for imaging were prepared following the methodology described in Miquel Guennoc et al. (2017). Briefly, fungal colonies or glass fibres were transferred to a new micro-plate. Then, samples were rinsed with NaCl (17 g l−1) then with PPB to detach planktonic and electrostatically attached bacterial cells as described by Toljander et al. (2006). For confocal imaging, fungal colonies were cut in half with a razor blade, stained then mounted on slide with Fluoromount-G anti-fading (Fisher Scientific). Fungal hyphae were stained with 10 µg ml−1 Wheat Germ Agglutinin coupled to Alexa Fluor 633 (WGA-633, Thermofisher Scientific) for 15 min. All bacteria, except P. fluorescens BBc6, B. ginsengisoli E3BF7_7 and Dyella sp. E3BF9_7 that constitutively expressed GFP, were stained with 0.3 µM 4’,6-Diamidino-2-Phenylindole (DAPI, Thermofisher Scientific) for 15 min. Several dyes were used to visualize matrix components: 1X SYPRO Ruby (15 min incubation; Thermofisher Scientific), DAPI, propidium iodide (1 µg ml−1, 15 min; Thermofisher Scientific) or TO-PRO-3 (1 µM, 15 min; Thermofisher Scientific) were used to stain proteins and extracellular DNA (eDNA), respectively (Table S1A, Supporting Information). Confocal imaging was performed with a LSM780 Axio Observer Z1 laser scanning confocal microscope (LSCM, CarlZeiss), equipped with 405, 488 and 633 nm excitation lasers and T-PMT and GaAsp PMT detectors, coupled to ZEN 2.1 lite black software (CarlZeiss). For all experiments, images were captured with 10x 0.3 NA objective to obtain a complete view of one fourth of the fungal using a combination of tile scan and Z stack functions (5×5 fields over the entire depth of the fungal colony). Then, images were taken with a 40x 1.2 NA objective to obtain high resolution zoom images of representative events within the imaged captured at 10x. Data visualization was performed by 2D maximum intensity projection, using the “Z project” function from Fiji free software (Schindelin et al. 2012, http://fiji.sc/Fiji). For electron microscopy imaging, fungal colonies or glass fibres were first rinsed with NaCl (17 g l−1) then with sterile water to avoid crystal formation during dehydration step. Then, samples were dehydrated by freeze-drying and coated with 2 nm of platinum (quartz measurement) under argon plasma (2.5 10-2 mbar, 35 mA) with a High Vacuum Coater Leica EM ACE600 (Leica). Coated samples were imaged first with a scanning electron microscope (SEM) equipped with a Field Emission Gun (SIGMA-HPSEM-FEG, Zeiss) using high resolution 'in lens' detector at 1 kV of accelerating voltage. In a second step, some samples (biofilms grown on glass fibres) were placed in a second SEM (LEO 1450VP W-SEM, Zeiss) to perform EDS micro-analysis and mappings of elements (20 kV of accelerating voltage at 1 nA of sample current; Oxford-Instruments INCA MAPS software). Effect of T. melanosporum Mel28 on the growth of P. fluorescens BBc6 Tuber melanosporum Mel28 was maintained in liquid modified MS medium (MES 1 g l−1, glucose 1 g l−1, sucrose 5 g l−1, macroelements 50 ml l−1, microelements 100 ml l−1, adjusted to pH7) at 20°C. About 150 mg (fresh weight) of T. melanosporum mycelium were distributed in 6 100 ml-flasks containing 20 ml of fresh MS medium and incubated for 5 days at 20°C. Spent media from 3 cultures were recovered by filtration (0.45µm) and transferred to new sterile flasks. Three additional flasks containing 20 ml of fresh MS medium were also prepared. Then, 10 µl of P. fluorescens BBc6 bacterial suspension (OD600nm 0.7 ∼ 109 cfu ml−1) were added to each flask (3 containing living mycelium of T. melanosporum, 3 containing spent medium and 3 containing fresh MS medium). Bacterial growth in each treatment was followed by measuring the OD600nm with a spectrophotometer plate reader (Tecan Infinite Pro M200, France) over 42 h. Biofilm formation in the presence of Populus roots and ectomycorrhizae Micro-propagated hybrid poplar (Populus tremula × Populus alba; clone INRA 717-1-B4) were used to form mycorrhiza with L. bicolor S238N following the method of 'in vitro sandwich co-culture system' developed by Felten et al. (2009). A mycelium-covered cellophane membrane was placed on fungus side down on the roots of three weeks old Populus seedlings. Petri dishes were closed with Band-Aids (ensuring high gas permeability). Cultures were arranged vertically, and the lower part of the dish was covered with a small black plastic bag to prevent light from reaching the fungus and roots. The co-cultures were incubated for one month at 24°C and under 16 h-photoperiod then the development of mature ectomycorrhizae was controlled under a stereoscope (CarlZeiss). At this stage, bacterial suspensions were prepared as described above. Seedlings of Populus colonized by L. bicolor were transferred in a large Petri dish filled with sterile PPB (Fig. S1, Supporting Information). The cellophane membranes were gently detached by agitation (1 min) and removed. Mycorrhizal seedlings were transferred into a double Petri dish setting containing the bacterial suspension (Fig. S1, Supporting Information). The double Petri dish was designed to prevent contact between plant shoot and bacterial suspension. The systems were then incubated at 20°C with gentle agitation (60 rpm) for 16 h. Control treatments (Populus plants not inoculated with L. bicolor) were treated similarly. After 16 h of incubation, mycorrhizal and non-mycorrhizal seedlings were transferred in a new double Petri dish to be washed with NaCl and PPB as described above to remove planktonic and electrostatically attached bacterial cells. Each sample was then examined by confocal microscopy. To obtain transversal cross sections of ectomycorrhizae, ectomycorrhizae were included in 4% agarose and sectioned with a vibratome (Leica VT1200S) at a thickness of 30 µm. For each treatment, two independent assays in duplicate were performed. Enzymatic treatment of biofilms To investigate the role of extracellular DNA (eDNA) in biofilms formation, DNase I (30 Kunitz units ml−1 Qiagen) was added to P. fluorescens BBc6 suspensions before the incubation with glass fibers. To analyze the composition of P. fluorescens BBc6 biofilm filaments, biofilms grown for 16 h on glass fibers were treated with proteinase K (60 mAnson units ml−1 for 1 h, ThermoFisher), DNase I (30 Kunitz units ml−1 Qiagen), RNase A (100 Kunitz units ml−1, ThermoFisher) or cellulase R10 (1 mg ml−1, from Trichoderma viride, SERVA). All the enzymatic treatments were performed at room temperature for 2 h. Samples were observed with laser scanning confocal microscopy (LSCM). Positive controls were treated similarly. Proteinase K activity was verified with non-fat dried skimmed milk powder as described by Nygren et al. (2007). Cellulase activity was verified with AZCL-HE-Cellulose per the manufacturer's instructions (0.2 % w/v, Megazyme). For RNase activity, total RNA from human placenta (1 µg µl−1, Clontech) was treated and degradation was verified by electrophoretic migration of treated and non-treated RNA (negative control) in 1%–agarose gel stained with ethidium bromide. RESULTS Characterization of the formation of biofilms by P. fluorescens BBc6 on hyphae of ECM and non-ECM soil fungi We previously reported that the MHB P. fluorescens BBc6 forms biofilm structures on the hyphae of L. bicolor S238N in vitro (Miquel Guennoc et al. 2017) but the dynamic of the process and the structure of the biofilms were not characterized. Using the same in vitro setting, we performed a detailed characterization of the process by Laser Scanning Confocal Microscopy (LSCM) and Scanning Electron Microscopy (SEM). The in vitro setting relies on a liquid co-culture of the microorganisms followed by a stringent rinsing to detach planktonic and electrostatically attached bacterial cells (Miquel Guennoc et al. 2017). Time course imaging of the process indicated that the colonization of hyphae started by the adhesion of individual bacterial cells to the hyphae a few minutes after the co-inoculation of the microorganisms (Fig. 1a and d). Micro-colonies engulfing the hyphae and made of several layers of cells were observed after few hours (Fig. 1b and e). The colonies kept building up to form mature biofilms after about 20 h (Fig. 1c and f). Bacterial cells were encased in a dense matrix of extracellular polymeric substances (Fig. 2a) made of extracellular DNA (eDNA) and proteins (Fig. 2b and c). Moreover, we observed the presence of filaments stained by DAPI that connected bacterial cells together and that anchored the biofilms to the hyphae (Fig. 2a and c). Figure 1. View largeDownload slide Time course of P. fluorescens BBc6 biofilm formation on the surface of L. bicolor S238N hyphae. Confocal microscopy images showing the spatial localization of BBc6 biofilms (green) on L. bicolor S238N hyphae (red) and their development over time from early stage attachment (30 min; A,D) to colony formation (6 h; B,E) and mature biofilms (20 h; C,F). The yellow arrow points toward the external edge of the fungal colony. Bottom panels are zoom in of the areas highlighted by white rectangles in top panels. Fungal hyphae were stained with Wheat Germ Agglutin-AlexaFluor 633 and bacterial cells were GFP-tagged. Images were obtained via 2D maximum intensity projection of 3D confocal microscopy images (z = 40.5µm (a), 30µm (b), 43.5µm (c)). Magnification 40×. Figure 1. View largeDownload slide Time course of P. fluorescens BBc6 biofilm formation on the surface of L. bicolor S238N hyphae. Confocal microscopy images showing the spatial localization of BBc6 biofilms (green) on L. bicolor S238N hyphae (red) and their development over time from early stage attachment (30 min; A,D) to colony formation (6 h; B,E) and mature biofilms (20 h; C,F). The yellow arrow points toward the external edge of the fungal colony. Bottom panels are zoom in of the areas highlighted by white rectangles in top panels. Fungal hyphae were stained with Wheat Germ Agglutin-AlexaFluor 633 and bacterial cells were GFP-tagged. Images were obtained via 2D maximum intensity projection of 3D confocal microscopy images (z = 40.5µm (a), 30µm (b), 43.5µm (c)). Magnification 40×. Figure 2. View largeDownload slide Characterization of the matrix components of P. fluorescens BBc6 biofilms on the surface of L. bicolor S238N. A. Scanning electron microscopy image of 24 h old biofilm showing bacterial cells (*) and fungal hyphae (f) encased in a matrix made of aggregates and filaments. B. Confocal microscopy image showing the presence of proteins aggregates (white) and eDNA (blue) in the matrix of 16 h old P. fluorescens BBc6 (green) biofilm on L. bicolor S238N hyphae (red). Proteins and eDNA were stained with SYPRO Ruby and DAPI, respectively. Magnification 40×. C. Confocal microscopy image showing the presence of filaments stained by DAPI (yellow arrows) connecting hyphae to bacterial cells. Magnification 40×. Figure 2. View largeDownload slide Characterization of the matrix components of P. fluorescens BBc6 biofilms on the surface of L. bicolor S238N. A. Scanning electron microscopy image of 24 h old biofilm showing bacterial cells (*) and fungal hyphae (f) encased in a matrix made of aggregates and filaments. B. Confocal microscopy image showing the presence of proteins aggregates (white) and eDNA (blue) in the matrix of 16 h old P. fluorescens BBc6 (green) biofilm on L. bicolor S238N hyphae (red). Proteins and eDNA were stained with SYPRO Ruby and DAPI, respectively. Magnification 40×. C. Confocal microscopy image showing the presence of filaments stained by DAPI (yellow arrows) connecting hyphae to bacterial cells. Magnification 40×. The substitution of L. bicolor hyphae by glass fibres (similar diameter) in controls also led to biofilm formation around the glass fibres by P. fluorescens BBc6 (Fig. 3a), potentially suggesting that L. bicolor hyphae may be nothing more than a physical support used by bacteria to establish biofilms. To further test this hypothesis, biofilm formation on living and dead fungal colonies was compared. P. fluorescens BBc6 formed biofilms on both living and dead hyphae but the distribution of the biofilms greatly differed between the two treatments. Although sparse attachment was detected all over the fungal living colonies, mature biofilms mainly developed at the actively growing margin of the fungal colony (Fig. 3b). In contrast, biofilms were found all over dead fungal colonies (Fig. 3c) suggesting that specific interactions between bacterial and fungal cells occur during the formation of biofilms and that fungal hyphae are more than physical supports. Figure 3. View largeDownload slide Distribution of P. fluorescens BBc6 biofilms on abiotic and biotic surfaces. A. Confocal microscopy image showing P. fluorescens BBc6 biofilm formed over glass fibers after 22 h. Magnification 40×. B, C. Confocal microscopy images showing differential distribution P. fluorescens BBc6 biofilms formed over alive (B) and dead hyphae (C) of L. bicolor S238N. L. bicolor S238N hyphae were killed by immersing fungal colonies in 3% paraformaldehyde for 1 h followed by three repeated washes in phosphate buffer before inoculating bacteria. Images were obtained via 2D maximum intensity projection of 3D mosaic confocal microscopy images (z = 69µm (b), 32 µm (c)). Magnification 10×. Figure 3. View largeDownload slide Distribution of P. fluorescens BBc6 biofilms on abiotic and biotic surfaces. A. Confocal microscopy image showing P. fluorescens BBc6 biofilm formed over glass fibers after 22 h. Magnification 40×. B, C. Confocal microscopy images showing differential distribution P. fluorescens BBc6 biofilms formed over alive (B) and dead hyphae (C) of L. bicolor S238N. L. bicolor S238N hyphae were killed by immersing fungal colonies in 3% paraformaldehyde for 1 h followed by three repeated washes in phosphate buffer before inoculating bacteria. Images were obtained via 2D maximum intensity projection of 3D mosaic confocal microscopy images (z = 69µm (b), 32 µm (c)). Magnification 10×. We next tested whether such biofilm formation on L. bicolor hyphae was a specificity of the interaction between the MHB P. fluorescens BBc6 and the ECM fungus. We measured the ability of the bacterial strain to form biofilm on the hyphae of eight other ECM and non-ECM fungi isolated from soils (Table 1). Bacterial biofilms were observed on the hyphae of all ECM strains except the Ascomycete T. melanosporum (Fig. 4). Conversely, the bacterial strain produced biofilms on the surface of the wood decaying Basidiomycete Phanerochaete chrysosporium (Fig. 4a) but not on the hyphae of the Ascomycete saprophytes Aspergillus ustus AU01 (Fig. 4f) and Penicillium funiculosum PF01 (Fig. 4g). In these latter cases P. fluorescens BBc6 only attached to the hyphae and never built multilayer biofilms (Fig. 4f and g). This contrasts with Tuber hyphae on which no attachment at all was visible (Fig. 4h). To further test whether the absence of biofilm formation on T. melanosporum hyphae could be due to antibiosis, we measured the growth of P. fluorescens BBc6 in the presence of T. melanosporum spent media and in liquid co-cultures. Tuber melanosporum did not inhibit the growth of P. fluorescens BBc6 in both conditions (Fig S2, Supporting Information), suggesting that absence of biofilm formation on the hyphae of T. melanosporum was not due to antibiosis. Figure 4. View largeDownload slide Biofilm formation by P. fluorescens BBc6-GFP on hyphae of soil fungi. Fungal hyphae were stained with Wheat Germ Agglutin-AlexaFluor 633 (red), bacterial cells were GFP-tagged and eDNA was stained with DAPI (blue). Images were obtained via 2D maximum intensity projection of 3D confocal microscopy images. Magnification 40×. Figure 4. View largeDownload slide Biofilm formation by P. fluorescens BBc6-GFP on hyphae of soil fungi. Fungal hyphae were stained with Wheat Germ Agglutin-AlexaFluor 633 (red), bacterial cells were GFP-tagged and eDNA was stained with DAPI (blue). Images were obtained via 2D maximum intensity projection of 3D confocal microscopy images. Magnification 40×. Finally, to furthermore assess the degree of specificity of the physical interaction between the MHB and the ECM fungus, 13 additional cultivable bacteria spanning over a wide range of taxa highly represented in soil or associated with plants, and with various ecological traits (e.g. MHB, biocontrol, pathogen, Table 1) were tested for their ability to form a biofilm on L. bicolor S238N hyphae. Lactococcus lactis MG1363, a bacterial strain that is not naturally found in soils and that is a poor biofilm former (Chuzeville et al. 2015) was used as a negative control. All bacteria, except L. lactis MG1363, formed biofilms around the hyphae at the edge of L. bicolor colonies (Fig. S3, Supporting Information). The biofilms were made of several layers of cells engulfed in a matrix containing eDNA and proteins. By contrast, only individual cells of L. lactis could be punctually spotted on the surface of the hyphae of L. bicolor (Fig. S3, Supporting Information). The presence of tree roots and ectomycorrhizae modifies P. fluorescens BBc6 behavior Roots and ectomycorrhizae—the mixed organ composed of root and fungal tissues in which nutritional exchanges take place (Martin et al. 2017)—are nutrient hotspots that chemoattract complex communities of bacteria and that can provide to bacteria alternative habitats from the hyphosphere (Danhorn and Fuqua 2007; Bonfante and Anca 2009). We tested whether the presence of these organs would modify the behaviour of P. fluorescens BBc6 using Poplar as a tree model organism. Poplar seedlings were grown in vitro and used to produce ectomycorrhizae with L. bicolor S238N. The root system, together with the associated mycelium, were then incubated with P. fluorescens BBc6 bacteria in a liquid setup without nutrients for 16 h to assess biofilm formation on free roots, ectomycorrhizae, short- and long-distance extramatrical mycelium (Fig. S1, Supporting Information). Bacteria heavily colonized the surfaces of free roots (Fig. 5a), ectomycorrhizae (Fig. 5b), and short-distance extramatrical mycelium (i.e. emerging from the ECM; Fig. 5c). By contrast, no biofilm was detected on distant hyphae (Fig. 5d). We hypothesized that the absence of biofilm could be either due to physiological differences between short- and long-distance extramatrical hyphae that would not favor biofilm formation on long distance hyphae, or to the presence of the root system that would offer a more attractive habitat for the bacteria than the hyphosphere. To test these hypotheses, part of the long-distance extramatrical mycelium was sampled and transferred to a new plate containing a bacterial inoculum. After 16 h, a biofilm had formed on the surface of this 'free' long-distance extramatrical mycelium (Fig. 5e). We conclude that short- and long-distance hyphae are both susceptible to support biofilms. The presence of ectomycorrhizae would induce a change of behaviour of the bacterial cells that would colonize roots, ECM and surrounding hyphae instead of long distance hyphae. Figure 5. View largeDownload slide Differential distribution of P. fluorescens BBc6 biofilms on Poplar roots, ectomycorhizae and extramatrical mycelium after 16 h of interaction. A. Confocal image showing P. fluorescens BBc6 (green) colonization of Poplar roots (blue). B. Transversal section of L. bicolor S238N (red)–Poplar (blue) ectomycorrhizae colonized by P. fluorescens BBc6 (green). C. Confocal image of L. bicolor S238N extramatrical hyphae (blue) surrounding Poplar root (red) and colonized by P. fluorescens BBc6 (green). D. Confocal image showing L. bicolor S238N extramatrical hyphae (red) distant from root and ectomycorhizae that are not colonized by P. fluorescens BBc6 (green) in the presence of Poplar root system. E. Confocal image showing L. bicolor S238N extramatrical hyphae (red) distant from root and ectomycorhizae colonized by P. fluorescens BBc6 (green) in the absence of Poplar root system. Magnification 40×. Laccaria bicolor S238N hyphae were stained with Wheat Germ Agglutin-AlexaFluor 633 (red), Poplar root cells and eDNA were visualized with a combination of DAPI staining and autofluorescence (blue) and bacterial cells were GFP-tagged (green). All images were obtained via 2D maximum intensity projection of 3D confocal microscopy images (z = 60 µm (a), 57 µm (b), 68 µm (c), 46 µm (d), 29 µm (e)). Magnification 40×. mr: main root, rh: root hairs, exm: extramatrical mycelium. Figure 5. View largeDownload slide Differential distribution of P. fluorescens BBc6 biofilms on Poplar roots, ectomycorhizae and extramatrical mycelium after 16 h of interaction. A. Confocal image showing P. fluorescens BBc6 (green) colonization of Poplar roots (blue). B. Transversal section of L. bicolor S238N (red)–Poplar (blue) ectomycorrhizae colonized by P. fluorescens BBc6 (green). C. Confocal image of L. bicolor S238N extramatrical hyphae (blue) surrounding Poplar root (red) and colonized by P. fluorescens BBc6 (green). D. Confocal image showing L. bicolor S238N extramatrical hyphae (red) distant from root and ectomycorhizae that are not colonized by P. fluorescens BBc6 (green) in the presence of Poplar root system. E. Confocal image showing L. bicolor S238N extramatrical hyphae (red) distant from root and ectomycorhizae colonized by P. fluorescens BBc6 (green) in the absence of Poplar root system. Magnification 40×. Laccaria bicolor S238N hyphae were stained with Wheat Germ Agglutin-AlexaFluor 633 (red), Poplar root cells and eDNA were visualized with a combination of DAPI staining and autofluorescence (blue) and bacterial cells were GFP-tagged (green). All images were obtained via 2D maximum intensity projection of 3D confocal microscopy images (z = 60 µm (a), 57 µm (b), 68 µm (c), 46 µm (d), 29 µm (e)). Magnification 40×. mr: main root, rh: root hairs, exm: extramatrical mycelium. Skeleton of bacterial biofilms on fungal hyphae are made of eDNA filaments Biofilms formed by P. fluorescens BBc6 on the hyphae of L. bicolor S238N and of the other fungi analyzed were characterized by the presence of complex networks of filaments stained by DAPI (Fig. 2c; Fig. S4, Supporting Information). Filaments started to form very early during the interactions as they could be visualized by CLSM 30 min after the co-inoculation of the microorganisms (Fig. 6a). These filaments could reach a length of several hundred micrometres, and were produced, at least, by the bacteria since they were also retrieved in BBc6 biofilms formed on glass fibres (Fig. 6b). In contrast, no filament could be seen in fungal colonies of L. bicolor cultured without bacteria (Fig. 6c). Figure 6. View largeDownload slide Production of eDNA filaments by P. fluorescens BBc6 during biofilm formation. A. Confocal images showing filaments stained by DAPI (blue) after 30 min of co-incubation between P. fluorescens BBc6 and L. bicolor. Laccaria bicolor was stained with Wheat Germ Agglutin-AlexaFluor 633 (red). The three panels show three different areas of the hyphal colony on which filaments were visible. Magnification 40×. B. Confocal images showing mm long filament structures stained by DAPI DNA marker in biofilms of P. fluorescens built on glass fibers. Magnification 40×. C. Hyphae of L. bicolor grown in the absence of P. fluorescens BBc6 and imaged by SEM. D, F. Confocal images of TO-PRO-3 stained filaments in P. fluorescens BBc6 16 h old biofilm on glass fibers before (D) and after DNAse I treatment (F). White arrows point at filament positions. Magnification 10×. E. Stitched confocal images (4×4 fields) of glass fibers 16 h after the inoculation of P. fluorescens BBc6 in the presence of DNAse I. DAPI staining was used to visualize potential filaments. Magnification 10×. Result of the control experiment performed without DNAse treatment is provided in Fig. S6, Supporting Information. Figure 6. View largeDownload slide Production of eDNA filaments by P. fluorescens BBc6 during biofilm formation. A. Confocal images showing filaments stained by DAPI (blue) after 30 min of co-incubation between P. fluorescens BBc6 and L. bicolor. Laccaria bicolor was stained with Wheat Germ Agglutin-AlexaFluor 633 (red). The three panels show three different areas of the hyphal colony on which filaments were visible. Magnification 40×. B. Confocal images showing mm long filament structures stained by DAPI DNA marker in biofilms of P. fluorescens built on glass fibers. Magnification 40×. C. Hyphae of L. bicolor grown in the absence of P. fluorescens BBc6 and imaged by SEM. D, F. Confocal images of TO-PRO-3 stained filaments in P. fluorescens BBc6 16 h old biofilm on glass fibers before (D) and after DNAse I treatment (F). White arrows point at filament positions. Magnification 10×. E. Stitched confocal images (4×4 fields) of glass fibers 16 h after the inoculation of P. fluorescens BBc6 in the presence of DNAse I. DAPI staining was used to visualize potential filaments. Magnification 10×. Result of the control experiment performed without DNAse treatment is provided in Fig. S6, Supporting Information. Filaments could only be visualized when stained with DNA specific dyes (DAPI, propidium iodide and TO-PRO-3, Table S1A, Supporting Information). None of the other dyes tested (e.g. cellulose specific dyes) gave a positive result (Table S1A, Supporting Information). In accordance to a DNA composition of the filaments, phosphorous and nitrogen, two major components of DNA, were both detected at the surface of the filaments by EDS-SEM elemental mapping (Fig. S5, Supporting Information). Lastly, a DNAse treatment disrupted the filaments and dismantled the biofilms (Fig. 6d and f) while proteinase K, RNAse and cellulase had no visible effect (Table S1B, Supporting Information). The filaments were also detected when 2% glucose was added to the incubation medium (data not shown), indicating that the DNA filaments were not produced as a substitution strategy for cellulose caused by the absence of carbon sources (Serra, Richter and Hengge 2013). Finally, the addition of DNAse at the time of inoculation of bacteria fully blocked the formation of filaments and of biofilms (Fig. 6e; Fig. S6, Supporting Information), suggesting that these eDNA filaments were cornerstone structural elements for biofilm assembly. To assess whether this eDNA skeleton was specific of the interaction between the MHB P. fluorescens BBc6 and fungi, we looked for eDNA filaments in the biofilms of the 13 additional bacterial strains that formed biofilms on the hyphae L. bicolor. All the biofilms formed by these bacteria around L. bicolor hyphae contained eDNA filaments (Fig. S3, Supporting Information). Altogether our data suggest that eDNA is frequently produced during the formation of bacterial biofilms on soil hyphae. DISCUSSION In many natural environments, bacteria preferentially live in biofilms that are built on abiotic surfaces but also on living tissues such as roots (Burmølle, Kjøller, Sørensen 2012; Flemming et al. 2016). Filamentous fungi represent up to 75% of the subsurface microbial biomass with extended networks of 102– 104 m length per g of topsoil (Ritz and Young 2004) and thus could provide immense surfaces for bacteria to form biofilms. Yet, little is known on the extent on such phenomenon and the consequences for the fitness of both microorganisms. Our data indicate that the MHB P. fluorescens BBc6, but also a large range of fungal interacting bacteria with various life styles and taxonomic affiliations, have the ability to form biofilms on the hyphae of the ECM fungus L. bicolor S238N. This generic behaviour contrasts with the one of arbuscular mycorrhizal associated bacteria that differed in their ability to form biofilm on the hyphae of Rhizophagus intraradices and Glomus sp. (Toljander et al. 2006; Scheublin et al. 2010). This may reflect different levels of specificity in the physical interactions, some bacteria being able to form biofilms on a large range of fungi and surfaces while other would rely on more specific and tightly regulated interactions with the fungi to form biofilms. By contrast, P. fluorescens BBc6 could not build biofilms on all fungal strains tested (Fig. 4). The mechanisms behind the inhibition of biofilm formationby certain fungi remain to be discovered. However, our data suggest that they rely on different processes depending on the fungal species. For instance, preliminary data in T. melanosporum suggest that the fungus would actively inhibit biofilm formation because biofilm formation was observed on dead hyphae (data not shown). Since antibiosis is not involved in the inhibition of biofilm formation (Fig. S2, Supporting Information)and eDNA production is necessary for biofilm formation (Fig 6d and e), it is tempting to speculate that T. melanosporum would secrete DNAse to prevent biofilm formation on its hyphae. This hypothesis is supported by the fact that one of the predicted DNAse encoded in the genome of T. melanosporum (Martin et al. 2010) is predicted to be secreted (data not shown). Furthermore, the gene encoding for the putative secreted DNase was actively transcribed in the free-living mycelium of T. melanosporum (Tisserand et al. 2011). Further experiments will be necessary to confirm this hypothesis. Bacterial biofilm formation on hyphae can be both advantageous and detrimental for the fungi: on one hand, bacterial biofilms could protect the fungal hyphae against grazing, toxic compounds and buffer environmental variations(Kuramitsu et al. 2007; Balbontín, Vlamakis and Kolter 2014; Nazir, Tazetdinova and van Elsas 2014). On the other hand, the presence of biofilms on the growing active area of fungal colonies may limit the capacity of fungi to degrade organic matter and induce a competition for nutrients between the bacterial community and the hyphae. In this regard, the saprotrophic fungus Coprinopsis cinerea has developed complex strategies to limit the formation of biofilms on its hyphae (Essig et al. 2014; Stöckli et al. 2016). Determining the effect of biofilm formation on the fitness of ECM and non-ECM fungi would therefore be important in the future to predict the output of such bacterial–fungal interactions. In the present study, P. fluorescens BBc6 preferentially formed biofilms at the edge of the actively growing colonies while they colonized the entire fungal colonies when those were killed by fixation in paraformaldehyde (Fig. 3) or ethanol (data not shown). Formation of biofilms on the growing tips of hyphae of fungi by the bacterium Burkholderia terrae was also previously reported (Nazir, Tazetdinova, van Elsas 2014). Such differential distribution could be caused by a polarized heterogeneity of the fungal colony, either in the composition of the fungal cell wall (Latgé 2007) or in the secretion of nutrients (Webster and Weber 2007). The localization of bacterial biofilms on fungal hyphae was also strongly influenced by external biotic cues such as the presence of roots or ectomycorrhizae (Fig. 5). Both plant roots and fungal hyphae produce various exudates that chemoattract bacteria and can be used as a nutrient source (Deveau et al. 2010; Nazir et al. 2010; Rudnick, van Veen and de Boer 2015; Stopnisek et al. 2016) but bacteria differ in their abilities to use these nutrients (Frey et al. 1997). As our results suggest, in the absence of bacterial competition, roots and ectomycorrhizae may support more bacterial biofilm formation than the fungus alone. However, competition between bacteria in natural environment is likely to hinder such behaviour (Förster et al. 2016; Kastman et al. 2016) and it will be necessary to integrate the multiple interactions that occur within multispecies mixed biofilms in further studies. In addition, settings mimicking thephysicochemical properties of soils and their texture will allow shedding light onto the parameters that drive bacterial–fungal interactions in soils. The large taxonomic distribution of bacteria able to form a biofilm on L. bicolor S238N hyphae suggest that there was a low degree of specificity of the bacteria towards the fungal host, and thus that the interaction potentially relied on a common mechanism. We observed that the biofilms produced by the 14 bacterial species were structured by a network of filaments that seemed to maintain cell together and to anchor the biofilms to the surface of hyphae and of glass fibres ( Figs. 2 and 4; Fig. S3, Supporting Information). Our data strongly suggest that these skeletons would be made of eDNA. Since its first discovery in the biofilms of the aquatic bacterium Reinheimera sp. F8 (Böckelmann et al. 2006), several studies have reported the presence of a similar organization of eDNA into filaments in biofilms of various bacterial species (Jurcisek and Bakaletz 2007; Barnes et al. 2012; Gloag et al. 2013; Novotny et al. 2013; Tang et al. 2013; Liao et al. 2014; Rose et al. 2015; Tran et al. 2016) and in biofilms produced by the fungus Aspergillus fumigatus (Rajendran et al. 2013). It is likely that the networks of eDNA filaments observed during the interaction between P. fluorescens BBc6 and the different fungi had a bacterial origin. Indeed, P. fluorescens BBc6 produced similar filaments while forming biofilms on glass fibres, while no filaments could be observed onfungal colonies grown alone (Fig. 6). However, we cannot exclude, at this stage, that the interaction with P. fluorescens BBc6 induced eDNA production by the fungi. The isolation and sequencing of the eDNA filaments will be necessary in the future to clearly identify the producers of the eDNA filaments. Further analyses are also needed to determine the origin of the eDNA filaments produced during the formation of biofilms on the hyphae of L. bicolor by the other bacteria analyzed here. Yet, since eDNA production is a very widespread phenomenon relying on a large variety of mechanisms in bacteria (Ibáñez de Aldecoa, Zafra and González-Pastor 2017), we hypothesize that the formation of eDNA filaments is also prevalent among bacteria. The role of such eDNA filaments in the formation and functioning of bacterial biofilms remains elusive. Consistently with the hypothesis that eDNA would serve as a cohesive molecule that maintain bacterial cells together and anchor them to a surface (Das et al. 2010; Tang et al. 2013), the addition of DNAse at the inoculation time of the bacteria fully blocked the formation of biofilm (Fig. 6e). Although eDNA skeletons may have additional functions in bacterial biofilms (Gloag et al. 2013; Doroshenko et al. 2014), it is noteworthy that a broad range of organisms belonging to the Animal, Plant, Protist and Eubacteria Kingdoms all uses DNA for additional purposes than coding genetic information (de Buhr, von Köckritz-Blickwede, Ckritz-Blickwede 2016; Tran et al. 2016; Zhang et al. 2016). DNA, thanks to its adhesive physicochemical properties, may also have an important role for the attachment of bacterial biofilms to fungal hyphae. Overall, our work indicates that ECM fungi may often serve as a support to biofilm formation for a wide range of soil bacteria, in addition to be a source of nutrients (Ballhausen, Vandamme and de Boer 2016; Worrich et al. 2017) and a potential vector for bacterial mobility (Simon et al. 2017). Such biofilm formation on ECM fungal hyphae is likely to be modulated by numerous biotic factors including roots exudates and fungal activities. Besides the importance of eDNA based filaments for the building of these biofilms, the molecular dialog involved in the formation of biofilms on fungal hyphae and the nature of the interaction engaged between the microorganisms will need to be further investigated. SUPPLEMENTARY DATA Supplementary data are available at FEMSEC online. ACKNOWLEDGEMENTS We would like to thank Francis Martin for helpful discussions and comments on the manuscript. We thank Frédéric Guinet for helping with the preparation of in vitro mycorrhized seedlings, Cyrille Bach for providing fresh cultures of T. melanosporum for additional experiments and Béatrice Palin for technical support along the experiments. We thank Philippe Moreillon (University of Lausanne, Switzerland) and Sophie Payot-Lacroix (University of Lorraine, France) for providing the L. lactis MG1363 strain. FUNDING This work was supported by the French National Research Agency through the Laboratory of Excellence ARBRE [ANR-11-LABX-0002-01] and by the Plant-Microbe Interfaces Scientific Focus Area in the Genomic Science Program, the Office of Biological and Environmental Research in the U.S. Department of Energy Office of Science. Oak Ridge National Laboratory is managed by UT-Battelle, LLC, for the U.S. Department of Energy [DE-AC05-00OR22725]. Conflict of interest. None declared. REFERENCES Bailey MJ , Lilley AK , Thompson IP et al. Site directed chromosomal marking of a fluorescent pseudomonad isolated from the phytosphere of sugar beet; stability and potential for marker gene transfer . Mol Ecol . 1995 ; 4 : 755 – 64 . Google Scholar CrossRef Search ADS PubMed Balbontín R , Vlamakis H , Kolter R . Mutualistic interaction between Salmonella enterica and Aspergillus niger and its effects on Zea mays colonization . Microb Biotechnol . 2014 ; 7 : 589 – 600 . 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FEMS Microbiology Ecology – Oxford University Press
Published: May 17, 2018