Soil Biology & Biochemistry 34 (2002) 1875–1881 www.elsevier.com/locate/soilbio
Phosphorus uptake of an arbuscular mycorrhizal fungus is not effected by the biocontrol bacterium Burkholderia cepacia Sabine Ravnskova,b,*, John Larsena, Iver Jakobsenb a
Department of Crop Protection, Danish Institute of Agricultural Sciences, Research Centre Flakkebjerg, DK-4200 Slagelse, Denmark b Plant Research Department, Risø National Laboratory, DK-4000 Roskilde, Denmark Received 27 February 2002; received in revised form 21 August 2002; accepted 3 September 2002
Abstract The biocontrol bacterium Burkholderia cepacia is known to suppress a broad range of root pathogenic fungi, while its impact on other beneficial non-target organisms such as arbuscular mycorrhizal (AM) fungi is unknown. Direct interactions between five B. cepacia strains and the AM fungus, Glomus intraradices (BEG87) were studied in root-free soil compartments separated from a rooting compartment by a fine nylon-mesh. B. cepacia had no effect on AM fungal biomass and energy reserves measured using the signature fatty acid 16:1v5 from phospholipid fatty acids (PLFAs) and neutral lipid fatty acids (NLFAs), respectively. Hyphal P transport was also unaffected by the biocontrol bacterium, which either stimulated, reduced or had no effect on length of the external mycelium of G. intraradices. The cyclic PLFAs cy17:0 and cy19:0 were suggested to be useful markers for estimation of biomass of B. cepacia. The presence of mycelium of G. intraradices reduced the biomass of three out of five B. cepacia strains as indicated by a reduction in PLFAs cy17:0 and cy19:0, while other bacterial PLFAs were unaffected by mycelium of G. intraradices. On the other hand, two out of five B. cepacia strains reduced the amount of branched PLFAs suggesting a reduction in the population of Gram-positive bacteria in these cases. In conclusion, the B. cepacia seems to have no impact on neither mycorrhiza formation nor on the functioning of the AM fungus G. intraradices in terms of P transport, whereas our results suggest that mycorrhiza might have adverse effects on B. cepacia. q 2002 Elsevier Science Ltd. All rights reserved. Keywords: Arbuscular mycorrhiza; Biocontrol bacteria; Microbial interactions; Signature fatty acids
1. Introduction Burkholderia cepacia is a rhizosphere-colonising bacterium with competitive saprobic ability, that is recognized as a potential biocontrol agent against several root pathogens (Bowers and Parke, 1993; Roberts et al., 1997; Mao et al., 1998). The mode of action of B. cepacia is not clear, but evidence of production of anti-fungal compounds of some strains of B. cepacia has been found, and direct interactions between B. cepacia and root pathogens in terms of antibiosis have been proposed (Homma et al., 1989; Burkhead et al., 1994). In order to use biocontrol agents in plant production, it is important to study their possible negative impact on other plant beneficials such as arbuscular mycorrhizal (AM) * Corresponding author. Address: Department of Crop Protection, Danish Institute of Agricultural Sciences, Research Centre Flakkebjerg, DK-4200 Slagelse, Denmark. Tel.: þ 45-58113469; fax: þ 45-58113301. E-mail address:
[email protected] (S. Ravnskov).
fungi, and if indigenous microorganisms associated with plant roots influence growth of the biocontrol agents. AM fungi are obligate, biotrophic symbionts associated with roots of most herbaceous plants. An external hyphal network proliferates from the root into the soil and transport phosphorus from the soil into the host plant (Smith and Read, 1997). Walley and Germida (1997) showed that B. cepacia inhibits spore germination of the AM fungus Glomus clarum, and this may have been the reason for a reduction in mycorrhiza colonisation of wheat roots in a field after application of B. cepacia as a seedcoat (Germida and Walley, 1996). Non-volatile diffusible substances produced by B. cepacia were suggested to be responsible for this suppression (Walley and Germida, 1997). Signature fatty acids has been used to study soil microbial communities (Frostega˚rd et al., 1993a, 1993b), and to estimate biomass and energy reserves of specific organisms in soil (Olsson et al., 1995; Larsen et al., 1998; Green et al., 1999) and roots (Larsen and Bødker, 2001).
0038-0717/02/$ - see front matter q 2002 Elsevier Science Ltd. All rights reserved. PII: S 0 0 3 8 - 0 7 1 7 ( 0 2 ) 0 0 2 0 1 - 8
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The fatty acid signature16:1v5 can be used to quantify biomass and amount of energy reserves of AM fungi in roots and soil (Olsson, 1999). The objective of the present work was to study possible interactions between the AM fungus G. intraradices and the biocontrol bacterium B. cepacia. The use of a compartmentalized growth system with root-free soil compartments allowed us to study direct interactions between these microbes without interference from roots. Functioning of the AM fungus was examined in terms of hyphal P transport, and phospholipid fatty acids (PLFAs) were used to quantify individual microbes and back ground microbial communities.
2. Materials and methods 2.1. Experimental design Non-mycorrhizal and mycorrhizal cucumber plants were grown in a compartmentalized, cross-shaped growth system with a central root compartment and two lateral root-free compartments. The root-free compartments were separated from the root compartment with nylon mesh with a 20 mm mesh diameter, which allowed passage of hyphae, but not roots. The experiment had 12 treatments; two of mycorrhiza (with and without) and six of bacteria (with five different strains and without) and each treatment had four replicates giving a total of 48 growth units. The experiment was replicated, but with two mycorrhizal treatments (with and without) and three bacterial treatments (none and strains Bc2 and MRL100), only. 2.2. Organisms involved The AM fungus, G. intraradices Schenck and Smith (BEG87), was grown in symbiosis with cucumber Cucumis sativus L. (Aminex, F1 hybrid). The inoculum of G. intraradices consisted of AM colonized roots, spores and soil from a dried pot culture with Trifolium subterraneum L. This inoculum was uniformly mixed into the soil in an amount corresponding to 10% (w/w) of the soil in the mycorrhizal treatments, whereas the soil in the non-mycorrhizal treatments consisted of the soil – sand mix only. Five strains of B. cepacia were used: Bc1, Bc2, BcF, MRL100 and HPQM100. All strains were obtained from Dr Daniel P. Roberts, United States Department of Agriculture, Beltsville, MD, USA. The bacterial strains were originally isolated from the rhizosphere of corn. Strains Bc1 and Bc2 belong to B. cepacia genomovar I –III. Strain MRL100 is taxonomically very different from the other strains and does not fall within any of the genomovars. Inoculum of the five strains of B. cepacia were prepared by growing them in nutrient broth (Difco 0003-15) at 37 8C on a rotary shaker (200 rpm). After 24 h the cells were harvested by centrifu-
gation for 10 min at 7000 rpm at 20 8C and resuspended in 0.9% NaCl. Following two washes in 0.9% NaCl, they were resuspended in sterile distilled water. The number of colony forming units (cfu) was counted after plating on nutrient agar (Difco 0001-15). In order to reintroduce soil microorganisms accompanying the AM inoculum all pots received 10 ml filtrate prepared by sieving a suspension of 100 g of mycorrhiza inoculum in 1 l distilled water through a nylon mesh (20 mm). 2.3. Experimental setup The soil was an irradiated (10 kGy, 10 MeV electron beam) 1/1 (w/w) mixture of sandy loam (Jensen and Jakobsen, 1982) and sand containing 8 mg P kg21 soil, determined after NaHCO3-extraction (Olsen et al., 1954). The following nutrients were mixed into the soil (mg kg21 dry soil): K2SO4, 71.0; CaCl2X5H2O, 71.0; CuSO4X5H2O, 2.0; ZnSO4X7H2O, 5.0; MnSO4XH2O, 10.0; CoSO4X7H2O, 0.35; Na2MoO4X2H2O, 0.18; MgSO4X7H2O, 20.0. Plants were grown in the main compartment of the growth unit in 740 g soil to sand mixture. The two lateral root-free compartments were each filled with 60 g of the soil to sand mixture. Two planting holes were made in the soil surface in the root compartments of each growth unit and one ml of a 1010 cfu bacterial suspension was added to each hole immediately before the pre-germinated seed were transferred. Two pre-germinated cucumber seeds were transferred to each growth unit and plants were thinned to one per growth unit after emergence. Plants were maintained in a growth chamber, equipped with Osram daylight lamps (HQI-T 250 W/D) providing a photosynthetic active radiation of 500 –550 mmol m22 s21 for 16 h per day. The day and night temperatures were 20 and 16 8C, respectively. Pots were watered daily to 65% of the water holding capacity of the soil. Twenty-five milligram of nitrogen was added to each pot as an NH4NO3 solution every week. After 14 d of plant growth, the soil in the root-free compartments was replaced with 60 g soil to sand mixture inoculated with the bacterial strains. This soil was watered to 20% of the water holding capacity with the bacterial suspension whereupon the bacteria were uniformly mixed into the soil to an average density of 3.4 £ 108 cfu g21 soil. One of the compartments was used for radioactive labeling, whereas the other was used for non-radioactive analyses. An aqueous solution of carrier-free H33 3 PO4 was mixed into the soil at 4 kBq g21 soil to a total of 240 kBq in one of the rootfree compartments of each growth system. Plants were harvested 5 weeks after planting. 2.4. Harvest and analyses Dry weights of shoots were determined after drying at 80 8C for 24 h and the shoots were then ground for analyses
S. Ravnskov et al. / Soil Biology & Biochemistry 34 (2002) 1875–1881 Table 1 Signature fatty acids used in the present experiment Fatty acid
iso15:0 anteiso15:0 iso16:0 16:1v5 10methyl16:0 iso17:0 anteiso17:0 cyclo17:0 17:0 cyclo19:0
Specificity Group
Within group
Signature
Bacteria Bacteria Bacteria Bacteria and fungi Bacteria Bacteria Bacteria Bacteria Bacteria Bacteria
Gram positive Gram positive Gram positive Gram negative, Zygomycetes Actinomycetes Gram positive Gram positive Gram negative
Burkholderia
Gram negative
Burkholderia
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automatically controlled all gas chromatography operations including calibration, subsequent sample sequencing, peak integration, and naming. Calibration standards contained a mixture of straight chain saturated and hydroxy fatty acid methyl esters with a length of 10 –20 carbon (MIDI Part No. 1200A). Specificity of signature fatty acids used in present
Glomus
of P and 33P contents. Fresh root subsamples were cleared and stained as described by Kormanik and McGraw (1982) except that trypan blue was used instead of acid fuchsin. Total root lengths and root lengths colonized by the AM fungus were measured in accordance with Newman (1965). Other fresh roots subsamples were dried at 80 8C for 24 h, weighed and ground for analyses of 33P content. Ground samples of shoots and roots were digested in a 4– 1 nitric acid to perchloric mixture and the 33P content in the digests was measured by liquid scintillation counting (Packard TR 1900). Hyphal length of G. intraradices in root-free soil samples was estimated using the membrane-filter technique as described by Jakobsen et al. (1992). The hyphal length density in root-free soil from treatments without mycorrhiza was subtracted from the values obtained in the corresponding treatments with mycorrhiza. Samples of three grams soil from the non-radioactive root-free compartments were subjected to lipid extraction according to the method used by Frostega˚rd et al. (1991). The extracted lipids were fractionated on silicic acid columns (100/200 mesh; Unisil) into neutral, intermediate, and polar fractions by elution with 5 ml of chloroform, 20 ml of acetone, and 5 ml of methanol, respectively. Phospholipids and neutral lipids were dried under nitrogen, with 25 mg ml21 nonadecanoate (fatty acid methyl ester 19:0) added as an internal standard. Lipids in both fractions were then transformed into free fatty acid methyl esters by a mild alkaline methanolysis (Dowling et al., 1986). Analysis of fatty acid methyl esters were performed using the software package Sherlock Version 6.0 (MIDI Inc.) with the HP Chemstation and a HP5890 CG fitted with a 25 m fused silica capillary column (HP part No.19091B-102) and hydrogen as carrier gas. The injector temperature was 250 8C and the detector temperature was 300 8C. The temperature program was as follows: initial temperature 170 8C increasing to 270 8C at 5 8C min21. One microliter of sample preparation was injected. The MIDI software
Fig. 1. Hyphal length density of the arbuscular mycorhhizal fungus, Glomus intraradices, (1a), concentration of the phospholipid fatty acid signature 16:1v5 (1b) and the neutral fatty acid 16:1v5 (1c) in soil from root-free compartments with or without inoculation of five different isolates of the bacterium B. cepacia. Bars represent standard error of the means of four replicates.
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experiment is summarized in Table 1. Fatty acid composition of pure cultures of the B. cepacia strains grown on tryptic soy agar for two days at 30 8C were also analyzed. Levels of significance of main treatments and their interactions were calculated by analysis of variance after testing for normality and variance homogeneity by Bartletts test.
3. Results The average shoot and root dry weight of all plants was 2.22 and 0.36 g, respectively, and the average root length of all plants was 14.1 m. Mycorrhizal plants developed 18% less shoot mass and 40% less root length compared to that of non-mycorrhizal plants, whereas root dry weight was unaffected by mycorrhizas (data not shown). All plant growth parameters were unaffected by bacterial inoculation. The average percent root colonisation by mycorrhiza was 65% in roots from treatments with mycorrhiza inoculum added and was unaffected by the bacterial treatments. Roots from treatments without mycorrhiza inoculum added remained non-mycorrhizal (data not shown). The hyphal length in soil from treatments without mycorrhiza was 2.6 m g21 soil and was unaffected by bacteria inoculations. This background was subtracted from hyphal length measured in the mycorrhizal treatments. Compared to treatments without bacteria added, the hyphal length density of G. intraradices in root-free soil was 43% higher in treatments with B. cepacia Bc2, 45% lower in treatments with B. cepacia MRL100 and 53% lower in treatments with B. cepacia HPQM100. The remaining two
B. cepacia strains Bc1 and BcF had no effect on G. intraradices hyphal length (Fig. 1(a)). The G. intraradices biomass as determined by the signature PLFA 16:1v5, after subtraction of the corresponding background value in nonmycorrhizal plants (0.35 nmol g21 soil), was also differentially affected by the B. cepacia strains. The presence of B. cepacia BcF increased the PLFA 16:1v5 by 49%, whereas B. cepacia HPQM100 decreased this signature by 44% (P ¼ 0.09) (Fig. 1(b)). On the other hand, the amount of the G. intraradices energy reserve marker neutral lipid fatty acids (NLFAs) 16:1v5 was unaffected by the bacterial inoculations (Fig. 1c). The amount of NLFA 16:1v5 in rootfree soil without mycorrhiza was 0.11 nmol g21 soil. Hyphal 33P transport of G. intraradices was unaffected by the bacterial inoculations (Fig. 2). 33P contents in nonmycorrhizal plants were 3.3 kBq and this background value was subtracted from hyphal P transport measured in mycorrhizal plants. The PLFA composition of cells from pure culture of the five B. cepacia strains revealed contents of the PLFAs cy17:0 and cy19:0 in the range 15– 28 and 8 – 24% of total content of PLFA’s, respectively (Table 2). PLFAs cy17:0 and cy19:0 was 97– 172 and 14 – 41% higher, respectively, in soil from treatments with B. cepacia, compared to that of soil without B. cepacia inoculation (Table 3). In treatments with the B. cepacia strains Bc2, BcF, and HPQM100 the presence of G. intraradices reduced the amount of PLFA cy17:0 by 22– 25%. The amount of the PLFA cy19:0 followed the same pattern as found with cy17:0. The presence of G. intraradices had no effect on the amount of PLFA cy17:0 in treatments without B. cepacia and in treatments with B. cepacia strains Bc1 and MRL100. In
Fig. 2. Contents of 33P in cucumber plants grown in symbiosis with the mycorrhizal fungus, G. intraradices and either not inoculated with bacteria or inoculated with five different isolates of the bacterium B. cepacia. Bars represent standard error of the means of four replicates.
S. Ravnskov et al. / Soil Biology & Biochemistry 34 (2002) 1875–1881
In the present work, however, we observed no impact of B. cepacia on the living biomass of the AM fungus, G. intraradices, and in the functioning in terms of P transport. Similarly, Ravnskov and Jakobsen (1999) found that Pseudomonas fluorescens DF57 had no effect on hyphal P transport by two AM fungi, and Barea et al. (1998) found that various Pseudomonas strains with biocontrol activity did not affect mycorrhiza formation by Glomus mosseae. The hyphal length of G. intraradices was either enhanced (Bc2) or reduced (MRL100) by B. cepacia. These findings were later confirmed in the replicate experiment using the same experimental conditions and organisms. Ravnskov and Jakobsen (1999) observed no effect of P. fluorescens DF57 on hyphal growth of G. intraradices, whereas hyphal length density of G. caledonium was higher in soil with P. fluorescens DF57 than in soil without the bacterial inoculation. Similarly, Vosatka and Gryndler (1999) found an enhancing effect of P. putida on growth and phosphatase activity of the external mycelium of G. fistulosum grown in symbiosis with maize, but not with potato. Their results suggest that the stimulating effect of the bacterium on the AM external mycelium was due to the low molecular (MW , 10 000) fraction of the liquid P. putida culture. Another mechanism could be the release of nutrients to the AM fungus via decomposition of organic material by B. cepacia. However, the mechanisms behind these different effects of the bacterial strains on AM fungal growth are not clear. Biomass and energy reserves of G. intraradices measured with specific fatty acids were, on the other hand, unaffected by B. cepacia. The contrasting result
Table 2 Composition (%) of PLFA in pure cultures of five strains of B. cepacia obtained from 2 d old cultures grown on tryptic soy agar B. cepacia strain PLFA
Bc1
Bc2
BcF
MRL 100
HPQM 100
14:0 15:0 16:1v7 16:1v5 16:0 17:0 cyclo 17:0 16:1 2OH 16:0 2OH 18:1v7 18:1v5 18:0 19:0 cyclo 18:1 2OH
0.2 0.2 9.1 0.3 18.9 14.5 0.3 2.7 2.6 33.5 0.2 1.1 11.2 4.8
0 0.3 4.4 0.3 19.1 18.3 0.4 2.8 3.4 18.7 0.4 0.9 23.5 6.7
0.3 0 7.9 0.4 16.9 18.5 0.3 1.0 1.4 37.6 0 0.8 10.1 4.4
0.2 0.2 8.7 0.3 20.6 18.2 0.3 1.8 1.9 26.0 0 1.1 15.2 5.5
0.2 0 10.1 0.3 15.4 15.7 0.4 2.8 2.0 37.9 0 0.7 7.7 6.8
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general, B. cepacia, especially the strains Bc1, MRL100 and HPQM100, reduced the signature fatty acids of Grampositive bacteria i17:0 and i16:0 (Table 1 and 3).
4. Discussion The use of B. cepacia as a biocontrol agent against root pathogens might have adverse effects on AM fungi (Germida and Walley, 1996), which play a key role in plant nutrition in transporting phosphorus to the host plant.
Table 3 Concentration (nmol g21 soil dwt) of common bacterial phospholipid fatty acids in soil from root-free compartments from treatments with and without mycorrhiza (M) and inoculation with different strains of B. cepacia (B) Treatments
i15:0
a15:0
i16:0
m16:0
i17:0
a17:0
cy17:0
17:0
cy19:0
LSD0.05
2.08 1.86 2.22 2.19 2.52 2.10 2.13 2.00 2.00 2.48 1.88 1.71 ns
1.58 1.60 1.44 1.44 1.75 1.58 1.57 1.53 1.58 1.75 1.57 1.42 0.24
0.92 0.92 0.75 0.74 0.94 0.88 0.94 0.88 0.86 0.92 0.86 0.77 0.12
1.52 1.60 1.25 1.22 1.37 1.27 1.31 1.10 1.22 1.57 1.73 1.26 0.29
0.91 0.73 0.52 0.71 0.76 0.67 0.64 0.63 0.65 0.70 0.76 0.75 0.16
0.40 0.51 0.35 0.40 0.48 0.45 0.45 0.43 0.41 0.43 0.49 0.44 ns
1.15 1.07 2.27 2.07 3.13 2.35 3.03 2.37 2.52 2.56 2.96 1.96 0.57
0.54 0.57 0.44 0.49 0.53 0.50 0.51 0.50 0.50 0.47 0.92 0.42 ns
1.50 1.26 1.71 1.58 2.12 1.78 2.12 1.87 1.74 1.71 2.06 1.51 0.40
ANOVA B M M*B
P values 0.13 0.51 0.38
* 0.54 0.36
*** 0.36 0.52
** 0.29 **
* 0.83 **
0.67 0.38 0.62
*** *** ***
0.39 0.84 0.53
*** ** 0.55
B
M
–
2 þ 2 þ 2 þ 2 þ 2 þ 2 þ
Bc1 Bc2 BcF MRL100 PHQM100
a (anteiso), cy (cyclo), i (iso), m (methyl).
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obtained from measuring G. intraradices with signature fatty acids or hyphal length measurements were later confirmed in a replicate experiment. Since the turnover of fatty acids are much faster than chitin, the fatty acid-based method gives a more precise measurement of the actual living biomass, whereas hyphal length measurements include both dead and living biomass (Olsson, 1999). Consequently, the measured hyphal length density of G. intraradices in the present experiment provides information about the influence of the B. cepacia strains on mycelial growth of G. intraradices in earlier phases of the experiment. Inoculating soil with B. cepacia coincided with an increase in the cyclic PLFAs cy17:0 and cy19:0. As these PLFAs constitute up to 28% of the total amount of PLFAs in B. cepacia, they might be considered as a method to estimate biomass of B. cepacia. The relatively high amount of these cyclic PLFAs in soil without B. cepacia inoculation originates from the indigenous populations of B. cepacia and/or other Gram-negative bacteria where cyclic PLFAs are common (Wilkinson, 1988). Signature fatty acids have been used to study interactions between specific microbes co-existing in roots (Larsen and Bødker, 2001) and soil (Larsen et al., 1998; Green et al., 1999). The present experiment adds further to the list of possible signature fatty acids such as the cyclic PLFAs cy17:0 and cy19:0 for B. cepacia that can be used in studies on soil microbial ecology under unsterile, but controlled experimental conditions. Co-inoculation of three out of five B. cepacia strains with G. intraradices reduced the amount of cy17:0 and cy19:0 suggesting that mycelium of G. intraradices reduced the growth of B. cepacia and/or other Gram-negative bacteria. Accordingly, previous experiments have reported negative effects of AM fungi on presence and activity of another bacterial biocontrol agent (Marschner and Crowley, 1996; Ravnskov et al., 1999). These results indicate that the effect of some biocontrol agents on pathogens might be reduced under natural conditions where most plants are mycorrhizal. It is, however, important to consider the environment in which the interactions are studied. The objective of the present work was to study the direct microbial interaction without interfering effects from roots. The outcome of the interaction between B. cepacia and G. intraradices might, however, be different in the rhizosphere, where B. cepacia has direct access to carbohydrates from the roots. Nevertheless, significant high amounts of the signature fatty acids cy17 and cy19 in treatments with B. cepacia, and the effect of B. cepacia on hyphal length of G. intraradices and on the bacterial community in the root-free soil showed, that the bacteria did function even in an environment with low content of carbohydrates. Other bacterial PLFAs not present in B. cepacia were unaffected by mycelium of G. intraradices, whereas two out of five B. cepacia strains reduced the amount of branched PLFAs common in Gram positive bacteria (O’Leary and Wilkinson, 1988), and thereby altered the composition of
the bacterial community in the soil. The different effects of the five B. cepacia strains on Gram-positive bacteria could, as well as the variable effects on AM hyphal length, be due to different ability of the bacterial strains to compete for nutrients, to decompose organic matter and to secrete secondary metabolites, which influence growth of other microorganisms. In conclusion, the B. cepacia seems to have no impact on either mycorrhiza formation or the functioning of the AM fungus G. intraradices in terms of P transport, whereas our results suggest that mycorrhiza might have adverse effects on B. cepacia.
Acknowledgements This work was supported by the Danish Agricultural and Veterinary Research Council, and by the Danish Ministry of Food, Agriculture and Fisheries. We thank Anette Olsen, Anne Olsen and Tina Tønnersen for excellent technical support. We thank Dr Daniel Roberts for providing strains of B. cepacia.
References Barea, J.M., Andrade, G., Bianciotto, V., Dowling, D., Lohrke, S., Bonfante, P., O’Gara, F., Azcon-Aguilar, C., 1998. Impact on Arbuscular mycorrhiza formation of Pseudomonas strains used as inoculants for biocontrol of soil-borne fungal plant pathogens. Applied and Environmental Microbiology 64, 2304–2307. Bowers, J.H., Parke, J.L., 1993. Epidemiology of Pythium damping-off and Aphanomyces root rot of peas after seed treatment with bacterial agents for biological control. Phytopathology 83, 1466–1473. Burkhead, K.D., Schisler, D.A., Slininger, P.J., 1994. Pyrrolnitrin production by biological-control agent Pseudomonas cepacia B37W in culture and in colonized wounds of potatoes. Applied and Environmental Microbiology 60, 2031–2039. Dowling, N.J.E., Widdel, F., White, D.C., 1986. Phospholipid ester-linked fatty acid biomarkers of acetate oxidizing sulphate reducers and other sulphide-formimg bacteria. Journal of General Microbiology 132, 1815–1820. ˚ ., Tunlid, A., Ba˚a˚th, E., 1991. Microbial biomass measured as Frostega˚rd, A total lipid phosphate in soils of different organic content. Journal Microbiological Methods 14, 151–163. ˚ ., Ba˚a˚th, E., Tunlid, A., 1993a. Shift in the structure of soil Frostega˚rd, A microbial communities in limed forests as revealed by phospholipid fatty acid analysis. Soil Biology & Biochemistry 25, 723–730. ˚ ., Tunlid, A., Ba˚a˚th, E., 1993b. Phospholipid fatty acid Frostega˚rd, A composition, biomass and activity of microbial communities from two soil types experimentally exposed to different heavy metals. Applied and Environmental Microbiology 59, 3605–3617. Germida, J.J., Walley, F.L., 1996. Plant growth-promoting rhizobacteria alter rooting patterns and arbuscular mycorrhizal fungi colonization of field-grown spring wheat. Biology and Fertility of Soils 23, 113–120. Green, H., Larsen, J., Olsson, P.A., Jakobsen, I., 1999. Suppression of the biocontrol agent Trichoderma harzianum by mycelium of the arbuscular mycorrhizal fungus Glomus intraradices in root-free soil. Applied and Environmental Microbiology 65, 1428–1434. Homma, Y., Sato, Z., Hirayama, F., Konno, K., Shirahama, H., Suzui, T., 1989. Production of antibiotics by Pseudomonas cepacia as an agent for
S. Ravnskov et al. / Soil Biology & Biochemistry 34 (2002) 1875–1881 biological control of soilborne plant pathogens. Soil Biology & Biochemistry 21, 723 –728. Jakobsen, I., Abbott, L.K., Robson, A.D., 1992. External hyphae of vesicular-arbuscular mycorrhizal fungi associated with Trifolium subterraneum L. 1. Spread of hyphae and phosphorus inflow into roots. New Phytologist 93, 410– 413. Jensen, A., Jakobsen, I., 1982. The occurence of vesicular–arbuscular mycorrhiza in barley and wheat grown in some Danish soils with different fertilizer treatments. Plant and Soil 55, 403–414. Kormanik, P.P., Mcgraw, A.C., 1982. Quantification of vesicular arbuscular mycorrhiza in plant roots. In: Schenck, N.C., (Ed.), Methods and Principles of Mycorrhizal Research, American Phytopathological Society, St paul, MN, pp. 37– 45. Larsen, J., Bødker, L., 2001. Interactions between pea root-inhabiting fungi examined using signature fatty acids. New Phytologist 149, 487–493. Larsen, J., Olsson, P.A., Jakobsen, I., 1998. The use of fatty acid signatures to study mycelial interactions between the arbuscular mycorrhizal fungus Glomus intraradices and the saprotrophic fungus Fusarium culmorum. Mycological Research 102, 1492–1496. Mao, W., Lewis, J.A., Lumsden, R.D., Hebbar, K.P., 1998. Biocontrol of selected soilborne diseases of tomato and pepper plants. Crop Protection 17, 535–542. Marschner, P., Crowley, D.E., 1996. Physiological activity of a bioluminescent Pseudomonas fluorescens (strain 2–79) in the rhizosphere of mycorrhizal and non-mycorrhizal pepper (Capsicum annuum L). Soil Biology & Biochemistry 28, 869 –876. Newman, E.I., 1965. A method of estimating the total length of roots in a sample. Journal of Applied Ecology 3, 139 –145. O’Leary, W.M., Wilkinson, S.G., 1988. Gram-positive bacteria. In: Ratledge, C., Wilkinson, S.G. (Eds.), Microbial Lipids, vol. 1. University of Hull, Hull, UK, pp. 117–185. Olsen, S.R., Cole, C.V., Watanabe, F.S., Dean, L.A., 1954. Estimation of
1881
available phosphorus in soils by extraction with sodium bicarbonate, Circular No. 939, United States Department of Agriculture, Washington, DC. Olsson, P.A., 1999. Signature fatty acids provide tools for determination of the distribution and interactions of mycorrhizal fungi in soil. FEMS Microbiology Ecology 29, 303– 310. Olsson, P.A., Ba˚a˚th, E., Jakobsen, I., So¨derstro¨m, B., 1995. The use of phospholipid and neutral lipid fatty acid analysis to estimate biomass of arbuscular mycorrhizal fungi. Mycological Research 99, 623 –629. Ravnskov, S., Jakobsen, I., 1999. Effects of Pseudomonas fluorescens DF57 on growth and P uptake of two arbuscular mycorrhizal fungi in symbiosis with cucumber. Mycorrhiza 8, 329– 334. Ravnskov, S., Nybroe, O., Jakobsen, I., 1999. Influence of an arbuscular mycorrhizal fungus on Pseudomonas fluorescens DF57 in rhizosphere and hyphosphere soil. New Phytologist 142, 113 –122. Roberts, D.P., Dery, P.D., Mao, W., Hebbar, P.K., 1997. Use of a colonization deficient strain of Escherichia coli in strain combinations for enhanced biocontrol of cucumber seedling diseases. Journal of Phytopathology 145, 461– 463. Smith, S., Read, D., 1997. Mycorrhizal Symbiosis, Academic Press, San Diego, CA. Vosatka, M., Gryndler, M., 1999. Treatment with culture fractions from Pseudomonas putida modifies the development of Glomus fistulosum mycorrhiza and the response of potato and maize plants to inoculation. Applied Soil Ecology 11, 245–251. Walley, F.L., Germida, J.J., 1997. Response of spring wheat (Triticum aestivum ) to interactions between Pseudomonas species and Glomus clarum NT4. Biology and Fertility of Soils 24, 365–371. Wilkinson, S.G., 1988. Gram-negative bacteria. In: Ratledge, C., Wilkinson, S.G. (Eds.), Microbial Lipids, vol. 1. University of Hull, Hull, UK, pp. 299 –557.