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Optimization of conditions for in vitro development of Trichoderma viride-based biofilms as potential inoculants Sodimalla Triveni, Radha Prasanna & Anil Kumar Saxena

Folia Microbiologica Official Journal of the Institute of Microbiology, Academy of Sciences of the Czech Republic and Czechoslavak Society for Microbiology ISSN 0015-5632 Folia Microbiol DOI 10.1007/s12223-012-0154-1

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Author's personal copy Folia Microbiol DOI 10.1007/s12223-012-0154-1

Optimization of conditions for in vitro development of Trichoderma viride-based biofilms as potential inoculants Sodimalla Triveni & Radha Prasanna & Anil Kumar Saxena

Received: 3 October 2011 / Accepted: 20 April 2012 # Institute of Microbiology, Academy of Sciences of the Czech Republic, v.v.i. 2012

Abstract Biofilms represent mixed communities present in a diverse range of environments; however, their utility as inoculants is less investigated. Our investigation was aimed towards in vitro development of biofilms using fungal mycelia (Trichoderma viride) as matrices and nitrogen-fixing and P-solubilizing bacteria as partners, as a prelude to their use as biofertilizers (biofilmed biofertilizers, BBs) and biocontrol agents for different crops. The most suitable media in terms of population counts, fresh mass and dry biomass for Trichoderma and Bacillus subtilis/Pseudomonas fluorescens was found to be Pikovskaya broth±1 % CaCO3, while for Trichoderma and Azotobacter chroococcum, Jensen’s medium was most optimal. The respective media were then used for optimization of the inoculation rate of the partners in terms of sequence of addition of partners, fresh/dry mass of biofilms and population counts of partners for efficient film formation. Microscopic observations revealed significant differences in the progress of growth of biofilms and dual cultures. In the biofilms, the bacteria were observed growing intermingled within the fungal mycelia mat. Further, biofilm formation was compared under static and shaking conditions and the fresh mass of biofilms was higher in the former. Such biofilms are being further characterized under in vitro conditions, before using them as inoculants with crops.

Electronic supplementary material The online version of this article (doi:10.1007/s12223-012-0154-1) contains supplementary material, which is available to authorized users. S. Triveni : R. Prasanna : A. K. Saxena (*) Division of Microbiology, Indian Agricultural Research Institute (IARI), New Delhi 110012, India e-mail: [email protected]

The use of biofertilizers as a nutrient management strategy needs to be revitalized through innovative technologies, which can reproduce their beneficial effects consistently and in diverse agro-ecological regions. In this context, biofilmed biofertilizers can be a suitable option. Biofilms represent complex communities of multiple microbial species which remain attached to surfaces or at the interfaces (Lynch et al. 2003), and possess the capacity to maintain the metabolic activity under adverse environmental conditions, exhibiting increased survival in a competitive environment (Stewart 2002). Biofilms comprise layers of prokaryotic or eukaryotic cells, which can also play a key role in plant–microbe interactions, promote plant growth and reduce detrimental effects of various stresses under controlled conditions (Lucy et al. 2004). Yet, one has to admit that in agro-ecological systems, successful biotechnological applications of natural microbial isolates are rare. This can be attributed to the complexity of natural microbial systems and also to the lack of comprehensive understanding of the dynamic interactions (Gamalero et al. 2003). Warmink et al. (2011) have pointed out that novel microhabitats are formed in soil via movement, growth and death of soil fungi, bacteria, plant roots etc., which is a dynamic process. Such microhabitats represent key phenomena which determine the ecological success of the inoculated bacteria and bacteria–fungi interactions are known to enhance the movement and survival of bacteria in soil, especially as biofilms (Allison 2000). From the perspective of the bacterium, there are a number of ways in which biofilm formation on fungi can be beneficial. Firstly, bacterial colonization of a fungal surface may enable the bacteria to exploit the fungus as a source of nutrients (Hogan et al. 2004). Bacteria may scavenge nutrients from the fungal cell wall, consume fungisecreted products, or induce lysis of the fungal cells thereby liberating the intracellular contents for consumption by the local bacterial population. Secondly, in communities where

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bacteria and fungi are in competition for nutrients, biofilm formation on fungal cells could enhance bacterial antagonism of fungi by concentrating bacterially derived antifungal compounds. Most importantly, biofilm formation on the surface of fungal hyphae would enable bacteria to “travel” with fungi as they extend into new areas in search of nutrients and enhance the synergistic actions of bacteria and fungi needed to breakdown complex substrates in soil. Several examples illustrate that soil pseudomonads exhibit chemotaxis towards plant and fungal exudates, thereby allowing bacteria to congregate around populations of fungi in soils (Cornelis 2008; Lugtenberg et al. 2001) or use their polar flagella to anchor to mycelia surfaces (Sen et al. 1996). The use of fungal hyphae as “highways” for movement of attached microorganisms is known to provide a useful mechanism for the co-migration and improved survival and an important intervention in microbial ecology (De Boer et al. 2005; Nazir et al. 2010). However, very few published works are available on the use of agriculturally important organisms for developing biofilms. Trichoderma-based formulations represent more than 60 % of the biofungicide market; but no published information is available regarding biofilms formation and interactions with agriculturally important organisms. In this context, our investigation focussed towards optimization of conditions for in vitro development of biofilms (Trichoderma viride-based) with different agriculturally important bacteria—Azotobacter chroococcum, Bacillus subtilis, and Pseudomonas fluorescens, as a means of developing effective inoculants, which can have dual functions as biocontrol agents and biofertilizers.

Materials and methods Growth and maintenance of cultures Commercial biofertilizer strains viz A. chroococcum strain W5 and P. fluorescens P27, along with B. subtilis RP24 were obtained from the germplasm of the Division of Microbiology, Indian Agricultural Research Institute, New Delhi. T. viride (ITCC 2211) was obtained from the Indian Type Culture Collection (ITCC), Division of Plant Pathology, IARI, New Delhi, India. The growth media used were Jensen’s medium for Azotobacter, King’s B medium for Pseudomonas, Nutrient Broth for B. subtilis and Potato Dextrose Agar for T. viride. The flasks were incubated in shaking incubator at 30°C, except for T. viride, which was maintained as static culture at 30°C. Optimization of medium for biofilm partners A. chroococcum, B. subtilis, P. fluorescens, and T. viride were grown in five different of media—Nutrient Broth, Jensen’s, Kings B, Pikovskaya and Potato dextrose media. Erlenmeyer flasks (100 mL) containing 50 mL broth were autoclaved at 103 kPa and 121°C, and then, after cooling, inoculated with 1 mL culture in triplicate. The flasks were incubated in

shaking incubator at 30°C and 120 rpm for 48 h. The flasks containing Potato dextrose broth were incubated at 30°C under static conditions for 1 week. After incubation, the bacterial growth was measured at 640 nm in spectrophotometer. The growth of T. viride was measured by taking fresh mass of the mycelial material. Optimization of protocol for preparation of biofilms The two biofilms of B. subtilis+T. viride and P. fluorescens+T. viride were prepared in Pikovskaya medium, while Jensen’s medium was used for Trichoderma and A. chroococcum biofilm. The inocula used for the preparation of different biofilms were 1-week-old culture of fungi and 48 h-old bacterial cultures. The method of preparation of different biofilms was also optimized in terms of sequence of addition of partners. For all biofilms, two sets of preliminary experiments were undertaken. The first experiment involved inoculation of bacterial cultures first, followed by growth at 30°C for 48 h, and inoculation with T. viride. The second experiment involved inoculation with Trichoderma and incubated at static conditions at 30°C for 2 days and then the bacterial cultures were added. For the Bacillus and Pseudomonas based biofilms, growth of bacteria for 48 h, followed by T. viride addition was found to be optimal, as recorded through visual and microscopic observations. In Azotobacter+Trichoderma biofilm preparation, inoculation with Trichoderma, followed by Azotobacter was observed to be most suitable for biofilm development. Further experiments were undertaken based on these observations. Optimization of inoculum rate Different volumes of liquid cultures (0.5, 1.0, 2.0, 3.0 mL with known cell concentrations, in CFU/mL) of each partner—A. chroococcum, B. subtilis, P. fluorescens, and T. viride were used to prepare the biofilms. The population counts were done using serial dilution-plate count method, after washing the biofilms repeatedly with sterile water to remove adhering bacteria. After 16 days incubation at 30°C, the biofilms were collected from the flasks and washed with sterile water two to three times to remove non adherent cells from biofilm, then centrifuged at 2,000 rpm for 10 min, homogenized by vortex with glass beads to provide a uniform suspension which was used for population counts. The fresh mass and oven-dried biofilm (for 24 h at 70°C) were measured. Standardization of growth conditions for biofilm development The flasks were inoculated with two partners for biofilm preparation and the flasks were incubated under static and shaking (with speed of 50 rpm) at 30°C for 16 days. Fresh mass and dry mass of biofilms were recorded. Microscopic observations The progress of biofilm development was recorded at 2 day intervals by visual and microscopic observations (at ×40 magnification) using a Zeiss Model Axio

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Scope A1 microscope. After harvesting, the biofilm was washed with sterile water to remove the non adherent cells on biofilm. A loopful of biofilm was taken and spread on the slide and air dried. Two drops of the Lactophenol cotton blue stain was added and after 1 min, washed and two drops of safranine added. After 1 min, the slide was washed and covered with cover slip and observed under a microscope using ×10, ×40, and oil immersion objectives. The growth of biofilm on solid media was compared with dual culture inoculation by visual and microscopic observations using Wild Stereozoom microscope.

Results Optimization of media for biofilm preparation After 48 h, the growth of A. chroococcum was highest in Jensen’s medium (Fig. 1). The growth of P. fluorescens was at par in Potato dextrose broth, Pikovskaya and King’s B broth, followed by Nutrient broth and Jensen’s medium. During the growth period of B. subtilis+T. viride biofilm in the broth, a pH change was recorded, which led to cessation of growth, hence addition of 1 % CaCO3 to Pikovskaya broth was done to aid in biofilm development (data not given). For developing biofilms of Trichoderma–Azotobacter sp., although the growth of

NA

Jensens

T. viride was highest in Pikovskaya and King’s B broth and much lower in Jensen’s broth, the addition of 1 % yeast extract to Jensen’s medium stimulated the formation of biofilms. Standardization of inocula addition A progressive increase in population counts was recorded with increase in volume of inoculum, with highest values recorded with 3.0 mL of both partners (Table 1; Fig. 2a–c); although in terms of mass (fresh and dry), a differential effect was recorded. In the case of A. chroococcum–T. viride biofilm, fresh and dry mass of biofilm was maximum (3.170, 0.262 g) with the inoculation of 2.0 mL volume of each partner (Table 1). For the B. subtilis–T. viride biofilm, a statistically significant enhancement in terms of population counts, fresh and dry mass of biofilm was observed with 3.0 mL of liquid culture of each partner in biofilm after incubation period of 16 days (Fig. 2b; Table 1). The P. fluorescens–T. viride biofilms recorded highest fresh and dry mass with 1.0 mL inocula levels. Growth conditions for biofilm development The static cultures of Trichoderma–Pseudomonas generated almost twofold-higher mass as compared to shaker cultures (Fig. 3). Visual observations revealed that the biofilm formation under shaking conditions took longer duration and were less robust, as compared to those grown under static conditions.

PDA

Piko

KB

3.50

a

Fresh weight (g) / Absorbance

3.00

2.50

2.00

a

a

a

1.50

1.00

0.50

0.00

A.chroococcum

B.subtilis

Fig. 1 Optimization of media for growth of biofilm partners (NA Nutrient Broth; Jensens Jensen’s medium; PDA Potato Dextrose Agar medium; Piko Pikovskaya medium; KB King’s B medium);

P.fluorescens

T.viride

observations based on absorbance (A640 nm) values for bacterial cultures and fresh mass (g) of Trichoderma viride were taken

Author's personal copy Folia Microbiol Table 1 Optimization of volume of inocula of bacteria and fungi for biofilm preparation Treatments

Volume of partners (mL)

P. fluorescens and T. viride biofilm (g)*

A. chroococcum and T. viride biofilm (g)

B. subtilis and T. viride biofilm (g)

Bacteria

Fungi

Fresh mass

Dry mass

Fresh mass

Dry mass

Fresh mass

Dry mass

0.5

0.5

2.18b

1.0 2.0 3.0

a

3.01 2.05c 1.38d 11.26

0.05c 0.10a 0.07b 0.07b 19.29

2.584c 2.952b 3.170a 2.310d 8.24

0.196d 0.220b 0.262a 0.218c 8.91

2.12d 2.80c 3.01b 4.19a 4.80

0.23c 0.27b 0.29b 0.38a 15.76

0.416 0.583

0.010 0.023

0.454 0.304

0.012 0.026

0.083 0.062

0.032 0.175

T1 T2 T3 T4 CV (%)

1.0 2.0 3.0

SEM ± CD (0.05) *

From 50 mL medium

The letters (based on Duncan’s multiple range test) in a column denote values which are significantly different at P≤0.05, with a being the highest

Visual and microscopic observations Visual observations of growth of biofilms and co-inoculated cultures on the

Population(cfu/ml)

300

A.chroococcum

a

T.viride

250 a

200 b

150 c

100 50

b

c

d d

0 T1

T2 T3 Volume of partners (ml) B.subtilis

T4

T.viride

800

b

Population (cfu/ml)

700 600

c

500 400

a

d

300

b

200 100

c

d

0 T1

T2

T3

optimized solid media revealed that the pattern of growth of biofilm was very different from that of co-inoculated/dual cultures (Fig. 4a–c). Visible growth was delayed in the dual culture and bacterial growth was recorded first after 24 h, followed by the growth of fungi after 48 h. Both the bacteria and fungi grew individually as distinct patches. In the case of biofilms, the two organisms started growing after 12 h as partners, with the bacteria seen growing within the fungal mycelium (Fig. 4d–f). The monitoring of biofilm development during the co-culturing of the two partners in the medium at 30°C revealed that the bacteria readily attached to the fungal filaments and subsequently, the attached cells formed a dense film over the fungal mycelia. Microscopic observations recorded good attachment under the optimized ratio of inocula. It was observed the cells were free floating for 1 week after inoculation, and then the attachment started from the 7th day onwards and good attachment observed on 13th and 14th day which continued to progress up to 16th day. Thereafter, the bacterial cells started to detach from the film (Fig. 5).

T4

Volume of partners (ml) 800

P.fluorescens

400

d

a

300

100

b c

d

Shaking

5.0

c

500

200

Static

6.0

b

600

Fresh weight(g)

Population (cfu/ml)

a

T.viride

700

4.0 3.0 2.0 1.0

0

T1

T2 T3 Volume of partners (ml)

T4

0.0 AzTv

BsTv

PsTv

Incubation conditions

Fig. 2 Enumeration of bacterial load in the biofilm as influenced by inoculum volume of partners. a Ps–Tv (T. viride–Pseudomonas fluorescens) biofilm; b Bs–Tv (T. viride–Bacillus subtilis) biofilm; c Az–Tv (T. viride–Azotobacter chroococcum) biofilm

Fig. 3 Standardization of growth conditions (static vs shaking with speed of 50 rpm) for biofilm development. B. subtilis+T. viride (Bs–Tv); P. fluorescens+T. viride (Ps–Tv); A. chroococcum+T. viride (Az–Tv)

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Fig. 4 Comparative evaluation of growth of dual cultures and biofilms in solid media, without (a, c, d, f) with zoom (b, e); a Trichoderma viride+ Pseudomonas fluorescens dual culture, initial stage; b Trichoderma viride+Pseudomonas fluorescens dual culture, initial stage; c Trichoderma

Fig. 5 Microphotographs of biofilms showing attachment of bacteria to T. viride mycelia; a only Trichoderma viride; b Trichoderma viride–Pseudomonas fluorescens biofilm; c Trichoderma viride– Azotobacter chroococcum biofilm; d Trichoderma viride–Bacillus subtilis biofilm

viride+Pseudomonas fluorescens dual culture mature stage; d Pseudomonas fluorescens–Trichoderma viride biofilm initial stage; e Pseudomonas fluorescens–Trichoderma viride biofilm initial stage; f Pseudomonas fluorescens–Trichoderma viride biofilm mature stage

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Discussion Microbial biofilms are communities of microorganisms adhering to abiotic/biotic surfaces and embedded in an organic matrix of biological origin which provides structure and stability to the community (Parsek and Fuqua 2004; Webb et al. 2003; Wargo and Hogan 2006). Such biofilms are highly structured and comprise microbial cells (algal, fungal, bacterial and/or other microbial) and an extracellular biopolymer these cells produce, known as EPS, which provides structure and protection to the community (Branda et al. 2005; Kolter and Greenberg 2006). Stevens et al. (2008) observed that Trichoderma biofilms exhibited promise in the biological control of soft rot of sweet potato, caused by Rhizopus stolonifer. Fungal biofilms are known to play a critical role in the nutrient cycling processes in soil. Recently, it has been shown that certain fungi in soil may be involved in a novel ecological phenomenon-migration helper effect, especially with bacteria (Warmink et al. 2011). Warmink et al. (2011) showed through culture independent approaches, using DGGE patterns, that the saprotrophic fungus Lyophyllum sp. stimulates the bacterial adherence and movement in soil. Bacterial colonization of a fungal surface may be a first step in more complex bacterial– fungal endosymbiont interaction such as those that are critical in the root rhizosphere (Dorr et al. 1998). Trichoderma spp. represent the most widely employed biocontrol agents, besides their role as plant growth promoters and potential as valuable sources of secondary metabolites/ enzymes which find use in pharmaceuticals, industry and agriculture (Harman 2006). Bahata (2008) evaluated the biofilm forming potential of several fungi, including T. viride and observed that light and time significantly influenced biofilm formation. Guetsky et al. (2002) have shown that combinations of biocontrol or plant growth promoting agents provide an additive effect in their mode of action, illustrating the need for developing consortia or mixed inoculum approach. Our study focused on supplementing the benefits of Trichoderma with different agriculturally important bacteria, used as nitrogen fixers, PGPs, P-solubilizers or biocontrol agents. The biofilms developed in our study differ from purely bacterial or fungal biofilms, since the fungus act as the biotic surface to which the bacteria adhere. Similar biofilms, using Bradyrhizobium–Penicillium combination have been observed to exhibit better growth, nitrogen-fixing ability and colonization abilities than their monocultures (Jayasinghearachchi and Seneviratne 2004; Seneviratne and Jayasinghearachchi 2003). However, our approach using T. viride as the matrix and agriculturally important bacteria, especially for potential use as biofertilizers and biocontrol agents is a novel one. Visual observations and observations of growth of dual cultures and biofilms using Stereozoom microscope revealed distinct differences in pattern of growth of biofilms and dual cultures on solid media. Biofilms grew as partners

with bacteria intermingled with the Trichoderma mycelia and visible growth was recorded earlier, while dual cultures grew more slowly, with both the cultures growing separately. Microscopic observations revealed that the attachment process was initiated from 7 days onwards and detachment of cells from biofilms was recorded after 16 days of coculturing. The growth of biofilms on solid media clearly revealed the concurrent growth and interactions among the partners vis a vis growth of dual cultures. Lower volume (0.5 and/or 1.0 mL) led to the formation of thin film while more than 3.0 mL inoculum recorded more free-floating spores of fungi. Exopolysaccharide production is known to contribute to surface-colonizing biofilms and play an important role in the colonization of bacterial biofilms by non-filamentous fungi (e.g., Candida albicans) and vice-versa (Lynch and Robertson 2008; Palleroni 2010; Toljander et al. 2006; Wargo and Hogan 2006). Microscopic observations confirmed our hypothesis that P. fluorescens, which produces biofilms may be an early starter as it started to aggregate early and form attachment when compared to the other two biofilms. Microscopic studies revealed that the bacterial cells also attached to the hyphae of T. viride, and as a mutual partnership, the bacterial cells showed good colonization of fungal spores. The need for determining the optimal sequence of partners was illustrated in our study, as Azotobacter behaved differently from the other two bacteria used in preferring addition into 48-h-old Trichoderma biofilm. The quality of the other two biofilms was found to be better when Trichoderma was inoculated into 48-h-old bacterial cultures. Comparative performance of the biofilm partners in different media revealed that Pikovskaya broth significantly enhanced the growth of all the three organisms—Bacillus, Pseudomonas, and Trichoderma. The addition of 1 % calcium carbonate was found help to maintain the pH, during the biofilm development. It can be surmised that the constituents present in the Pikovskaya broth supported the growth of these three organisms. No significant differences were recorded among the media tested in terms of growth of A. chroococcum, but higher values were recorded when it was grown in Jensen’s broth. Addition of 1 % yeast extract was found to enhance the growth of the T. viride, facilitating the growth of both the partners in the A. chroococcum+T. viride biofilm. Trichoderma is known to exhibit the ability to utilize a variety of carbon and inorganic/organic nitrogen compounds as a sole source of carbon and nitrogen. The wide occurrence of T. viride attests to its ability to grow in a wide range of environmental conditions, but exhibit ecological preferences. Yeast extract was found to be the most promising combination of carbon and nitrogen for growth stimulation and cellulose production by T. viride (Gautam et al. 2010; Rossi-Rodrigues et al. 2009). Jayaswal et al. (2003) observed growth and sporulation in a broad range of physiological and environmental factors, which revealed its versatility as a biocontrol agent.

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The present study revealed that the selected bacteria and fungi were compatible, showing a progressive increase in population of both the partners, with the microscopic observations revealing parallel growth patterns of both organisms, as a biofilm. Inoculated flasks incubated under static conditions significantly showed good biofilm population when compared to the static conditions and shaking with speed of 50 rpm, especially for Trichoderma–Pseudomonas, in which the static cultures recorded twofold-higher biomass as compared to shaker cultures. Microscopic observations also revealed more number of cells attached to the mycelia under static conditions. One of the major benefits of this environment is the increased resistance to detergents and antibiotics, as the dense extracellular matrix and the outer layer of cells protect the interior of the community, besides abiotic stress (Stewart and Costerton 2001). This is supported by most of the studies on biofilms, in which the static conditions allowed better development of biofilms, as it permitted gradual attachment and colonization and the common presence of biofilms on rocks and pebbles at the bottom of most streams or rivers and often observed on the surface of stagnant pools of water (Allison 2000; Lynch et al. 2003). Future research is needed to analyze the characteristics of such biofilms in depth, so as to stimulate the formation of such biofilms and their deployment in agriculture and industry. Acknowledgments The authors are thankful to the Post Graduate School and Director, Indian Agricultural Research Institute (New Delhi, India) for providing for fellowship towards PhD program to the first author. We are thankful to the Division of Microbiology (IARI, New Delhi) for providing necessary facilities for undertaking this study.

References Allison DG (2000) Community structure and co-operation in biofilms. Cambridge University Press, Cambridge, UK Bahata YT (2008) The biofilm forming potential of potential of moulds commonly isolated from Norwegian drinking water. Thesis, Norwegian School of Veterinary Sciences, Oslo Branda SS, Vik S, Friedman L, Kolter R (2005) Biofilms: the matrix revisited. Trends Microbiol 13:20–26 Cornelis P (2008) Pseudomonas genomics and molecular biology. Caister Academic Press De Boer W, Folman LB, Summerbell RC, Boddy L (2005) Living in a fungal world: impact of fungi on soil bacterial niche development. FEMS Microbiol Rev 29:795–811 Dorr J, Hurek T, Reinhold-Hurek B (1998) Type IV pili are involved in plant–microbe and fungus–microbe interactions. Mol Microbiol 30:7–17 Gamalero E, Lingua G, Berta G, Lemanceau P (2003) Methods for studying root colonization by introduced beneficial bacteria. Agronomie 23:407–418 Gautam SP, Bundela PS, Pandey AK, Jamaluddin AMK, Sarsaiya S (2010) Optimization of the medium for the production of cellulase by the Trichoderma viride using submerged fermentation. Intl J Environ Sci 1:656–665

Guetsky R, Stienberg D, Elad Y, Fischer E, Dinoor A (2002) Improving biological control by combining biocontrol agents each with several mechanisms of disease suppression. Phytopathology 92:976– 985 Harman GE (2006) Overview of mechanisms and uses of Trichoderma spp. Phytopathology 96:190–194 Hogan DA, Vik A, Kolter R (2004) A Pseudomonas aeruginosa quorum-sensing molecule influences Candida albicans morphology. Mol Microbiol 54:1212–1223 Jayasinghearachchi HS, Seneviratne G (2004) A bradyrhizobialPenicillium spp. biofilm with nitrogenase activity improves N2-fixing symbiosis of soybean. Biol Fertil Soils 40:432–434 Jayaswal RK, Singh R, Lee YS (2003) Influence of physiological and environmental factors on growth and sporulation of an antagonistic strain of Trichoderma viride RSR 7. Mycobiology 31:36–41 Kolter R, Greenberg EP (2006) Microbial sciences: the superficial life of microbes. Nature 441:300–302 Lucy M, Reed E, Glick BR (2004) Applications of free living plant growth-promoting rhizobacteria. Antonie Van Leeuwenhoek 86:1–25 Lugtenberg BJ, Dekkers L, Bloemberg GV (2001) Molecular determinants of rhizosphere colonization by Pseudomonas. Annu Rev Phytopathol 39:461–490 Lynch AS, Robertson GT (2008) Bacterial and fungal biofilm infections. Annu Rev Med 59:415–428 Lynch, JF Lappin S, Hilary M, Costerton, JW (2003) Microbial biofilms. Cambridge University Press, Cambridge, UK Nazir R, Warmink JA, Boersma FGH, van Elsas JD (2010) Mechanisms that promote bacterial fitness in fungal-affected soil microhabitats. FEMS Microbiol Ecol 71:169–185 Palleroni NJ (2010) The Pseudomonas story. Environ Microbiol 12 (6):1377–1383 Parsek MR, Fuqua C (2004) Biofilms 2003: Emerging themes and challenges in the study of surface-associated microbial life. J Bacteriol 186:4427–4440 Rossi-Rodrigues BC, Bruchetto-Bragga MR, Tauk-Tornisielo SM, Carmona EC, Arruda VM, Netto JS (2009) Comparative growth of Trichoderma strains in different nutritional sources, using bioscreen automated system. Braz J Microbiol 40:404–410 Sen R, Nurmiaho-Lassila EL, Haahtela K, Korhonen T (1996) Specificity and mode of primary attachment of Pseudomonas fluorescens strains to the cell walls of ectomycorrhizal fungi. In: Azcon-Aguilar C, Barea JM (eds) Mycorrhizas in integrated systems from genes to plant development, 4th European symposium on Mycorrhizae. European Commission, Granada, pp 661–664 Seneviratne G, Jayasinghearachchi HS (2003) Mycelial colonization by bradyrhizobia and azorhizobia. J Biosci 28:243–247 Stevens A, Fyffe A, Khan A, Wilson VA, Williams J, Winsniewski M (2008) Culturable bacterial and Trichoderma biofilms isolated from sweet potato as it relates to biological control of rhizopus soft rot. J Plant Pathol 90:507–510 Stewart PS (2002) Mechanisms of antibiotic resistance in bacterial biofilms. Int J Med Microbial 292:107–113 Stewart PS, Costerton, JW (2001) Antibiotic resistance of bacteria in biofilms. Lancet 358:135–138 Toljander JF, Artursson V, Paul LR, Janson JK, Finlay RD (2006) Attachment of different soil bacteria to arbuscular mycorrhizal fungal extraradical hyphae is determined by hyphal vitality and fungal species. FEMS Microbiol Lett 254:34–40 Wargo MJ, Hogan DA (2006) Fungal–bacterial interactions: a mixed bag of mingling microbes. Curr Opin Microbiol 9:359–364 Warmink JA, Nazir R, Corten B, van Elsas JD (2011) Hitchhikers on the fungal highway: the helper effect for bacterial migration via fungal hyphae. Soil Biol Biochem 43:760–765 Webb JS, Thompson LS, James S, Charlton T, Tolker-Nielsen T, Koch B, Givskov M, Kjelleberg S (2003) Cell death in Pseudomonas aeruginosa biofilm development. J Bacteriol 185:4585–4592