Ectomycorrhizal symbiosis affects functional ... - Wiley Online Library

Research

Ectomycorrhizal symbiosis affects functional diversity of rhizosphere fluorescent pseudomonads Blackwell Publishing, Ltd.

Pascale Frey-Klett1, Michaël Chavatte1, Marie-Lise Clausse1, Sébastien Courrier1, Christine Le Roux2, Jos Raaijmakers3, Maria Giovanna Martinotti4, Jean-Claude Pierrat5 and Jean Garbaye1 1

UMR INRA–UHP “Interactions Arbres /Micro-organismes”, 54280 Champenoux, France; 2Laboratoire des Symbioses Tropicales et Méditerranéennes,

TA10/J, Campus international de Baillarguet, 34398 Montpellier cedex 5, France; 3Laboratory of Phytopathology, Wageningen University, the Netherlands; 4

DiSCAFF, Faculty of Pharmacy, University of East Piedmont ‘Amedeo Avogadro’, Via Bivio 6 Novara, Italy; 5INRA-ENGREF, Equipe Dynamique des

Systèmes Forestiers, 54042 Nancy, France

Summary Author for correspondence: Pascale Frey-Klett Tel. 33-(0)3-83-39-41–49 Fax: 33-(0)3-83-39-40–69 Email: [email protected] Received: 7 May 2004 Accepted: 9 July 2004

• Here we characterized the effect of the ectomycorrhizal symbiosis on the genotypic and functional diversity of soil Pseudomonas fluorescens populations and analysed its possible consequences in terms of plant nutrition, development and health. • Sixty strains of P. fluorescens were isolated from the bulk soil of a forest nursery, the ectomycorrhizosphere and the ectomycorrhizas of the Douglas fir (Pseudostuga menziesii) seedlings-Laccaria bicolor S238N. They were characterized in vitro with the following criteria: ARDRA, phosphate solubilization, siderophore, HCN and AIA production, genes of N2-fixation and antibiotic synthesis, in vitro confrontation with a range of phytopathogenic and ectomycorrhizal fungi, effect on the Douglas fir–L. bicolor symbiosis. • For most of these criteria, we demonstrated that the ectomycorrhizosphere significantly structures the P. fluorescens populations and selects strains potentially beneficial to the symbiosis and to the plant. • This prompts us to propose the ectomycorrhizal symbiosis as a true microbial complex where multitrophic interactions take place. Moreover it underlines the fact that this symbiosis has an indirect positive effect on plant growth, via its selective pressure on bacterial communities, in addition to its known direct positive effect. Key words: bacteria, ectomycorrhizosphere effect, functional diversity, Laccaria bicolor S238N, Pseudomonas fluorescens, Pseudotsuga menziesii. New Phytologist (2005) 165: 317–328 © New Phytologist (2004) doi: 10.1111/j.1469-8137.2004.01212.x

Introduction Due to the ubiquity of mycorrhizal symbioses in terrestrial ecosystems, most of the actively absorbing rootlets are connected with the surrounding soil through an interface called the mycorrhizosphere ( Johansson et al., 2004). In the case of the ectomycorrhizal symbiosis, which dominates temperate and boreal forest ecosystems, the fungal mantle even creates a physical barrier between the soil and the rootlets. The physical, biochemical and microbiological features of the rhizosphere and mycorrhizosphere are very different. From a

www.newphytologist.org

physical point of view, the emanating fungal mycelium improves soil aggregation and thus modifies soil structure in comparison with rhizosphere soil (Schreiner & Bethlenfalvay, 1995). In addition, because the ectomycorrhizal symbiosis modifies root morphology by promoting root branching (Linderman, 1988), the volume of the mycorrhizosphere soil is larger than the rhizosphere one. From a biochemical point of view, root exudation in the mycorrhizosphere is quantitatively and qualitatively different from that in the rhizosphere because mycorrhizal fungi use some of the root exudates and modify the root metabolic functions (Rambelli, 1973; Leyval

317

318 Research

& Berthelin, 1993; Rygiewicz & Andersen, 1994). Furthermore, mycorrhizal fungi associated with plant roots can produce antibiotics (Olsson et al., 1996). These differences between rhizosphere and mycorrhizosphere explain the socalled ‘mycorrhizosphere effect’ defined by Linderman (1988), which qualifies modifications of the microbial equilibrium induced by the mycorrhizal symbiosis. The mycorrhizosphere effect concerns a number of different organisms such as protozoa ( Jentschke et al., 1995; Wamberg et al., 2003), microarthropods (Cromack et al., 1988), microfungi (Neal et al., 1964), bacteria (Ames et al., 1984; Christensen & Jakobsen, 1993; Olsson et al., 1996; Timonen et al., 1998; Heinonsalo et al., 2000, Wamberg et al., 2003) and more particularly pseudomonads, which populations are quantitatively and qualitatively regulated by the symbiosis (Meyer & Linderman, 1986; Waschkies et al., 1994; Andrade et al., 1997). From a population structure point of view, Frey et al. (1997) demonstrated that the symbiosis between Douglas fir (Pseudostuga menziesii) and the ectomycorrhizal fungus Laccaria bicolor S238N very significantly modifies the metabolic diversity of the soil fluorescent pseudomonads. For instance, strains of Pseudomonas fluorescens isolated from Douglas fir–L. bicolor mycorrhizas differ from those of the bulk soil by their preferential utilization of carbohydrates, particularly trehalose. Since trehalose is the most abundant carbohydrate accumulated in the mycelium of L. bicolor S238N in vitro and since it is rarely detected in plants (Eastmond et al., 2002), Frey et al. (1997) suggested that L. bicolor exerts a trehalose-mediated selection on the soil fluorescent pseudomonads. Up to now, it is unclear whether plants can selectively favour microbial communities most beneficial to themselves (Denison et al., 2003). In this context, our aim was to further characterize the genotypic and functional diversity of the P. fluorescens strains associated with the Douglas-fir-L. bicolor symbiosis, in comparison with the bulk soil, in order to reveal functional bacterial traits potentially involved in plant nutrition, development and health, which might be selected by the ectomycorrhizal symbiosis. For this purpose, we selected 60 strains of P. fluorescens among 300 strains isolated from bulk soil, mycorrhizosphere and mycorrhizas of Douglas fir–L. bicolor (Frey et al., 1997), in a forest nursery. We genotypically characterized them by amplified 16S rDNA restriction analysis (ARDRA). We also determined functional activity potentials of these bacterial isolates by a series of in vitro assays related to the following important functions in the mycorrhizosphere: mineral nutrition (phosphate solubilization, siderophore production, N2-fixation), development (AIA synthesis), health (HCN production, antibiotic synthesis, in vitro confrontation with different phytopathogenic fungal strains from nursery soil) and effect on the ectomycorrhizal symbiosis (in vitro confrontation with different ectomycorrhizal fungal strains, effects on the Douglas fir–L. bicolor symbiosis).

New Phytologist (2005) 165: 317–328

Materials and Methods Collection of bacterial strains Three hundred strains of fluorescent pseudomonads were isolated from a Douglas fir (Pseudotsuga menziesii ) experimental nursery in north-eastern France in winter, in the following three zones: first mycorrhizosphere, that is soil adherent to the root system of Douglas fir seedlings presenting a high percentage (71%) of short roots mycorrhizal with L. bicolor, second L. bicolor mycorrhizas collected from these root systems, and third bulk soil, that is soil from unplanted plots adjacent to planted plots. As described in Frey et al. (1997), the experimental design consisted of four independent couples of 1-m2 planted and unplanted plots, 6 m distant from each other in a nursery bench. For bacterial isolation, one sample (10 g) of bulk soil was taken in each of the four unplanted plots and four independent suspensions were prepared by vigorously shaking 1 g of soil in 9 ml of sterile distilled water for 60 s. Similarly, one plant was sampled in each of the four planted plots and four independent mycorrhizosphere suspensions were prepared by vigorously shaking the entire root system with adhering soil in 100 ml of sterile distilled water for 60 s. For each root system, 10 mycorrhizas typical of L. bicolor were collected and washed together 10 times in 1 ml of sterile distilled water, then crushed with a sterile pestle in a microtube containing 1 ml of sterile distilled water. The isolation procedure of the bacterial strains was described in Frey et al. (1997). The isolates were characterized with the BIOLOG® method then classified into four homogeneous metabolic groups according to the KMACL4 software, a classification method for building non-overlapping clusters of bacterial isolates (Lelu, 1994; Frey et al., 1997). BIOLOG revealed that 90% of the isolates belonged to the Pseudomonas fluorescens species with a majority (81.5%) of the P. fluorescens biovar I and V (Frey et al., 1997). The present study focused on these two main biovars, which corresponded to 220 isolates. Sixty isolates of P. fluorescens biovar I and biovar V (29 and 31 isolates, respectively) were subsampled at random among the four metabolic groups revealed by the KMACL4 analysis, in order to be representative of the whole collection of the 300 isolates. In the subcollection, the proportion of the selected biovar I and V isolates in each sampling zone (bulk soil, mycorrhizosphere, mycorrhizas) was chosen to be as close as possible to the one in the whole collection in order to form a subsample representative of the actual population (Table 1). Genotypic fingerprinting of the isolates by ARDRA Bacterial isolates were grown in 3 ml of shaken (300 rpm) liquid King B medium (King et al., 1954), at 25°C for 24 h. They were harvested by centrifugation (12 000 × g, 6 min) and the pellet was washed twice in 1 ml of sterile deionized water (18 °C, Millipore-Q reagent water). The resulting cell

www.newphytologist.org © New Phytologist (2005)

Research

Table 1 Sampling design of the 60 Pseudomonas fluorescens isolates from the subcollection in relation to the zones of origin of the isolates and their belonging to the two biovars I et V Number of isolate (percent per each biovar type) Origin of the isolate

Biovar

In subsample

In total sample

Bulk soil

I V I V I V I V

1 (3%) 22 (71%) 19 (65%) 8 (26%) 9 (31%) 1 (3%) 29 31

1 (1%) 91 (71%) 62 (67%) 36 (28%) 29 (31%) 1 (1%) 92 128

Mycorhizosphere soil Mycorrhiza Total

suspensions, with A600nm 2, were kept at −20°C. Before PCR amplification, 100 µl of these cell suspensions were digested in 100 µl of Tris-HCl buffer (10 m, pH 9) and 13 µl of proteinase K (1 mg ml−1) at 55°C for 2 h. Then, digested bacterial cells were incubated at 100°C for 10 min to stop the reaction. Five µl of digested bacterial cells suspensions were added to 50 µl of the PCR mix (Laguerre et al., 1994). The 16S rDNAs were amplified by PCR, with the primers fD1 and rD1 (Weisburg et al., 1991), and digested with four discriminative restriction endonucleases AluI, DdeI, Hinf I and MspI, as described by Laguerre et al. (1994). Digested PCR products were separated by horizontal electrophoresis with 3% Nusieve agarose (Tebu-bio, Le Perray en Yvelines, France) in Tris-borate-EDTA buffer (Sambrook et al., 1989) at 60 V cm−1 over 4 h. The 1-kb DNA ladder (Invitrogen, Cergy Pontoise, France) was used as a molecular weight marker. Gels were stained with ethidium bromide and photographed under UV illumination with Polaroid type 665 positivenegative film. Depending on their ARDRA profiles, the bacterial isolates were distributed into five genotypic groups named according to Laguerre et al. (1994) and Frey et al. (1997). Functional fingerprinting of the isolates Preparation of bacterial inocula Bacteria were grown on 10% TSA medium (3 g l−1 Tryptic Soy Broth from Difco and 15 g l−1 agar) at 25°C for 36 h. To prepare the bacterial inocula used in the following tests, three to four bacterial colonies were picked and suspended in 2 ml of sterile deionized water in order to obtain a suspension with A600nm0.7 (about 109 cfu ml−1). In vitro inorganic phosphorus solubilization assay The ability of the bacterial isolates to solubilize tricalcium orthophosphate (TCP) was tested on the TCP medium containing 4 g Ca3(PO4)2, 10 g glucose, 5 NH4Cl, 1 g NaCl, 1 g MgSO4

© New Phytologist (2005) www.newphytologist.org

and 20 g agar per litre at pH = 7.2, distributed in Petri dishes (5.5 cm diameter, 10 ml of medium per dish). For each bacterial isolate, 10 µl of inoculum were dropped in the center of three plates. After incubation at 25°C for 3 d, the clearing of the initially turbid medium indicated phosphate solubilizing bacterial isolates. All the bacterial isolates grew on the TCP agar medium and their solubilizing capacity was the same whatever the replicate. The bacterial isolates were distributed into two classes referred to ‘0’ and ‘+’ depending on their phosporous solubilizing ability. The experiment was repeated twice at two different dates. In vitro siderophore production assay Semi-quantitative analysis of siderophore production by the bacterial isolates was performed following the chrome azurol S (CAS) method of Alexander & Zuberer (1991). The CAS agar medium was prepared according to Alexander and Zuberer’s protocol and distributed in Petri dishes (5.5 cm diameter, 10 ml of medium per dish). For each bacterial isolate, 10 µl of inoculum were dropped in the center of three plates. After incubation at 25°C for 3 d, the discoloration of the medium (blue to orange) indicated siderophore-producing bacterial isolates. All the bacterial isolates grew on the CAS agar medium. Two perpendicular diameters of the discoloration area were measured. According to mean values of the diameters of the discoloration area, the bacterial isolates were distributed into three classes referred to ‘+, ++, +++’, depending on the intensity level of siderophore production. The experiment was repeated twice on two different dates. In vitro IAA production assay Qualitative analysis of indole3-acetic acid (IAA) production by the bacterial isolates was performed following the method of Bric et al. (1991). Bacterial isolates were grown on TSA medium (3 g l−1 Tryptic Soy Broth from Difco and 15 g l−1 agar) at 28°C for 24 h. Then, they were striked as spots with sterile toothpicks on 10% TSA medium containing 5 m tryptophan. After inoculation, plates were immediately overlaid with a nitrocellulose membrane and incubated at 28°C for 72 h. Then the membranes were removed from the plates and dipped in Salkowski reagent (2% 0.5  FeCl3 in 35% perchloric acid) at room temperature for about 1 h. Bacterial strains producing IAA were identified by the formation of red halo on the membrane immediately surrounding the bacterial spots. The bacterial isolates were distributed into two classes referred to ‘0’ and ‘+’ depending on their IAA synthesis ability. The experiment was repeated twice on two different dates. In vitro HCN production assay Bacterial isolates were grown on a HCN induction medium (30 g Tryptic Soy Broth, 4.4 g Glycine, 15 g agar per litre) at 28°C for 4 d (Castric, 1977). For each bacterial isolate, 10 µl of inoculum were dropped in the center of one plate (5.5 cm of diameter, 10 µl of HCN induction medium). Then, a disk of Whatman paper (5.5 cm


319

320 Research

diameter) previously dipped in a HCN revealing solution (0.5% picric acid and 2% Na2CO3) was placed in the lid of the Petri dish and the plate was sealed with air-tight tape. After 4 d incubation at 28°C, an orange-brown color of the paper indicated HCN producing bacterial isolates. The bacterial isolates were distributed into two classes referred to ‘0’ and ‘+’ depending on their HCN synthesis ability. The experiment was repeated twice on two different dates. In vitro fungal growth inhibition assay The 60 bacterial isolates were screened for their ability to inhibit the growth of seven phytopathogenic fungi isolated from forest nurseries or plantations: Rhizoctonia solani strains 263 and 264 obtained from Dr Sen (University of Helsinki, Finland), Fusarium avenaceum D2 and Botrytis cinerea 98G1 obtained from Dr Stenström (Agricultural University of Uppsala, Sweden), one strain of Botrytis spp. obtained from Dr Alabouvette (INRA Dijon, France), one strain of Phytophthora spp. obtained from Dr Hansen (INRA Nancy, France) and one strain of Heterobasion annosum obtained from our collection in Alessandria (Italy). They were grown on Potato Dextrose Agar (PDA) from Difco, at 25°C for 6 d. Six mm-diameter plugs were then cut out from the periphery of the fungal colonies and transferred in the center of four 5.5 cm-diameter Petri dishes containing 10 ml of rich medium (39 g PDA, 5 g agar per litre). For each bacterial isolate, four 10-µL droplets of inoculum were distributed 1.5 cm from the center, in four perpendicular directions. In the control treatment without bacteria, 10-µl droplets of sterile deionized water were used. The fungal colony extension was measured along the four radii after 3–22 d of incubation at 25°C, depending on the growth speed of the fungus. In vitro ectomycorrhizal fungal growth stimulation assay The 60 bacterial strains were screened for their ability to stimulate the growth of three ectomycorrhizal fungi isolated from forest nurseries or plantations: Laccaria bicolor strain S238N, which derives from strain S238 provided by R. Molina and J.M. Trappe (USA) (Di Battista et al., 1996), Thelephora terrestris strain CHA obtained from J.L. Churin (INRA Nancy, France) and Hebeloma crustuliniforme strain KAPP obtained from J. Menez (INRA Nancy, France). Laccaria bicolor and H. crustuliniforme were grown on Potato Dextrose Agar (PDA) from Difco, at 25°C for 1 month. Thelephora terrestris was grown on P5 Agar medium (Pachlewski & Pachlewska, 1974). The protocol used was the same as for the fungal growth inhibition assay, except that the fungal-bacterial confrontations were performed on a poor medium (15 g agar per litre in distilled water), with a 4-wk incubation period. Effect on Douglas fir–L. bicolor mycorrhizal establishment Laccaria bicolor S238N inoculum was prepared by aseptically growing the mycelium in a peat-vermiculite nutrient mix (Duponnois & Garbaye, 1991). Seeds of Douglas fir from


provenance zone 422 (Washigton State, USA) were pretreated in moist peat at 4°C for 1 month to break dormancy. Seedlings were then grown in 80 ml polyethylene containers filled with a peat-vermiculite mix (1 : 1, v/v; pH 5.5), watered to field capacity and mixed with 2.5% (v/v) fungal inoculum. Bacterial inocula were prepared as described by Frey-Klett et al. (1997). The absorbance of each bacterial suspension at 600 nm was adjusted to 1 in 0.1  MgSO4 buffer; then the suspensions were diluted 10 times. Five ml of each bacterial suspension (2 × 107−8 × 107 CFU ml−1, depending on the bacterial isolate) was poured on top of each container immediately after sowing; the controls without bacteria received only 0.1  MgSO4 buffer. One seedling was grown per container. The containers were arranged in trays containing 40 containers. One tray was inoculated for each bacterial isolate. The control treatment comprised three trays. In the glasshouse, the trays were distributed at random on three tables and were moved around weekly according to a circular permutation. Watering with distilled water or nutrient solution was performed as described by Frey-Klett et al. (1997). Fourteen weeks after inoculation, 20 seedlings per tray (i.e. 50%) were randomly sampled. The proportion of short roots forming mycorrhizas with L. bicolor was determined by randomly examining 100 short roots per seedling with a stereomicroscope; L. bicolor ectomycorhizas can be identified on morphological bases. Searching for genes of specific functions Total DNA of each bacterial isolate was purified using DNeasy Tissue Kit according to the manufacturer (Qiagen, Dusseldorf, Germany). Searching for genes of antibiotic synthesis The oligonucleotide primers Phl2a and Phl2b were used to amplify sequences from the gene phlD, which is involved in the synthesis of 2,4-diacetylphloroglucinol (Phl) (Raaijmakers et al., 1997). The oligonucleotide primers PCA2a and PCA3b were used to amplify sequences within phzC and phzD, two genes involved in the synthesis of phenazine-1-carboxylic acid (Phz) (Raaijmakers et al., 1997). The oligonucleotide primers PRND1 (De Souza & Raaijmakers, 2003) and PRN2 () were used to amplify pyrrolnitrin (Prn) genes. The oligonucleotide primers PLTC1 and PLTC2 were used to amplify pyoluteorin (Plt) genes (De Souza et al., 2003). PCR amplification was carried out in 25-µl reaction mixture, which contained approximately 20 ng of total DNA, according to the protocol described by De Souza et al. (2003), with Taq DNA polymerase (Q-Biogen, Illkirch, France) 1 U of for PRN and PLT or 2 U for PHL and PHZ. The PCR program used for PHL and PHZ was described by Raaijmakers et al. (1997). The one for PRN and PLT was described by De Souza et al. (2003). The strains P. fluorescens CHA0, 2–79 and SSB17, from our collection in Wageningen, are known to have one or several of the following antibiotic genes, PHZ,


Research

PHL, PRN or PLT. They were used as positive controls. Samples of the PCR products were separated on a 1.2% agarose gel (Sigma, Saint Quentin Fallavier, France) in 1× TBE buffer (Sambrook et al., 1989) at 75 V cm−1 over 3 h. The 1-kb DNA ladder (Invitrogen, Cergy Pontoise, France) was used as a molecular weight marker. Gels were stained with ethidium bromide and photographed under UV illumination with Polaroid type 665 positive-negative film. Genes for nitrogen fixation A 370 bp fragment of nifH genes was amplified by using nested PCR according to Widmer et al. (1999). The first step of the PCR amplification was performed with the forward primer nifH (forA) and the reverse primer nifH (rev). The second step of the PCR amplification was performed with the forward primer nifH (forB) and the same reverse primer nifH (rev) as in the first step. PCR amplification was carried out in 25-µl reaction mixture which contained 200 µ of each dXTP, 1XPCR Buffer, 0.8 µ of each primer, 1.5 m of MgCl2, 1.25 U of Taq DNA polymerase (Q-Biogen, Illkirch, France). Both PCR amplification steps were done as follows: initial denaturation at 95°C for 5 min followed by 40 cycles (for the first PCR) or 35 cycles (for the nested PCR), of 94°C for 11 s, 92°C for 15 s, 48°C for 8 s, 50°C for 30 s, 74°C for 10 s, 72°C for 10 s, and followed by a 10-min final extension at 72°C (Widmer et al., 1999). One µl of the first PCR product was used as a template for the nested PCR. After the last step, the PCR products were separated on a 2% agarose gel (Sigma) in 1× TAE buffer (Sambrook et al., 1989). The DNA size standard Eurogentec Smartladder was used as a molecular weight marker. Gels were stained with ethidium bromide. The amplified 370-bp fragment obtained was purified with a QIAquick Gel Extraction Kit (Qiagen), then sequenced using the sequencing primer nifH (forB) () on the Applied Biosystems model 310 DNA automated sequencer with BigDye Terminator chemistry. The obtained sequences were subjected to the nonredundant GenBank nucleic acid database by using the algorithm BLASTN (Altschul et al., 1997) for assessing the identity of the sequences. The bacterial isolates were distributed into two classes: ‘−’ = no nifH gene detected, ‘+’ = detection of a sequence which is homologous to the nifH sequences of the NCBI database (% of homology higher than 88%).

treatment) . The mean diameter values of the different bacterial treatments were then compared with those of the control treatment using the Bonferoni-Dunn test (P = 0.05), in order to identify the bacterial isolates that significantly inhibited/stimulated fungal growth. According to these results, for the inhibition assay, the bacterial isolates were distributed into three classes corresponding to the number (n) of phytopathogenic fungal strains significantly inhibited by the same bacterial isolate: ‘0 ≤ n < 3′’, 3 ≤ n < 5′- and ‘5 ≤ n < 8’. Concerning the stimulation assay, the bacterial isolates were distributed into two classes: ‘n < 2’ and ‘2 ≤ n’ with n = number of ectomycorrhizal fungal strains significantly stimulated by the same bacterial isolate. In the specific case of L. bicolor S238N, the bacterial isolates were distributed into two classes depending on their effect on the fungal growth: ‘0’ = no significant effect, ‘+’ = significant stimulation. The percentages of mycorrhizal short roots were arcsin transformed and analysed with one-factor s (bacterial treatment or origin of the strains) and the means were compared with the control or compared two by two using the Bonferoni-Dunn test (P = 0.05). Bacterial distributions Whatever the assays, the bacterial isolates were distributed into different classes according to their behaviour in the in vitro tests. The proportion of bacterial isolates per class were compared between the two zones of origin of the isolates, bulk soil and mycosphere (mycorrhizosphere + mycorrhiza), using a χ2 test (P = 0.05).

Results Genotypic fingerprinting of the bacterial isolates The 60 P. fluorescens isolates studied belonged to five 16S rDNA genotypic groups (Laguerre et al., 1994; Frey et al., 1997). In the bulk soil, only the two genotypic groups 22 and 23 were identified (Fig. 1). In the mycorrhiza and mycorrhizosphere zones, at least one additional genotypic group was detected. Furthermore, the distribution of groups 22 and 23 was highly significantly affected by the zone the isolates originated from (P = 0.0008); the frequency of group 23 was reduced in the mycosphere (mycorrhizosphere + mycorrhiza) zone, whereas that of group 22 was enhanced.

Statistical analyses

Functional fingerprinting of the bacterial isolates

Continuous variables The two variables: mean diameters of the discoloured areas on CAS medium and mean diameters of the L. bicolor colonies, were analysed with one-factor (origin of the bacterial isolate) . The three zones of origin (bulk soil, mycorrhizosphere, mycorrhiza) were then compared using the Bonferoni-Dunn test (P = 0.05). The mean diameters of the pathogenic and ectomycorrhizal fungal colonies were analysed with one-factor (bacterial

In vitro inorganic phosphorus solubilization assay The majority of the bacterial isolates from the mycosphere zone were able to solubilize tri-calcium phosphate in the culture medium, whereas the majority of the isolates from the bulk soil did not (P = 0.03, Fig. 2a).


In vitro siderophore production assay As expected, our 60 isolates of P. fluorescens produced siderophores. Moreover,


321

322 Research

Fig. 1 Distribution of the 60 bacterial isolates according to their zone of origin and their belonging to four 16S rDNA genotypic groups. The proportion of bacterial isolates in the two genotypic groups 22 and 23 (the only ones with at least four isolates for each zone) was significantly different between the bulk soil and the mycosphere zones, according to a χ2 test (P = 0.0008). Light shading, genotype groups 1 and/or 2; dark shading, genotype group 21; solid, genotype group 22; no shading, genotype group 23.

Fig. 2 Distribution of the 60 bacterial isolates according to their zone of origin and following functional traits: (a) solubilisation of inorganic phosphorus, (b) siderophore production, (c) AIA production, and (d) and HCN production. (a) The proportion of bacterial isolates in each of the two classes (solubilisation or nonsolubilisation of inorganic phosphorus) was significantly different between the bulk soil and the mycosphere zones, according to a χ2 test (P = 0.03). No shading, no solubilisation of inorganic phosphorus in vitro; solid, solubilisation of inorganic phosphorus in vitro. (b) The proportion of bacterial isolates in each of the three classes related to the intensity of siderophore production was significantly different between the bulk soil and the mycosphere zones, according to a χ2 test (P = 0.0151). Increasing intensity level of siderophone production in vitro: no shading, lowest intensity; shading, increased intensity; solid, highest intensity. (c) The proportion of bacterial isolates in each of the two classes (production or nonproduction of AIA) was not significantly different between the bulk soil and the mycosphere zones, according to a χ2 test (P = 0.36). No shading, no production of IAA in vitro; solid, production of IAA in vitro. (d) The proportion of bacterial isolates in each of the two classes (production or nonproduction of HCN) was significantly different between the bulk soil and the mycosphere zones, according to a χ2 test (P = 0.0011). No shading, no production of HCN in vitro; solid, production of HCN in vitro.

this semiquantitative test allowed us to classify the bacterial strains according to the quantity/efficiency of the siderophore production, which clearly depended on the origin of the bacterial isolates (Fig. 2b). The frequency of the isolates producing the highest levels of siderophores (and/or the most efficient ones) was significantly higher in the mycosphere zone than in the bulk soil zone (P = 0.0151). This effect of the origin


of the isolates on siderophore production was statistically significant according to one-factor  (P = 0.001). The Bonferoni-Dunn test showed that the bulk soil isolates (mean = 2.54) were significantly different from the mycorrhiza isolates (mean = 5.25). By contrast, the mycorrhizosphere isolates (mean = 2.83) were not significantly different either from those of the bulk soil or those of the mycorrhizas.


Research

Fig. 3 Distribution of the 60 bacterial isolates according to their zone of origin and their following functional traits: (a) inhibition of the in vitro growth of phytopathogenic fungi, (b) stimulation of the in vitro growth of ectomycorrhizal fungi, (c) stimulation of the in vitro growth of L. bicolor S238N, and (d) and effect on Douglas fir–L. bicolor ectomycorrhiza formation. (a) The proportion of bacterial isolates in each of the three classes (no shading, 0 ≤ n < 3; shading, 3 ≤ n < 5; solid, 5 ≤ n < 8; n = number of phytopathogenic fungi significantly inhibited by each bacterial isolate) was significantly different between the bulk soil and the mycosphere zones, according to a χ2 test (P = 0.013). (b) The proportion of bacterial isolates in each of the two classes (no shading, 0 ≤ n < 2; solid, 2 ≤ n < 4; n = number of ectomycorrhizal fungi significantly stimulated by each bacterial isolate) was significantly different between the bulk soil and the mycosphere zones, according to a χ2 test (P = 0.0101). (c) The proportion of bacterial isolates in each of the two classes (stimulation, solid, or nonstimulation, no shading, of the in vitro growth of L. bicolor S238N) was significantly different between the bulk soil and the mycosphere zones, according to a χ2 test (P = 0.0249). (d) The effect on mycorrhiza formation of the bulk soil isolates was significantly different from that of the mycorrhiza and mycorrhizosphere isolates, according to a one-factor ANOVA (origin of the strains) and the Bonferoni-Dunn test (P = 0.0001). No shading, no significant effect on ectomycorrhiza formation; solid, inhibition of ectomycorrhiza formation.

In vitro IAA production assay In the three zones studied, the majority of the bacterial isolates were able to produce IAA (Fig. 2c). There was no significant difference in the proportion of IAA-producing strains between the bulk soil and the mycosphere zones (P = 0.36). In vitro HCN production assay The majority of the bacterial isolates from the bulk soil were able to synthetise HCN, whereas the large majority of the isolates from the mycorrhiza and the mycorrhizosphere zones were not (P = 0.0011, Fig. 2d). In vitro fungal growth inhibition assay Twenty-six percent of the bacterial isolates from the bulk soil zone were unable to inhibit the in vitro growth of the phytopathogenic fungal strains tested. By contrast, all the isolates from the mycorrhizosphere zone inhibited at least one fungal strain and all the isolates from the mycorrhiza zone restricted the growth of at least three fungal strains. The distribution of the


frequency of the bacterial isolates in each class was significantly different between the bulk soil and the mycosphere zones (P = 0.013, Fig. 3a). In vitro ectomycorrhizal fungal growth stimulation assay None of the 60 bacterial isolates was able to promote the in vitro growth of the three ectomycorrhizal fungal strains tested. All the bacterial isolates from the bulk soil zone promoted the growth of one or two fungal strains. By contrast, the majority of the bacterial isolates from the mycorrhiza zone stimulated the growth of one fungal strain only; the others were unable to promote the growth of any fungus. The frequency of the different classes of bacterial isolates was intermediate in the mycorrhizosphere zone (Fig. 3b). This distribution was significant according to a χ2 test (P = 0.0101). In the particular case of the ectomycorrhizal fungus L. bicolor S238N, the bulk soil zone was also characterized by a significantly higher proportion of bacterial isolates able to promote the fungal growth in comparison with the mycosphere zone (P =


323

324 Research

Fig. 4 Distribution of the 60 bacterial isolates according to their zone of origin and the nifH gene detection. The proportion of bacterial isolates in each of the two classes (detection or not of the nifH gene) was not significantly different between the bulk soil and the mycosphere zones according to a χ2 test (P = 0.66). No shading, no detection of the nifH gene; solid, detection of the nifH gene.

0.0249, Fig. 3c). This effect of the origin of the isolates on L. bicolor growth enhancement was statistically significant according to one-factor  (P = 0.002). The BonferoniDunn test showed that the bulk soil isolates were significantly different from the mycorrhiza and mycorrhizosphere isolates. Effect on Douglas fir–L. bicolor mycorrhizal establishment None of the bacterial isolates from the mycorrhiza zone significantly inhibited ectomycorrhiza formation, whereas in the bulk soil zone, 30% of the bacterial isolates did so (Fig. 3d). This effect of the origin of the isolates on mycorrhiza formation was statistically significant according to one-factor  (P = 0.0001). The Bonferoni-Dunn test showed that the effect of the bulk soil isolates was significantly different from that of the mycorrhiza and mycorrhizosphere isolates. Searching for genes of antibiotic synthesis None of the genes of antibiotic synthesis studied could be detected in the 60 bacterial isolates tested, although the genes had been detected in the control strains. Searching for nifH genes The proportion of the bacterial isolates harboring a nifH gene was not significantly different in the mycosphere compared to the bulk soil zone (P = 0.66). Whatever the zone of origin, the proportion of bacterial isolates with a nifH gene was lower than the one without (Fig. 4).

Discussion The tree root–soil interface of forest ecosystems in the boreal and temperate regions consists in a diverse community of ectomycorrhizal short roots. Together with their extramatrical mycelium and associated microorganisms, they form a multitrophic ectomycorrhizal complex, which plays a central role in gross production and nutrient cycling. In this study, we analysed the effect of the ectomycorrhizal symbiosis on the genotypic and functional bacterial diversity and its possible consequences in terms of tree nutrition, development and health. The rhizosphere is generally considered as a zone with higher microbial density and activity but lower specific and


intraspecific diversity than in the bulk soil, resulting from the oriented selection pressure exerted by plant roots (Davet, 1996). Such a decrease of the rhizosphere genetic diversity was demonstrated by Mavingui et al. (1992), who showed that the genetic diversity within populations of Bacillus polymyxa, isolated from the rhizoplane of wheat, was lower than that of the bulk soil. In the same way, Marilley & Aragno (1999), using a culture-independent approach, described a decrease of the phylogenetic diversity of bacterial communities in the rhizosphere of Trifolium and Lolium. By contrast, in the rhizosphere of pine seedlings, Bomberg et al. (2003) observed an increase of the Crenarchaeal diversity. In the present work also, performed with another tree species, the genetic diversity of culturable P. fluorescens was higher in the Douglas fir–L. bicolor mycosphere (mycorrhiza + mycorrhizosphere) than in the bulk soil. The aim of the second part of this work was thus to determine if this increase of the bacterial genotypic diversity in the mycosphere was parallelled with an increase of the bacterial functional diversity. It is well known that, among other plant-beneficial activities, ectomycorrhizal fungi improve the phosphorus uptake of their associated host plants (Read & Perez-Moreno, 2003). This has been mainly ascribed to the fact that the extramatrical mycelium, which extends out from the roots, first absorbs soluble phosphorus (a rare and poorly mobile resource) in a very large volume of soil and transfers it to the plant roots (Smith & Read, 1997), and second mobilizes phosphorus directly from organic matter through the excretion of phosphatases or from minerals through the excretion of organic acids (Landeweert et al., 2001). Biogeochemical weathering of minerals is also performed by other microorganisms, especially phosphate-solubilizing bacteria, which are commonly associated with the roots of a wide range of species (Wenzel et al., 1994; Kim et al., 1997). We showed in the present study that most of the fluorescent pseudomonad isolates from the mycosphere were able to solubilize inorganic phosphate, by contrast to the majority of the bacterial isolates from the bulk soil. It has been shown that some phosphate-solubilizing bacterial strains can interact synergistically with mycorrhizal fungi and promote sustainable phosphorus supply to plants (Kim et al., 1997; Toro et al., 1997; Muthukumar et al., 2001). Therefore, our results suggest that the enrichment of the


Research

mycosphere with phosphate-solubilizing bacterial isolates could contribute to improve the nutrition of Douglas fir seedlings in the nursery soil. By contrast to the commonly accepted role of mycorrhizal fungi in phosphorus nutrition, very few data are available on the effect of these fungi on iron nutrition of plants. In the case of the endomycorrhizal fungi, Caris et al. (1998), who used radiolabelled Fe, demonstrated that the fungus Glomus mosseae can mobilise and/or take up iron from soil and translocate it to the plant. In the case of the ectomycorrhizal fungi, however, the role of the fungi in the root Fe-uptake remains unclear. Ectomycorrhizal fungi produce hydroxamate siderophores (Szaniszlo et al., 1981; Haselwandter, 1995; Renshaw et al., 2002), which are efficient at dissolving minerals, such as biotite, and releasing Fe (Watteau & Berthelin, 1994). Moreover, iron concentration in mycorrhizal plants, grown in nongnotobiotic conditions, is higher than in nonmycorrhizal ones (Leyval & Reid, 1991; Hauer & Dawson, 1996). These data suggest that ectomycorrhizal fungi play a major role in iron nutrition, either by themselves or via their associated microorganisms. In the present work, we demonstrated that the bacterial isolates from the mycosphere produced higher levels of siderophore and/or more efficient ones than the bulk soil isolates. This suggests that the ectomycorrhizal symbiosis has not only a direct effect on iron nutrition, but also an indirect one via the Fe-uptake properties of the mycosphere bacterial communities that are selected by the symbiosis. This hypothesis is in agreement with the results of Crowley et al. (1992), who demonstrated that rhizosphere microorganisms significantly increased the apparent rates of plant iron acquisition from phytosiderophores. Ectomycorrhizal fungi also have access to inorganic and organic soil nitrogen, thus improving nitrogen nutrition of their plant host (Martin & Plassard, 2001; Read & PerezMoreno, 2003). Obviously, these fungi are unable to fix N2 by themselves. However nitrogenase activities were detected in sporocarps of different ectomycorrhizal fungi due to the presence of associated N2-fixing bacteria (Li & Hung, 1987). In forest ecosystems, high rates of N2-fixation were already reported (Limmer & Drake, 1996), but their origin remains unclear. Indeed, it is difficult to distinguish the contribution of the free-living N2-fixing bacteria that live in the soil, from that of the diazotrophs located in the mycorrhizosphere, a niche that is suited for these bacteria (Sen, 2000). The present work revealed that the proportion of the P. fluorescens isolates with a nifH gene, the unique gene required for nitrogen fixation (Widmer et al., 1999), did not differ significantly between the mycosphere and the bulk soil zones. Therefore the Douglas fir–L. bicolor symbiosis did not enrich the mycosphere with potential N2-fixing P. fluorescens isolates. By contrast, Rózycki et al. (1999) showed a significant increase in the N2-fixing bacteria, mainly Pseudomonas, in the mycorrhizosphere of pine and oak trees in a mixed conifer-hardwood forest.


Ectomycorrhizal fungi synthesise different phytohormones, especially indole-3-acetic acid (IAA), which is involved in the formation and the functioning of the ectomycorrhizal symbiosis (Karabaghli et al., 1997). Likewise, many species of rhizosphere and soil bacteria synthesise IAA, which can mediate plant growth promotion (Patten & Glick, 1996). In the present work, contrary to the majority of the other criteria analysed, we did not observe any significant effect of the symbiosis on the distribution of the IAA-producing Pseudomonads. This result contrasts with the one of Lebuhn et al. (1997) who demonstrated that the proportion of IAA producing Paenibacillus polymyxa strains decreases from the bulk soil zone to the rhizosphere and the rhizoplane of wheat. Mycorrhizal fungi are known to enhance plant resistance to soil-borne pathogens (Duchesne, 1994; Azcón-Aguilar & Barea, 1996; Graham, 2001). Several direct mechanisms are involved such as: first, production of antibiotic substances by the mycorrhizal fungi (Sylvia & Sinclair, 1983; Tsantrizos et al., 1991) and/or the associated plant roots (Duchesne, 1994); second, mechanical exclusion of the pathogens by the mantle of the ectomycorrhizas (Duchesne, 1994); and third, competition between mycorrhizal fungi and other biotrophic pathogens for the root cortex and its nutrient resources (Graham, 2001). The increased vigour and the modification of the physiology of mycorrhizal plants also indirectly contribute to the disease suppression. In the same way, changes in the population of pathogen antagonists in the mycorrhizosphere could indirectly explain many reported protective effects of the mycorrhizal symbiosis against plant diseases (Linderman, 1994). In the present study, the antagonistic ability of the 60 bacterial isolates was assayed in vitro. In the context of screening biocontrol agents against soil-borne pathogens, the significance of the in vitro antagonism assays is often controversial because the efficiency of biocontrol agents depends not only on their antagonism ability but also on their rhizosphere competence (Whipps, 1987; Lugtenberg et al., 2001). Nevertheless, the in vitro antibiosis assays constitute an efficient way to analyse the antagonistic potential of a bacterial community (Whipps, 1987). Interestingly, we demonstrated that the Douglas fir–L. bicolor mycorrhizas and ectomycorrhizosphere selected P. fluorescens isolates that had a broader spectrum of antagonistic ability than the bulk soil isolates. This suggests that the ectomycorrhizal symbiosis causes a shift in the P. fluorescens populations that could contribute to the known protection effect of the symbiosis. These results are in accordance with those of Citernesi et al. (1996) who detected many bacteria antagonistic to Fusarium and Phytophthora in the mycorrhizosphere of Glomus mosseae grown in pot cultures for 17 yr. Other bacteria isolated from the roots of Douglas fir seedlings, harvested from nurseries and forest sites, also exhibited broad-spectrum antagonistic activities, not only against fungal pathogens, but also against human ones (Axelrood et al., 1996).


325

326 Research

Among the antibiotic substances involved in the biocontrol of phytopathogens, HCN is a toxic secondary metabolite that is produced by many rhizosphere microorganisms, especially fluorescent pseudomonads (Dowling & O’Gara, 1994). Here, the proportion of HCN producing bacterial isolates was significantly higher in the bulk soil than in the mycosphere. Because L. bicolor is very sensitive to HCN (data not shown), this suggests that the Douglas fir–L. bicolor symbiosis counter-selected HCN-producing bacterial isolates. Laccaria bicolor may thus exert a negative selection pressure on the bacterial communities. Many fluorescent pseudomonads also produce a variety of other antimicrobial metabolites, such as phloroglucinols, phenazines, pyoluteorin and pyrrolnitrin, which are major determinants of the antagonism of these bacteria against phytopathogens (Dowling & O’Gara, 1994). The biosynthetic genes of these antibiotics are conserved in pseudomonads from diverse geographic locations (Raaijmakers et al., 1997) but none of the genes coding for these antibiotics was detected in our 60 P. fluorescens isolates. This result contrasts with that of Bedini et al. (1999), who isolated phloroglucinol-producing Pseudomonads strains from fruit bodies of the ectomycorrhizal fungus Tuber borchii. Our results showed therefore that the broad antagonistic spectrum against phytopathogenic fungi, which characterizes the P. fluorescens isolates from the mycorrhiza and the mycorrhizosphere zones is not correlated to the capacity of these isolates to synthesis antibiotics such as HCN, phloroglucinols, phenazines, pyoluteorin and pyrrolnitrin. Further studies will therefore be necessary to elucidate the mechanisms of the antagonistic activity of the mycosphere pseudomonad isolates and perhaps to detect some still unknown antibiotics or other mechanisms. Some bacteria isolated from the soil, the ectomycorrhizas or the sporocarps of some ectomycorrhizal fungi can significantly promote the growth of these fungi in vitro (Garbaye & Bowen, 1989; Varese et al., 1996; Founoune et al., 2002). However, to our knowledge, nothing is known about the proportion of these helper bacterial isolates in the different soil zones (bulk soil, mycorrhizosphere soil, mycorrhizas). In our study, we showed that the proportion of bacterial isolates able to promote the growth of different ectomycorrhizal fungi was significantly higher in the bulk soil than in the mycosphere zones. This suggests that the soil acts as a reservoir for bacteria having a beneficial effect on the presymbiotic growth of the ectomycorhizal fungi, before the root-fungus encounter. In the same way, we showed that in the particular case of L. bicolor, the proportion of fungal growth promoting bacteria is significantly higher in the bulk soil zone compared with the mycosphere one, where the proportion is quite low (10%). This small proportion of fungal growth promoting bacteria in the mycosphere zone is in accordance with the results of Sbrana et al. (2002) who demonstrated that 19% of the bacterial strains isolated from Quercus robur – T. borchii ectomycorrhizas were able to significantly promote the growth of T. borchii in vitro.


The promotion of the survival and /or the presymbiotic growth of the ectomycorrhizal fungi in the soil was proposed by Garbaye (1994) as one of the five hypotheses that could explain the positive effect of mycorrhiza helper bacteria on the establishment of the mycorrhizal symbiosis. This hypothesis was confirmed in the case of the Douglas fir–L. bicolor S238N symbiosis and the mycorrhiza helper P. fluorescens strain BBc6R8 (Brulé et al., 2001). Duponnois (1992) found a highly significant correlation between the effect of bacterial isolates from Douglas fir – L. bicolor S238N ectomycorrhizas or associated sporocarps on the in vitro growth of the fungus, and their effect on mycorrhiza formation. In this study, as the percentage of short roots mycorrhizal with L. bicolor in the control treatment without inoculated bacteria was close to 100%, we were unable to detect any positive effect of the bacterial inoculation. However, we could demonstrate that there was a significantly higher proportion of inhibiting bacteria in the bulk soil zone, in comparison with the mycosphere one. This result reveals that the Douglas fir – L. bicolor symbiosis counter-selects the bacteria that have a negative impact on mycorrhiza formation. To conclude, our work demonstrates that, in addition to its direct positive effect on plant growth via nutritional and hormonal mechanisms, the ectomycorrhizal symbiosis also has an indirect positive effect via its selective pressure on bacterial communities, which should be considered when studying the functional role of the ectomycorrhizal symbiosis in situ. Indeed, we showed that the ectomycorrhiza symbiosis determines the structure of the P. fluorescens populations in the nursery soil and selects bacteria potentially beneficial to the symbiosis and to the plant. This underlines the existence of multitrophic microbial associations in the ectomycorrhizal complex and completely supports Linderman’s metaphor: ‘mycorrhizas are like the quaterback of the microbial herd: mycorrhizas call out signals to the microbial herd, which in turn acquires nutrients to feed back to the plant’ (R.G. Linderman, unpublished).

Acknowledgements This study was funded by the Lorraine Region and by the French and Italian Ministries of Foreign Affairs through the “Galilée” programme. F. Martin is acknowledged for useful discussions on the manuscript and E. Gamalero for her technical assistance. We thank R. Molina and J.M. Trappe (Corvallis, OR, USA) for providing the strain S238, from which S238N was derived. We also thank R. Sen (University of Helsinki, Finland), E. Stenström (Agricultural University of Uppsala, Sweden), C. Alabouvette (INRA Dijon, France), and E. Hansen (INRA Nancy, France), for providing the phytopathogenic fungal strains.

References Alexander DB, Zuberer DA. 1991. Use of chrome azurol S reagents to evaluate siderophore production by rhizosphere bacteria. Biology and Fertility of Soils 12: 39–45.


Research Altschul SF, Madden TL, Schäffer AA, Zhang J, Zhang Z, Miller W, Lipman DJ. 1997. Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Research 25: 3389–3402. Ames RN, Reid CPP, Ingham ER. 1984. Rhizosphere bacterial population responses to root colonization by a vesicular-arbuscular mycorrhizal fungus. New Phytologist 96: 555 – 563. Andrade G, Mihara KL, Linderman RG, Bethlenfalvay GJ. 1997. Bacteria from rhizosphere and hyphosphere soils of different arbuscular-mycorrhizal fungi. Plant and Soil 192: 71 – 79. Axelrood PE, Clarke AM, Radley R, Zemcov SJV. 1996. Douglas-fir root-associated microorganisms with inhibitory activity towards fungal pathogens. Canadian Journal of Microbiology 42: 690 – 700. Azcón-Aguilar C, Barea JM. 1996. Arbuscular mycorrhizas and biological control of soil-borne plant pathogens-an overview of the mechanisms involved. Mycorrhizas 6: 457 – 464. Bedini S, Bagnoli G, Sbrana C, Leporini C, Tola E, Dunne C, Filippi C, D’Andrea F, O’Gara F, Nuti MP. 1999. Pseudomonads isolated from within fruit bodies of Tuber borchii are capable of producing biological control or phytostimulatory compounds in pure culture. Symbiosis 26: 223–236. Bomberg M, Jurgens G, Saano A, Sen R, Timonen S. 2003. Nested PCR detection of Archaea in defined compartments of pine mycorrhizospheres developed in boreal forest humus microcosms. FEMS Microbiology Ecology 43: 163–171. Bric JM, Bostock RM, Silverstone SE. 1991. Rapid in situ assay for indolacetic acid production by bacteria immobilized on a nitrocellulose membrane. Applied and Environmental Microbiology 57: 535 – 538. Brulé C, Frey-Klett P, Pierrat JC, Gérard F, Lemoine MC, Rousselet JL, Sommer J, Garbaye J. 2001. Survival in the soil of the ectomycorrhizal fungus Laccaria bicolor and effect of a mycorrhiza helper Pseudomonas fluorescens. Soil Biology and Biochemistry 33: 1683 – 1694. Caris C, Hördt W, Hawkins HJ, Römheld V, George E. 1998. Studies of iron transport by arbuscular mycorrhizal hyphae from soil to peanut and sorghum plants. Mycorrhiza 8: 35 – 39. Castric PA. 1977. Glycine metabolism by Pseudomonas aeruginosa: hydrogen cyanide biosynthesis. Journal of Bacteriology 130: 826 – 831. Christensen H, Jakobsen I. 1993. Reduction of bacterial growth by a vesicular-arbuscular mycorrhizal fungus in the rhizosphere of cucumber (Cucumis sativus L.). Biology and Fertility of Soils 15: 253 – 258. Citernesi AS, Fortuna P, Filippi C, Bagnoli G, Giovannetti M. 1996. The occurrence of antagonistic bacteria in Glomus mosseae pot-cultures. Agronomie 16: 671–677. Cromack JK, Fichter BL, Moldenke AM, Entry JA, Ingham ER. 1988. Interactions between soil animals and ectomycorrhizal fungal mats. Agriculture and Ecosystem Environments 24: 161 – 168. Crowley DE, Römheld V, Marschner H, Szaniszlo PJ. 1992. Root-microbial effects on plant iron uptake from siderophores and phytosiderophores. Plant and Soil 142: 1 – 7. Davet P. 1996. Vie microbienne du sol et production végétale. Paris, France: Editions INRA. De Souza JT, Raaijmakers JM. 2003. Polymorphisms within the prnD and pltC genes from pyrrolnitrin and pyoluteorin-prducing Pseudomonas and Burkholderia spp. FEMS Microbiology Ecology 43: 21 – 34. Denison RF, Bledsoe C, Kahn M, O’Gara F, Simms EL, Thomashow LS. 2003. Cooperation in the rhizosphere and the ‘free rider’ problem. Ecology 84: 838–845. Di Battista C, Selosse MA, Bouchard D, Stenström E, Le Tacon F. 1996. Variations in symbiotic efficiency, phenotypic characters and ploidy level among different isolates of the ectomycorrhizal basidiomycete Laccaria bicolor strain S238. Mycological Research 100: 1315 – 1324. Dowling DN, O’Gara F. 1994. Metabolites of Pseudomonas involved in the biocontrol of plant disease. Tibtech 12: 133 – 141. Duchesne LC. 1994. Role of ectomycorrhizal fungi in biocontrol. In: Pfleger FL, Linderman RG, eds. Mycorrhizae and plant health. St Paul, MN, USA: APS Press, 27–45.


Duponnois R. 1992. Les bactéries auxiliaires de la mycorhization du Douglas (Pseudotsuga menziesii (Mirb.) Franco) par Laccaria Laccata souche S238. PhD Thesis, University of Nancy, France. Duponnois R, Garbaye J. 1991. Mycorrhization helper bacteria associated with the Douglas fir-Laccaria laccata symbiosis: effects in aseptic and in glasshouse conditions. Annales des Sciences Forestières 48: 239–251. Eastmond PJ, van Dijken AJH, Spielman M, Kerr A, Tissier AF, Dickinson HG, Jones JDG, Smeekens SC, Graham IA. 2002. Trehalose-6-phosphate syntase 1, which catalyses the first step in trehalose synthesis, is essential for Arabidopsis embryo maturation. Plant Journal 29: 225–235. Founoune H, Duponnois R, Bâ AM, Sall S, Branget I, Lorquin J, Neyra M, Chotte JL. 2002. Mycorrhiza Helper Bacteria stimulate ectomycorrhizal symbiosis of Acacia holosericea with Pisolithus albus. New Phytologist 153: 81–89. Frey P, Frey-Klett P, Garbaye J, Berge O, Heulin T. 1997. Metabolic and genotypic fingerprinting of fluorescent pseudomonads associated with the Douglas-fir–Laccaria bicolor mycorrhizosphere. Applied and Environmental Microbiology 63: 1852–1860. Frey-Klett P, Pierrat JC, Garbaye J. 1997. Location and survival of mycorrhiza helper Pseudomonas fluorescens during establishment of ectomycorrhizal symbiosis between Laccaria bicolor and Douglas fir. Applied and Environmental Microbiology 63: 139–144. Garbaye J. 1994. Helper bacteria: a new dimension to the mycorrhizal symbiosis. New Phytologist 128: 197–210. Garbaye J, Bowen GD. 1989. Stimulation of ectomycorrhizal infection of Pinus radiata by some microorganisms associated with the mantle of ectomycorrhizas. New Phytologist 112: 383–388. Graham JH. 2001. What do root pathogens see in mycorrhizas? New Phytologist 149: 357–359. Haselwandter K. 1995. Mycorrhizal fungi siderophore production. Critical Reviews in Biotechnology 15: 287–291. Hauer RJ, Dawson JO. 1996. Growth and iron sequestring of pin oak (Quercus palustris) seedlings inoculated with soil containing ectomycorrhizal fungi. Journal of Arboriculture 22: 122–130. Heinonsalo J, Jorgensen KS, Haahtela K, Sen R. 2000. Effects of Pinus sylvestris root growth and mycorrhizosphere development on bacterial carbon source utilization and hydrocarbon oxidation in forest and petroleum-contaminated soils. Canadian Journal of Microbiology 46: 451–464. Jentschke G, Bonkowski M, Godbold DL, Scheu S. 1995. Soil protozoa and forest tree growth: non-nutritional effects and interaction with mycorrhizae. Biology and Fertility of Soils 20: 263–269. Johansson JF, Paul LR, Finlay RD. 2004. Microbial interactions in the mycorrhizosphere and their significance for sustainable agriculture. FEMS Microbiology Ecology 48: 1–13. Karabaghli C, Sotta B, Gay G. 1997. Hormones fongiques, ectomycorhizes et rhizogénèse. Revue Forestière Française. XLIX: 99–109. Kim KY, Jordan D, McDonald GA. 1997. Effect of solubilizing bacteria and vesicular-arbuscular mycorrhizae on tomato growth and soil microbial activity. Biology and Fertility of Soils 26: 79–87. King EO, Ward MK, Raney DE. 1954. Two simple media for the demonstration of pyocyanine and fluorescein. Journal of Laboratory and Clinical Medicine 44: 301–307. Laguerre G, Rigottier-Gois L, Lemanceau P. 1994. Fluorescent Pseudomonas species categorized by using polymerase chain reaction (PCR) /restriction fragment analysis of 16S rDNA. Molecular Ecology 3: 479–487. Landeweert R, Hoffland E, Finlay R, Kuyper TW, van Breemen N. 2001. Linking plants to rocks: ectomycorrhizal fungi mobilize nutrients from minerals. Trends in Ecology and Evolution 16: 248–254. Lebuhn M, Heulin T, Hartmann A. 1997. Production of auxin and other indolic and phenolic compounds by Paenibacillus polymyxa strains isolated from different proximity to plant roots. FEMS Microbiology Ecology 22: 325–334.


327

328 Research Lelu A. 1994. Clusters and factors: neural algorithms for a novel representation of huge and highly multidimensional data sets. In: Diday Y, Lechevallier M, Schader M, Bertrand P, Burtchy B, eds. New approaches in classification and data analysis. Berlin, Germany: Springer-Verlag KG, 241–248. Leyval C, Berthelin J. 1993. Rhizodeposition and net release of soluble organic compounds by pine and beech seedlings inoculated with rhizobacteria and ectomycorrhizal fungi. Biology and Fertility of Soils 15: 259–267. Leyval C, Reid CPP. 1991. Utilization of microbial siderophores by mycorrhizal and non-mycorrhizal pine roots. New Phytologist 119: 93–98. Li CY, Hung LL. 1987. Nitrogen-fixing (acetylene-reducing) bacteria associated with ectomycorrhizae of Douglas fir. Plant and Soil 98: 425–428. Limmer C, Drake HL. 1996. Nonsymbiotic N2 fixation by acidic and pH-neutral forest soils: aerobic and anaerobic differentials. Soil Biology Biochemistry 28: 177–183. Linderman RG. 1988. Mycorrhizal interactions with the rhizosphere microflora: the mycorrhizosphere effect. Phytopathology 78: 366 –371. Linderman RG. 1994. Role of VAM Fungi in Biocontrol. In: Pfleger FL, Linderman RG, eds. Mycorrhizae and plant health. St Paul, MN, USA: APS Press, 1–25. Lugtenberg BJJ, Dekkers L, Bloemberg GV. 2001. Molecular determinants of rhizosphere colonization by Pesudomonas. Annual Review of Phytopathology 39: 461–490. Marilley L, Aragno M. 1999. Phylogenetic diversity of bacterial communities differing in degree of proximity of Lolium perenne and Trifolium repens roots. Applied Soil Ecology 13: 127 –136. Martin F, Plassard C. 2001. Nitrogen assimilation by ectomycorrhizal symbiosis. In: Morot-Gaudry J-F, ed. Nitrogen assimilation by plants. Enfield, NH, USA: Science Publishers, 169 –183. Mavingui P, Laguerre G, Berge O, Heulin T. 1992. Genetic and phenotypic diversity of Bacillus polymyxa in soil and in the wheat rhizosphere. Applied and Environmental Microbiology 58: 1894 – 1903. Meyer JR, Linderman RG. 1986. Selective influence on populations of Rhizosphere or rhizoplane bacteria and actinomycetes by mycorrhizas formed by Glomus fasciculatum. Soil Biology and Biochemistry 18: 191–196. Muthukumar T, Udaiyan K, Rajeshkannan V. 2001. Response of neem (Azadirachta indica A. Juss) to indigenous arbuscular mycorrhizal fungi, phosphate-solubilizing and asymbiotic nitrogen-fixing bacteria under tropical nursery conditions. Biology and Biofertility of Soils 34: 417–426. Neal JL, Bollen JR, Bollen WB. 1964. Rhizosphere microflora associated with mycorrhizae of Douglas fir. Canadian Journal of Microbiology 10: 259–265. Olsson PA, Chalot M, Bååth E, Finlay RD, Söderström B. 1996. Ectomycorrhizal mycelia reduce bacterial activity in sandy soil. FEMS Microbiology Ecology 21: 77 – 86. Pachlewski R, Pachlewska J. 1974. Studies on symbiotic properties of mycorrhizal fungi of pine ( Pinus Sylvestris) with the aid of the method of mycorrhizal synthesis in pure culture on agar. Warsaw, Poland: Forest Research Institute. Patten CL, Glick BR. 1996. Bacterial biosynthesis of indole-3-acetic acid. Canadian Journal of Microbiology 42: 207 –220. Raaijmakers JM, Weller DM, Thomashow LS. 1997. Frequency of antibiotic- producing Pseudomonas spp. in natural environments. Applied and Environmental Microbiology 63: 881 – 887. Rambelli A. 1973. The rhizosphere of mycorrhizae. In: Marks GC, Kozlowski TT, eds. Ectomycorrhizae: their ecology and physiology. New York, USA: Academic Press, 299 – 343. Read DJ, Perez-Moreno J. 2003. Mycorrhizas and nutrient cycling in ecosystems – a journey towards relevance? New Phytologist 157: 475–492.


Renshaw JC, Robson GD, Trinci APJ, Wiebe MG, Livens FR, Collison D, Taylor RJ. 2002. Fungal siderophores: structures, functions and applications. Mycological Research 106: 1123–1142. Rózycki H, Dahm H, Strzelczyk E, Li CY. 1999. Diazotrophic bacteria in root-free soil and in the root zone of pine (Pinus sylvestris L.) and oak (Quercus robur L.). Applied Soil Ecology 12: 239–250. Rygiewicz PT, Andersen CP. 1994. Mycorrhizae alter quality and quantity of carbon allocated below ground. Nature 369: 58–60. Sambrook J, Fritsch EF, Maniatis T. 1989. Molecular cloning: a laboratory manual, 2nd edn. New York, NY, USA: Cold Spring Harbor Laboratory. Sbrana C, Agnolucci M, Bedini S, Lepera A, Toffanin A, Giovannetti M, Nuti MP. 2002. Diversity of culturable bacterial populations associated to Tuber borchii ectomycorrhizas and their activity on T. borchii mycelial growth. FEMS Microbiology Letters 211: 195–201. Schreiner RP, Bethlenfalvay GJ. 1995. Mycorrhizal interactions in sustainable agriculture. Critical Review in Biotechnology 15: 271–285. Sen R. 2000. Budgeting for the wood-wide web. New Phytologist 145: 161–163. Smith SA, Read D. 1997. Mycorrhizal symbiosis, 2nd edn. London, UK: Academic Press. Sylvia DM, Sinclair WA. 1983. Suppressive influence of Laccaria laccata on Fusarium oxysporum and on Douglas-fir seedlings. Phytopathology 73: 384–389. Szaniszlo PJ, Powell PE, Reid CPP, Cline GR. 1981. Production of hydroxamate siderophore iron chelators by ectomycorrhizal fungi. Mycologia 73: 1158–1174. Timonen S, Jorgensen KS, Haahtela K, Sen R. 1998. Bacterial community structure at defined locations of Pinus sylvestris-Suillus bovinus and Pinus sylvestris-Paxillus involutus mycorrhizospheres in dry pine forest humus and nursery peat. Canadian Journal of Microbiology 44: 499–513. Toro M, Azcón R, Barea JM. 1997. Improvement of arbuscular mycorrhiza development by inoculation of soil with phosphate-solubilizing rhizobacteria to improve rock phosphate bioavailability (32P) and nutrient cycling. Applied and Environmental Microbiology 63: 4408–4412. Tsantrizos YS, Kope HH, Fortin JA, Ogilvie KK. 1991. Antifungal antibiotics from Pisolithus tinctorius. Phytochemistry 30: 1113–1118. Varese GC, Portinaro S, Trotta A, Scannerini S, Luppi-Mosca AM, Martinotti G. 1996. Bacteria associated with Suillus grevillei sporocarps and ectomycorrhizae and their effects on in vitro growth of the mycobiont. Symbiosis 21: 129–147. Wamberg C, Christensen S, Jakobsen I, Müller AK, Sorensen SJ. 2003. The mycorrhizal fungus (Glomus intraradices) affects microbial activity in the rhizosphere of pea plants (Pisum sativum). Soil Biology and Biochemistry 35: 1349–1357. Waschkies C, Schropp A, Marschner H. 1994. Relations between grapevine replant disease and root colonization of grapevine (Vitis sp.) by fluorescent pseudomonads and endomycorrhizal fungi. Plant and Soil 162: 219 – 227. Watteau F, Berthelin J. 1994. Microbial dissolution of iron and aluminium from soil minerals: efficiency and specificity of hydroxamate siderophores compared to aliphatic acids. European Journal of Soil Biology 30: 1–9. Weisburg WG, Barns SM, Pelletier DA, Lane DJ. 1991.16S ribosomal DNA amplification for phylogenetic study. Journal of Bacteriology 173: 697–703. Wenzel CL, Ashford AE, Summerell BA. 1994. Phosphate-solubilizing bacteria associated with proteid roots of seedlings of waratah (Telopea speciosissima (Sm.) R.Br.). New Phytologist 128: 487–496. Whipps JM. 1987. Effect of media on growth and interactions between a range of soil-borne glasshouse pathogens and antagonistic fungi. New Phytologist 107: 127–142. Widmer F, Shaffer BT, Porteous LA, Seidler RJ. 1999. Analysis of nifH gene pool complexity in soil and litter at a Douglas fir forest site in the Oregon cascade montain range. Applied and Environmental Microbiology 65: 374–380.
