Plant root carbohydrates affect growth behaviour of endophytic microfungi

FEMS Microbiology Ecology 41 (2002) 161^170

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Plant root carbohydrates a¡ect growth behaviour of endophytic microfungi Franz Hadacek  , Gu«nther F. Kraus Department of Comparative and Ecological Phytochemistry, Institute of Botany, University of Vienna, Rennweg 14, A-1030 Vienna, Austria Received 23 January 2002; received in revised form 25 April 2002; accepted 30 April 2002 First published online 28 June 2002

Abstract Peucedanum alsaticum and Peucedanum cervaria represent characteristic umbellifers (Apiaceae) of calcareous grasslands in Central and Eastern Europe. Both accumulate glucose, fructose, mannitol and sucrose as dominant carbohydrates in their roots. The objective of the study was to determine if endophytes utilise host plant carbohydrates differently than rhizosphere and bulk soil microfungi. Inula ensifolia (Asteraceae), Lathyrus latifolius (Fabaceae) and Bromus erectus (Poaceae), all plants that grow along with the two umbellifers, accumulated only sucrose as major sugar in their roots. Germ tube growth of 30 microfungal isolates, recovered from various rhizosphere habitats, was quantified in microdilutions (10^5000 Wg ml31 ) of a number of substrates, including glucose, sucrose, mannitol and the water-soluble plant root carbohydrate mixtures. Multivariate analysis of variance and subsequent least significant difference analysis of isolate group means revealed that sucrose and mannitol utilisation affected affiliation to a certain fungal lifestyle. Endophytes utilised host plant carbohydrates more efficiently at lower concentrations. Conversely, higher concentrations slowed their growth. Non-host carbohydrates did not cause comparable effects. The results suggest that root carbohydrate diversity may determine fungal diversity in natural rhizosphere environments. 8 2002 Federation of European Microbiological Societies. Published by Elsevier Science B.V. All rights reserved. Keywords : Plant^fungus interaction ; Endophyte; Rhizosphere soil ; Bulk soil; Sucrose; Glucose; Mannitol; Substrate utilisation ; Germ tube growth; Peucedanum alsaticum ; Peucedanum cervaria; Inula ensifolia ; Lathyrus latifolius ; Bromus erectus

1. Introduction Sloughed-o¡ cells, mucilage and exudates from plant roots represent essential carbon sources for soil microbes such as fungi and bacteria. Root exudates are plant-speci¢c and their composition may also be in£uenced by physiological conditions or abiotic factors [1^4]. Roots of intact plants harbour an extensive diversity of fungi: arbuscular mycorrhizal fungi of the order Glomales occur in roots of nearly all plants; various asco- and basidiomycetes form an ectomycorrhizal mantle of hyphae around the roots of trees and shrubs; and endophytes comprise asexually reproducing fungi that spend the whole or greater part of their life cycle inside healthy plant tissues but cause no obvious disease symptoms [5^7].

* Corresponding author. Fax: +43 (1) 4277 9541. E-mail address : [email protected] (F. Hadacek).

Plant biodiversity is generally considered an important factor that contributes to the functioning and stability of terrestrial ecosystems [8,9]. Microbial communities in the rhizosphere may a¡ect plant communities [10,11] and, conversely, plants may in£uence microbial communities in their rhizosphere [12,13]. These ¢ndings have provided a ¢rst step in understanding mechanisms that regulate structure and function of natural plant^microbe communities [14]. Association of fungi with plants dates far back to the arrival of vascular plants [15]. Many studies focus on mycorrhizal fungi as crucial components in the rhizosphere [16]. Endophytes that are the focus of this study are necrotrophic microfungi. The majority of them are common soil fungi that can be found in the rhizosphere as well as in the bulk soil. Such microfungi do not only constitute a big portion of fungal biodiversity in soils, a number of them may develop into perthotrophic pathogens that cause severe damage to crop plants and dramatically reduce yields [17]. In natural systems, pathogen outbreaks do not compare, but still are known to occur [18]. Thus, the diversity of their lifestyles

0168-6496 / 02 / $22.00 8 2002 Federation of European Microbiological Societies. Published by Elsevier Science B.V. All rights reserved. PII : S 0 1 6 8 - 6 4 9 6 ( 0 2 ) 0 0 2 9 2 - 1

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provides us with a unique scenario to study mechanisms involved in adaptation to host plants. Conventional isolation techniques prevent us from recovering the whole spectrum of fungal diversity from plant tissues [19]. Anamorph microfungi that can be recovered by soil-washing include genera such as Acremonium, Cylindrocarpon or Penicillium. Without too much e¡ort, these strains can also be maintained on arti¢cial media. This provides us with a spectrum of test organisms that allows us to identify possible low molecular signal molecules involved in adaptation towards a lifestyle of an endophyte. Plant compounds comprise primary metabolites that fungi utilise as nutrients, such as sugars and amino acids [4]. The major portion of chemical biodiversity in plants, however, is created by secondary metabolism. Plant roots also accumulate and may also exude numerous terpenoids, coumarins, £avonoids or polyacetylenes, just to name a few of them. The majority of authors concur that the main function of secondary metabolites is to contribute to the plant’s survival by acting as chemical defence against microbial and herbivore predators [20,21]. Independently, plant metabolites may also adopt a further function: as an attractive or behaviour-changing signal to various micro-organisms or animals [22,23]. Peucedanum alsaticum and Peucedanum cervaria represent typical umbellifers of calcareous grasslands in Central and Eastern Europe [24]. Preliminary experiments indicated that, despite the structural diversity of secondary metabolites present in the roots of these two umbellifers, inhibition caused by non-water-soluble secondary metabolites was more uniform than growth stimulation by water-soluble carbohydrates. Thus we focused the role of the water-soluble carbohydrates in the adaptation of microfungi to our model plants, P. alsaticum and P. cervaria. For comparative purposes, Bromus erectus, the dominant grass species, pea Lathyrus latifolius and composite Inula ensifolia were also included. No e⁄cient in situ method for capture, identi¢cation and quanti¢cation of root exudates exists [25]. In an e¡ort to make our experiments practicable and detect di¡erences in utilisation spectra as e⁄ciently as possible, carbohydrate mixtures isolated directly from the plant roots and selected major components of those extracts were o¡ered in concentration gradients to selected microfungal isolates recovered from bulk soil, rhizosphere soil and root tissues of the two Peucedanum species.

2. Materials and methods 2.1. Plant material Eichkogel (367 m asl; long. 16‡17P, lat. 48‡03P) is a small hill located near the town of Mo«dling, south of Vienna, Austria, and still carries more or less undisturbed grassland areas. Tap root material of the two umbellifers,

P. alsaticum L. and P. cervaria (L.) Lapeyr. was uncovered to about 30 cm below the soil surface and coarsely freed from surrounding soil. Similarly, underground organs of the dominant grass species, B. erectus Huds. (Poaceae) and two other nearby growing herbs, I. ensifolia L. (Asteraceae) and L. latifolius L. (Fabaceae), were also collected. Voucher specimens (P13278BIO-01^P13278BIO-05) were deposited at the herbarium of the Institute of Botany, University of Vienna (WU). 2.2. Extraction of water-soluble root carbohydrates Ground root material (10^20 g) was exhaustively extracted with methanol (MeOH) at room temperature for 3 days. The whole crude extract was evaporated to a watery residue on a rotary evaporator, dissolved in 100 ml deionised water, and twice extracted with approximately 300 ml chloroform to remove lipophilic compounds. The water-soluble phase was then evaporated to about 20 ml and stored at 4‡C for further use. 2.3. High performance liquid chromatography (HPLC)^UV analyses of the water-soluble root extract fraction Portions of the extracts were evaporated to dryness, dissolved in methanol and adjusted to a concentration of 10 mg ml31 , of which 10 Wl was injected. A Hewlett Packard 1090 Series II HPLC equipped with an UV diode array detector was used. The column was a C18 Hypersil BDS (250U4.6 mm, 5 Wm), the column oven was thermostatted at 40‡C ; binary solvent gradient started at 100% aqueous bu¡er (o-phosphoric acid (Merck) 0.015 mol, tetrabutylammonium hydroxide (Sigma) 0.0015 mol, pH 3) and rose to 40% acetonitrile (Merck, gradient grade) in 20 min, £ow rate was 1 ml min31 ; signal wavelength was 230 nm, UV spectra were recorded from 220 to 450 nm. 2.4. Gas chromatography coupled with mass spectrometry (GC^MS) analyses of the water-soluble extract fraction Water-soluble extract fractions were derivatised into their trimethylsilyl (TMS)-oxime ether/esters as described [26]. A Perkin Elmer GC AutoSystem gas chromatograph (GC) coupled to a Perkin Elmer Turbomass quadrupole mass spectrometer (MS) was used for analysis. The column was a Perkin Elmer PE-5ms (20 mU0.18 mm, 0.18 Wm ¢lm thickness); the GC oven was programmed to stay at 60‡C for 2 min, to rise to 100‡C at a rate of 20‡C min31 , to rise to 250‡C at a rate of 5‡C min31 , to rise to 330‡C at a rate of 8‡C min31 , and to hold 330‡C for 4 min; helium was used as carrier gas at a constant £ow rate of 1 ml min31 ; the transfer line to the MS was

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thermostatted at 280‡C and the ion source at 180‡C. MS spectra were recorded from 30 to 620 amu per 0.6 s in the electron ionisation mode at an electron energy of 70 eV and a ¢lament emission of 200 WA. One Wl of sample was injected in the split mode, the split £ow was adjusted to 10 and 30 ml min31 , depending on analyte concentration. Temperature of the injector was programmed to stay at 60‡C for 2 min, and then to rise to 320‡C at a rate of 180‡C min31 . Identi¢cation of sugar, sugar alcohol, organic acid and amino acid trimethylsilyl-oxime ether/esters was achieved (1) by comparison with MS spectra and retention times of authentic standards, (2) tentatively, by comparison of mass spectra to the Wiley Registry of Mass Spectral Data, 6th edition or the NIST/EPA/NIH Mass Spectral Library 1.5a, and to a MS library and retention time database of TMS compounds o¡ered for free download from the web site of the Max Planck Institute of Molecular Plant Physiology, Potsdam, Germany for metabolic pro¢ling [27].

163

tria. Identi¢cation was carried out based on morphological characters [34,35]. 2.6. Conidia suspensions Petri dishes containing 2% malt extract agar (20 ml) were three-point inoculated with cryoconserved strains and grown for 2^4 weeks at room temperature in the dark. Conidia were harvested by suspending in 1% (w/v) sodium chloride solution containing 5% (v/v) dimethylsulfoxide. Conidia suspension was ¢ltered (pore width 53 or 105 Wm, depending on conidia size), concentrated by centrifuging (4000Ug) and transferred into 1.8 ml cryotubes and stored at 320‡C. Colony forming units (CFU) were determined by plating 20 Wl of 10U serially diluted suspensions and counting germinated spores/dilution concentration. Procedures were replicated three times for each suspension. 2.7. Germ tube growth assays

2.5. Fungal isolates Recovery of microfungi from soil, rhizosphere and root tissue was carried out several times from April to November in two consecutive years. Root pieces (ca. 5 cm long and 0.5^1.5 cm in diameter) were transferred into glass vials containing 1% (w/v) sodium chloride solution. Soil samples were taken during the digging up of roots ; 1^2 g of bulk soil was transferred into glass vials. Processing of the samples on the same day turned out to be mandatory. Soil fungi were isolated by the soil-plating [28] and soilwashing technique [29,30]. The latter method was modi¢ed by using sieve sizes of 0.8, 0.4, 0.2 and 0.1 mm. Mineral and organic particles retained on the smallest-sized sieves (0.2 and 0.5 mm) were transferred to 2% malt extract (Merck) and corn meal (Difco) agar-plates. All media used for isolation purposes contained chloramphenicol (Merck) or validamycin (Solacol0 , Aventis Crop Science) as antibiotic additives (0.1 g ml31 ) [31]. Rhizosphere soil was recovered by vigorously shaking roots in 1% (w/v) sodium chloride solution for 15 min and subjected to the same isolation procedures. Root tissue was thoroughly washed in tap water to remove soil and mucilage as e⁄ciently as possible. Standard surface sterilisation was performed by treatment with 70% ethanol and subsequent £aming [32,33]. Root pieces (5^10U2^3 mm) were transferred to agar plates containing medium identical to that used for isolation of soil microfungi. Conidia and hyphae of pure isolates were suspended in an aqueous solution of sucrose (140 g l31 ) and peptone (10 g l31 ) and stored at 380‡C. Transfer to freezing temperatures was performed by decreasing temperature not faster than 1‡C min31 . For documentation and further use, all isolated strains were deposited in the culture collection of the Institute of Applied Microbiology of the University of Agricultural Sciences (VIAM), Vienna, Aus-

The liquid medium was an aqueous solution of sodium nitrate (3.0 g l31 ), magnesium sulfate (0.5 g l31 ), potassium chloride (0.5 g l31 ), iron(III) sulfate (0.01 g l31 ) and dipotassium hydrogen phosphate (1.0 g l31 ). Sodium nitrate constituted the sole nitrogen source; variable carbon sources were o¡ered by adding pure compounds or root extract fractions. Stock solutions were prepared using this medium containing the test compound or extract at a concentration of 10 mg ml31 and ¢ltered over sterile ¢ltration units (Nalgene). Assays were carried out in sterile, disposable microtitre plates (Greiner; 96 U-bottomed wells). For result comparability, glucose, sucrose and mannitol were assayed together with the water-soluble root extract fractions within the same plate. Stock solutions were serially diluted. Stock inoculum suspension was adjusted to 105 CFU ml31 . Final concentrations ranged from 10 to 5000 Wg ml31 for each compound/extract. Microtitre plates were incubated at room temperature until growth was visible under the microscope (magni¢cation 140U); plates were regularly checked after 24, 48 and 72 h. To stop germ tube growth and enhance contrast for scoring, 10-Wl aliquots of lactophenol blue (Merck) were dispensed into each well. To quantify germ tube growth, a representative section of the well covered with germ tubes was captured to hard disk. Germ tube density was determined by image analysis (Scion Image 3b for Windows [36]) in a representative 150U150-pixel sector of the digitised image. Within the identical microtitre plate, the well showing the highest pixel number was designated as 100%. Pixel numbers obtained from other wells were expressed as percentage of that value, respectively. The 100% value represents also the maximum growth performance of an isolate within a given time period. Replicate experiments were carried out and scored visually to con¢rm previously observed growth behaviour.

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ple Kolmogorov^Smirnov test for normal distribution and Levene’s test of equality of error variances).

2.8. Statistical analysis Statistical analyses were carried out using SPSS 10.0.5 for Windows [37]. Growth rates obtained from di¡erent concentrations in the dilution series represented repeated measurements that may be analysed by one-factor-repeated-measures analysis of variance (ANOVA). However, this type of ANOVA assumes sphericity of variable variances, and present data violate this assumption. Instead, a multivariate analysis of variance (MANOVA) is used that treats the concentration series as multiple independent variables [38]. Assumptions of normal distribution and equality of error variances for variables were ful¢led (one-sam-

3. Results 3.1. Chemical analyses of water-soluble root components Water-soluble root extract fractions of B. erectus, I. ensifolia, L. latifolius and P. alsaticum amounted to at least two thirds of the whole crude MeOH extract. Watersoluble extract fractions were analysed by HPLC^UV and

Table 1 GC^MS analysis of the trimethylsilyl/trimethylsilyl-oxime ethers/esters of organic acids, sugars, sugar alcohols and amino acids from the water-soluble fraction of the methanolic root extract Retention time 5.47 6.65 7.31 7.93 8.08 8.19 8.85 8.96 9.40 11.44 13.16 14.24 15.18 15.74 16.37 17.04 17.21 17.39k 17.54k 18.15 18.17k 18.42k 18.44k 18.62k 19.74 28.65 30.03 31.35k 31.49k 37.34 43.04

B. erectus unknowna unknownb unknownc unknownd homoserinee unknownf malic acide unknowng pyroglutamic acide unknownh xylitole arabinosei citric acide unknownj mannosee mannitoli sorbitole fructosei

I. ensifolia

L. latifolius

5.5

0.5 1.5 0.3 0.4 1.1 0.2 1.0 0.4 0.2 0.5

0.3 tr 0.4

1.4 0.1 6.9

P. alsaticum

P. cervaria 13.5

tr 0.2 0.2 0.3

0.5

37.5

3.3

1.1

22.5

quebrachitole galactosei

0.2

glucosei

7.4

7.0

inositole sucrosei trehalosee maltosee

0.1 72.5 7.5

83.2 0.8

ra⁄nosee maltotriosee

tr 0.7

1.3 19.6 1.7 19.0

2.4 1.0 3.9

28.0

24.8

0.1 85.4

11.0

20.0

0.1 tr

0.2

Relative quantities have been calculated from integrated peak areas of the TIC signal; amounts of single compounds represent percentages of the total area of all peaks detected ; identi¢ed compounds detected in amounts lower than 0.1% are designated as trace (tr) and compounds present in amounts larger than 1% are marked in bold. a MS [m/z(%)] : 70(29), 74(29), 130(31), 203(100), 204(33), 275(89), 292(56). b MS [m/z(%)]: 32(100), 45(38), 56(26), 73(99), 75(92), 103(68), 130(22), 146(79), 248(1). c MS [m/z(%)] : 43(37), 56(100), 73(17), 75(24), 116(5), 130(5). d MS [m/z(%)]: 73(100), 75(47), 77(26), 103(11), 147(34), 212(17), 254(9), 344(6). e Tentative identi¢cation by retention time and MS spectrum comparison with libraries and literature. f MS [m/z(%)] : 43(76), 76(100), 75(64), 77(19), 117(12), 133(14), 152(7), 212(11), 254(22). g MS [m/z(%)] : 43(30), 73(63), 75(34), 100(7), 128(100), 147(10), 188(9), 202(6). h MS [m/z(%)]: 43(47), 73(100), 75(68), 103(23), 147(23), 158(54), 188(21), 229(13), 290(5). i Identi¢cation by comparison of retention times and mass spectra of authentic compounds. j MS [m/z(%)]: 73(100), 75(76), 103(21), 147(25), 217(14), 259(3). k Derivatisation yielded more than one peak in the GC chromatogram.

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GC^MS. UV spectra of peaks present in the HPLC chromatogram indicated presence of glycosides of various natural products, e.g. cinnamic acid derivatives. GC^MS chromatograms, however, suggested that their amounts were negligible in comparison to sugars and sugar alcohols. In selected cases, e.g. for ca¡eic and chlorogenic acid, detectability by GC^MS was additionally con¢rmed by analyses of derivatised authentic compounds. Consequently, quanti¢cation of compounds present in the water-soluble extract fraction was carried out on the basis of results exclusively obtained by GC^MS analysis. Table 1 lists the composition of sugars, sugar alcohols, organic and amino acids that were identi¢ed in the extracts o¡ered

165

in the growth assays. Observed accumulation trends were also con¢rmed by analysis of additional individuals. The two umbellifers, P. alsaticum and P. cervaria, showed different patterns of carbohydrates compared to other investigated plants: mannitol, glucose and fructose were accumulated in similar amounts as sucrose. This disaccharide constituted the main sugar derivative ( s 70%) in the water-soluble carbohydrate fractions of B. erectus, I. ensifolia and L. latifolius. Besides malic acid, the latter plant also contained amino acids in detectable amounts, of which homoserine and pyroglutamic acid were tentatively identi¢ed. MS spectra of all unknowns are listed in the footnotes to Table 1.

Table 2 Microfungal isolates from bulk soil, rhizosphere soil and root tissues of two umbellifers from calcareous grasslands, P. alsaticum L. and P. cervaria (L.) Lapeyr Isolate P. alsaticum Rhizosphere isolates Acremonium strictum W. Gamsa Doratomyces stemonitis (Pers. ex Steud.) Morton and G. Sm.a Fusarium sp.a Penicillium citrinum Thomb Penicillium manginii Duche¤ and Heima P. manginii Duche¤ and Heima Penicillium vulpinum (Cooke and Massee) Seifert and Sampsona Endophytes Cylindrocarpon destructans (Zinssm.) Scholtenb C. destructans (Zinssm.) Scholtenb Penicillium sp.b Penicillium sp.b P. cervaria Rhizosphere isolates Acremonium murorum (Corda) W. Gamsa Arthrinium arundinis Statea Beauveria bassiana (Bals.) Vuill.a C. destructans (Zinssm.) Scholtena Fusarium tricinctum (Corda) Sacc.a Paecilomyces marquandii (Massee) Hughesa Penicillium expansum Link ex Graya Trichoderma koningii Oudem.a Endophytes C. destructans (Zinssm.) Scholtenb C. destructans (Zinssm.) Scholtenb Penicillium sp.b Talaromyces £avus (Klo«cker) Stolk and Samsonb ( = Penicillium vermiculatum Dangeard)b Bulk soil isolates A. murorum (Corda) W. Gamsc Aspergillus ochraceus K. Wilh.c C. destructans (Zinssm.) Scholtena D. stemonitis (Pers. ex Steud.) Morton and G. Sm.a Penicillium corylophilum Dierckxc P. expansum Link ex Grayc Phoma sp.a

Conserved strain

Growth duration (h)

100% growth (pixels)

VIAM-MA2350 VIAM-MA2808 VIAM-MA1084 VIAM-MA2328 VIAM-MA2014 VIAM-MA2016 VIAM-MA2810

24 48 24 24 24 24 24

28 184 24 986 25 641 17 992 11 560 17 262 18 394

VIAM-MA3414 VIAM-MA3415 VIAM-MA3327 VIAM.MA3450

24 24 24 24

13 113 12 151 13 868 3 349

VIAM-MA2817 VIAM-MA2814 VIAM-MA2815 VIAM-MA1059 VIAM-MA2776 VIAM-MA1965 VIAM-MA2811 VIAM-MA1091

48 24 48 48 24 48 24 48

37 124 34 335 34 771 36 011 27 033 28 244 21 412 29 451

VIAM-MA2819 VIAM-MA2825 VIAM-MA2824 VIAM-MA3427

48 48 24 72

30 833 30 893 9 055 42 501

VIAM-MA1937 VIAM-MA1945 VIAM-MA1959 VIAM-MA1987 VIAM-MA1970 VIAM-MA1969 VIAM-MA1981

72 48 24 48 24 24 24

23 417 30 500 15 278 34 455 32 776 22 490 17 699

Strain numbers, assay duration and maximum pixel number determined by image analysis of germ tubes present in a 150U150-pixel section of that well within the microtitre plate that showed maximum growth within the given growth period. All fungal strains have been deposited at the culture collection of the Institute of Applied Microbiology, University of Agricultural Sciences, Vienna (VIAM). a Recovered by soil-washing. b Recovered from surface-sterilised root tissue. c Recovered by soil-plating.

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3.2. Selection of fungal isolates from root tissues, rhizosphere and bulk soil The total number of isolates recovered from the two Peucedanum species were too numerous to be included into this study. Consequently, a number of strains were chosen as representatives of three distinct isolate groups. The three distinguished isolate groups comprised endophytic, rhizosphere soil and bulk soil microfungi (Table 2). Of those identi¢ed, C. destructans was the only species occurring ubiquitously, also within root tissues of both Peucedanum species. Penicillium species were recovered as endophytes, rhizosphere soil and bulk soil fungi. Endophytes of this genus, of which we have been able to identify only one isolate so far, were di¡erent from those species recovered from rhizosphere and bulk soil. Moreover, morphologically, all four isolates seemed to belong to separate species. Apart from identi¢ed isolates, a big number of sterile mycelia also occurred in root tissues. The percentage of such isolates was notably higher among endophytic isolates than rhizosphere soil and bulk soil isolates. However, conidia and/or spores constituted an indispensable prerequisite for germ tube growth assays. Rhizo-

sphere soil contained the greatest diversity of genera and species. Altogether, eight endophytes, 15 rhizosphere soil isolates and seven isolates from bulk soil were chosen for the bioassays. 3.3. Germ tube growth assays to determine utilisation of various carbohydrate sources At ¢rst impression, fungal isolates notably di¡ered in developing variable-sized germ tubes on the various substrates. In general, growth was more vigorous on plantderived carbohydrates than on pure compounds. A few isolates failed to germinate at lower concentrations. MANOVA was carried out to determine if utilisation of a particular substrate corresponds to a⁄liation to a particular isolate group (factor lifestyle) or to association with a particular plant species (factor plant). Table 3 lists the tested substrates by increasing Wilks’ V values of the factor lifestyle. Sucrose and mannitol showed the lowest Wilks’ V values that were also signi¢cant (P 6 0.05). The other substrates only showed non-signi¢cant Wilks’ V values for the same factor. Among those, the substrate with lowest Wilks’ V was glucose, followed by the water-soluble

Table 3 MANOVA to test e¡ects of isolate origin (plant: P. alsaticum, P. cervaria, bulk soil) and isolate lifestyle (endophyte, rhizosphere soil, bulk soil) on growth rates on various substrates Carbon source Sucrose Plant Lifestyle* PlantUlifestyle Mannitol Plant Lifestyle* PlantUlifestyle Glucose Plant Lifestyle PlantUlifestyle P. alsaticum Plant Lifestyle PlantUlifestyle P. cervaria Plant Lifestyle PlantUlifestyle B. erectus Plant Lifestyle PlantUlifestyle I. ensifolia Plant Lifestyle PlantUlifestyle L. latifolius Plant Lifestyle PlantUlifestyle

Wilks’ V

F

Hypoth. df

Error df

P

0.672 0.334 0.412

0.779 3.193 2.281

10 10 10

16 16 16

0.648 0.019 0.068

0.628 0.386 0.548

0.949 2.546 1.319

10 10 10

16 16 16

0.518 0.046 0.300

0.601 0.420 0.547

1.062 2.206 1.322

10 10 10

16 16 16

0.441 0.076 0.298

0.596 0.481 0.539

1.085 1.726 1.367

10 10 10

16 16 16

0.427 0.159 0.279

0.737 0.543 0.509

0.570 1.347 1.541

10 10 10

16 16 16

0.815 0.287 0.213

0.400 0.563 0.787

2.396 1.241 0.434

10 10 10

16 16 16

0.058 0.338 0.908

0.781 0.687 0.531

0.448 0.730 1.415

10 10 10

16 16 16

0.900 0.688 0.258

0.625 0.696 0.664

0.960 0.699 0.811

10 10 10

16 16 16

0.510 0.713 0.623

Plant-derived carbon sources comprise water-soluble constituents of the methanolic root extract. Signi¢cant factors are marked with an asterisk.

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Table 4 Pair-wise comparisons of group means of endophyte (E, n = 8), rhizosphere soil (R, n = 15) and bulk soil (S, n = 7) isolates grown in a dilution series of concentrations on eight di¡erent carbon sources J

5000

E

2500

E

1250

E

625

E

312

E

156

E

78

E

39

E

20

E

10

E

5000

E

2500

E

1250

E

625

E

312

E

156

E

78

E

39

E

20

E

10

E

R S R S R S R S R S R S R S R S R S R S R S R S R S R S R S R S R S R S R S R S

Mean di¡erence I3J glucose

4.83 7.34 2.19 0.93 30.57 30.90 7.30 7.15 2.65 1.66 1.15 0.67 6.92 3.74 12.89* 6.28 11.23 8.49 8.17 4.96 sucrose 4.77 10.21 3.46 5.90 4.05 4.35 13.00 14.17 7.38 10.18 8.00 10.08 14.59* 12.96 13.47* 7.79 7.79 3.06 5.77 1.00

Standard P error 8.56 10.11 7.83 9.25 7.48 8.83 6.90 8.15 6.56 7.75 5.87 6.93 6.12 7.23 5.60 6.61 5.65 6.68 6.12 7.23 8.40 9.92 7.71 9.11 8.02 9.47 7.26 8.58 7.91 9.35 7.17 8.47 6.46 7.63 5.73 6.77 5.57 6.58 4.88 5.77

0.58 0.47 0.78 0.92 0.94 0.92 0.30 0.39 0.69 0.83 0.85 0.92 0.27 0.61 0.03 0.35 0.06 0.22 0.19 0.50 0.58 0.31 0.66 0.52 0.62 0.65 0.09 0.11 0.36 0.29 0.28 0.25 0.03 0.10 0.03 0.26 0.17 0.65 0.25 0.86

mannitol

P. alsaticum

Mean di¡erence I3J

Standard P error

37.41 36.35 11.14 8.67 12.73 14.37 20.00* 14.50 10.12 12.44 8.38 7.96 4.79 5.83 4.04 8.25 4.26 1.65 6.19 7.01 322.84* 317.50* 320.56* 318.11 322.01* 319.55 320.39* 313.06 315.32 312.87 312.07 311.33 34.79 3.26 34.00 2.23 1.16 2.52 33.77 32.87

10.01 11.82 9.52 11.25 8.63 10.19 9.31 11.00 7.75 9.15 7.14 8.43 7.30 8.62 7.22 8.53 6.18 7.30 5.65 6.67 5.68 6.71 8.64 10.21 8.07 9.53 9.16 10.82 10.37 12.25 9.36 11.05 9.60 11.34 8.45 9.98 6.25 7.38 5.22 6.16

0.47 0.60 0.25 0.45 0.15 0.17 0.04 0.20 0.20 0.19 0.25 0.35 0.52 0.51 0.58 0.34 0.50 0.82 0.28 0.30 0.00 0.02 0.03 0.09 0.01 0.05 0.04 0.24 0.15 0.30 0.21 0.32 0.62 0.78 0.64 0.82 0.85 0.74 0.48 0.64

B. erectus

P. cervaria

Mean di¡erence I3J

Standard P error

35.26 30.37 34.39 11.69 7.15 15.84 31.31 12.92 10.36 14.80 9.53 10.38 11.51 11.29 7.29 13.30 15.67* 14.61* 9.91 5.78 316.81 316.36 38.01 36.70 33.35 35.63 36.71 34.15 1.89 31.01 31.87 31.24 1.62 34.85 3.96 0.91 10.18* 8.26 13.11* 5.78

7.58 8.95 7.91 9.34 9.44 11.15 9.49 11.22 9.83 11.61 9.95 11.76 9.30 10.98 9.36 11.05 6.45 7.62 5.62 6.64 8.77 10.36 10.46 12.36 11.83 13.97 10.71 12.65 9.37 11.06 7.83 9.25 7.92 9.35 6.44 7.60 4.79 5.66 4.98 6.64

0.49 0.97 0.58 0.22 0.46 0.17 0.89 0.26 0.30 0.21 0.35 0.39 0.23 0.31 0.44 0.24 0.02 0.07 0.09 0.39 0.07 0.13 0.45 0.59 0.78 0.69 0.54 0.75 0.84 0.93 0.81 0.89 0.84 0.61 0.54 0.91 0.04 0.16 0.01 0.39

I. ensifolia

L. latifolius

Mean di¡erence I3J

Standard error

0.77 31.87 12.19 10.20 31.48 1.74 15.57 17.40 10.28 7.79 6.00 5.31 5.69 31.05 4.07 0.72 4.85 2.92 30.08 3.97 31.88 34.13 39.80 35.04 33.64 31.47 32.15 5.64 4.73 6.85 3.53 8.03 7.98 7.83 3.15 5.10 6.25 8.00 2.86 1.74

210.8 12.78 9.20 10.87 8.29 9.79 9.35 11.05 10.84 12.81 8.97 10.60 9.07 10.72 7.94 9.38 6.87 8.11 6.47 7.64 8.33 9.84 10.25 12.11 8.70 10.27 9.30 10.98 37.79 4.73 9.54 11.27 8.17 9.65 7.74 9.14 6.48 7.66 5.85 6.91

P

0.94 0.88 0.20 0.36 0.86 0.86 0.11 0.13 0.35 0.55 0.51 0.62 0.54 0.92 0.61 0.94 0.49 0.35 0.99 0.61 0.82 0.68 0.35 0.68 0.68 0.89 0.82 0.61 0.72 0.81 0.71 0.48 0.34 0.42 0.69 0.58 0.34 0.31 0.63 0.80

F. Hadacek, G.F. Kraus / FEMS Microbiology Ecology 41 (2002) 161^170

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Concentration I (Wg ml31 )

Di¡erences signi¢cant at the 0.05% level are marked with an asterisk. Data represent the least signi¢cant di¡erence post hoc analysis of the factor lifestyle in the MANOVA shown in Table 3. 167

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carbohydrates of P. alsaticum and P. cervaria, the two rhizosphere hosts. The carbohydrate fractions of B. erectus, I. ensifolia and L. latifolius, which represented those plants growing along with the two focussed umbellifers, showed the highest Wilks’ V and P values. For the other factor, plant, no signi¢cant Wilks’ V was obtained, and values were generally higher. A subsequent pair-wise comparison of least signi¢cant di¡erences of group means for the factor lifestyle aimed to explore the nature of existing group di¡erences (Table 4). On the whole, signi¢cant pair-wise di¡erences of group means were only detected between endophytes on one hand and rhizosphere and bulk soil isolates on the other hand. Endophytes grew signi¢cantly better than rhizosphere soil fungi at 39 and 78 Wg ml31 sucrose, 39 Wg ml31 of glucose, 625 Wg ml31 mannitol and 20 Wg ml31 carbohydrates from B. erectus. Neighbouring concentrations also showed low P values but fell short of the 0.05 signi¢cance level due to large standard errors caused by isolate variability. The water-soluble carbohydrates of P. alsaticum reduced the growth of endophytes in the range of 625^5000 Wg ml31 . The data of P. cervaria suggested a similar e¡ect. However, di¡erences between group means were not signi¢cant. Instead, endophytes grew better at 10 and 20 Wg ml31 . The water-soluble carbohydrates of I. ensifolia and L. latifolius did not a¡ect growth of endophytes discernibly di¡erent compared to rhizosphere soil and bulk soil isolates.

4. Discussion Multivariate analyses of variance and subsequent least signi¢cant di¡erence analyses of group means of fungi recovered from di¡erent rhizosphere habitats suggested that plant sugars or sugar alcohols may constitute signals that facilitate adaptation of certain fungi to a speci¢c host plant. Various results support this interpretation: (1) a⁄liation to a particular isolate group was most notably re£ected by growth patterns on those substrates that contained a single sugar or sugar alcohol (MANOVA) ; (2) carbohydrates of the two host plants, P. alsaticum and P. cervaria, yielded the lower Wilks’ V values with lower P values than non-host plants; and (3) at various concentrations, extracts of host plants changed growth behaviour of endophytes. The two umbellifers, P. alsaticum and P. cervaria, differed from other plants growing around them by accumulating large amounts of fructose, glucose and mannitol in addition to sucrose in their roots (Table 1). However, sucrose was also detected as the major sugar derivative in the roots of B. erectus, I. ensifolia and L. latifolius, despite the fact that these three plant species are classi¢ed in quite heterogeneous plant families. Recent investigations demonstrated that external sugar concentrations may a¡ect gene expression in ectomycorrhizal basidiomycetes [39].

The present results also suggest that sugars or sugar alcohols may constitute important signals to soil fungi. Higher plants are also capable of sensing changing levels of sugars, especially of glucose and sucrose [23,40]. The occurrence of sucrose in the roots of diverse plants might turn e⁄cient utilisation of this monosaccharide into a general constraint for the endophytic lifestyle of a fungus. Low concentrations of sucrose stimulated the growth of endophytes. Raw data suggested a similar e¡ect for glucose. However, signi¢cance closely missed the 0.05% level. Interestingly, the sugar alcohol mannitol also stimulated growth of the endophytic isolates, but only at higher concentrations. High amounts of mannitol were only detected in the two umbellifers, the host plants of the tested endophytic isolates. These ¢ndings suggest that species-speci¢c components of the root carbohydrates, that are accumulated in larger amounts, may also a¡ect endophytes. Mixtures of sugars cause e¡ects that cannot be explained merely by adding the e¡ects of the major single constituents. This becomes evident in the comparison of the variation in the responses of the endophytic isolates to the carbohydrates of the two Peucedanum species that show variable quantities of the same major sugars. Carbohydrates of the dominant grass species, B. erectus, stimulated the growth of the majority of the tested isolates most e⁄ciently of all tested substrates. This concurs with expectations that plant species present in large numbers of individuals may shape the utilisation behaviour of soil microfungi more than those present in lower numbers. The results point to a mechanism of adaptation that may be facilitated by ¢ne-tuning the combination of variable substrate utilisation and/or signal perception capabilities of a single fungal strain. These factors may also participate in the postulated reciprocal determination of plant and fungal diversity. Endophytes constitute an essential proportion of fungal diversity. Among the fungi recovered from the root tissues of the two Peucedanum species, a large number were sterile isolates. A big proportion of these were dark-pigmented mycelia, many more compared to those recovered from rhizosphere and bulk soil. This suggests that endophytes comprise more highly specialised strains than rhizosphere and bulk soil isolates. Our results also corroborate the general impression that rhizosphere soil harbours a higher proportion of fungal diversity than bulk soil [2,3]. However, isolates from both soil habitats did not markedly di¡er in their bandwidth of substrate utilisation. Among those endophytic isolates that we could identify on the basis of morphological characteristics, we found strains belonging to the genera Cylindrocarpon and Penicillium. Cylindrocarpon isolates have frequently been reported as colonisers of plant root surfaces. C. destructans and its teleomorph, Nectria radicicola Gerlach and L. Nilsson, are known to cause root rot, together with various Fusarium species [34]. Other identi¢ed endophytes comprised four Penicillium species. Penicillium represented

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the most species-rich genus among the isolates, especially in the rhizosphere of P. alsaticum and, to a lesser extent, in the bulk soil. The four endophytes were not a⁄liated to any of the soil-derived Penicillium isolates and we have failed to identify three of them so far. Penicillum species have already been reported to occur as endophytes in various trees and shrubs [41^43] and may be more widespread than we currently assume. The biology of endophytic fungi within plants is still not understood completely. Endophytes are considered to be latent pathogens on one hand [44] ; on the other hand, they are also regarded to be mutualists that, by producing toxic secondary metabolites, may even protect their host plants against herbivores or nematodes [45,46]. The majority of anamorphic microfungi in the soil are necrotrophs [34]. The endophytes in our model umbelllifers obviously adapt to their host plants by utilising root carbohydrates more e⁄ciently at lower concentrations. This might be helpful during the infection process. Concurrently, once within the root’s tissue, the growth rate of the endophyte is slowed by higher concentrations of the same compounds. This reduced growth might help to avoid the induction of plant defence mechanisms that are usually triggered by more vigorously growing pathogenic fungi [47]. Later, often many years after infection, the endophytes might pro¢t from the fact that they are able to exploit the dying plant from within its tissues. O¡ering substrates in a range of concentrations o¡ers additional parameters and thus improves the detection of di¡erences in the responses of fungal isolates to a range of di¡erently composed substrates. Usually, growth of a speci¢c fungus or fungal community on a single concentration of de¢ned sugars, sugar alcohols, organic and amino acids after a given period or in a de¢ned time course is compared [13,48]. Although the presented study is of pilot character, conclusions drawn from the obtained results o¡er an interesting working hypothesis for similarly aimed investigations in the future and may also bene¢t research in discovering and understanding gene regulation mechanisms governing plant^fungus interactions. Secondary metabolites have not been the focus of this study, but they certainly are not to be neglected because they may play an essential role in the ¢ne tuning of the species composition within a fungal community. Flavonoids have already been identi¢ed as serving as important stimulant signals to arbuscular mycorrhizal fungi [49].

Acknowledgements This research was supported by Grant P13278-BIO from the Austrian Science Fund. We are grateful to Dr W. Gams (CBS, Utrecht, The Netherlands) and Dr. Katja Ster£inger (Institute of Applied Microbiology, University of Agricultural Sciences, Vienna) for advice on isolate identi¢cation, Prof. K.-H. Prillinger (Institute of Applied

169

Microbiology, University of Agricultural Sciences, Vienna) for general support, and Dr Elvira Ho«randl (Institute of Botany, University of Vienna) for critical comments.

References [1] Whipps, J.M. and Lynch, J.M. (1985) Energy losses by the plant in rhizodeposition. In: Plant Products and the New Technology, Annu. Proc. Phytochem. Soc. Eur. (Fuller, K.W. and Gallon, J.R., Eds.), pp. 59^71. Clarendon Press, Oxford. [2] Brimecombe, M.J., De Leij, F.A. and Lynch, J.M. (2001) The e¡ect of root exudates on rhizosphere microbial communities. In: The Rhizosphere. Biochemistry and Organic Substances at the Soil^Plant Interface (Pinton, R., Varanini, Z. and Nannipieri, P., Eds.), pp. 95^140. Marcel Dekker, New York. [3] Newman, E.I. (1985) The rhizosphere: carbon sources and microbial populations. In: Ecological Interactions in Soil (Fitter, A.H., Ed.), pp. 107^121. Blackwell Scienti¢c, Oxford. [4] Neumann, G. and Ro«mheld, V. (2001) The release of root exudates as a¡ected by the plant’s physiological status. In: The Rhizosphere. Biochemistry and Organic Substances at the Soil^Plant Interface (Pinton, R., Varanini, Z. and Nannipieri, P., Eds.), pp. 41^94. Marcel Dekker, New York. [5] Reid, C.P.P. (1990) Mycorrhizas. In: The Rhizosphere (Lynch, J.M., Ed.), pp. 281^316. John Wiley and Sons, Chichester. [6] Martin, F.M., Perotto, S. and Bonfante, P. (2001) Mycorrhizal fungi: a fungal community at the interface of soil and roots. In: The Rhizosphere. Biochemistry and Organic Substances at the Soil^Plant Interface (Pinton, R., Varanini, Z. and Nannipieri, P., Eds.), pp. 263^296. Marcel Dekker, New York. [7] Petrini, O. (1986) Taxonomy of endophytic fungi in aerial plant tissues. In: Microbiology of the Phyllosphere (Fokkema, N.J. and van den Heuvel, J., Eds.), pp. 175^187. Cambridge University Press, Cambridge. [8] Schulze, E.D., and Mooney, H.A. (1993) Biodiversity and Ecosystem Function. Springer, New York. [9] Hooper, D.U. and Vitousek, P.M. (1997) The in£uence of plant composition and diversity on ecosystem processes. Science 277, 1302^1305. [10] Bever, J.D., Westover, K.M. and Antonovics, J. (1997) Incorporating the soil community into population dynamics: the utility of the feedback approach. J. Ecol. 85, 561^573. [11] Van der Heijden, M.G.A., Klironomos, J.N., Ursic, M., Moutoglis, P., Streitwolf-Engel, R., Boller, T., Wiemken, A. and Sanders, I.R. (1998) Myccorhizal fungal diversity determines plant biodiversity, ecosystem variability and productivity. Nature 396, 69^72. [12] Grayston, S.J., Wang, S., Cambpell, C.D. and Edwards, A.C. (1998) Selective in£uence of plant species on microbial diversity in the rhizosphere. Soil Biol. Biochem. 30, 369^378. [13] Westover, K.M., Kennedy, A.C. and Kelley, S.E. (1997) Patterns of rhizosphere microbial community structure associated with co-occurring plant species. J. Ecol. 85, 863^873. [14] Watkinson, A.R. (1998) The role of the soil community in plant population dynamics. Trends Ecol. Evol. 13, 171^172. [15] Southwood, T.R.E. (1985) Interactions of plants with animals : patterns and processes. Oikos 44, 5^11. [16] Read, D.J. (1997) Mycorrhizal fungi: the ties that bind. Nature 388, 517^518. [17] Prell, H.H. and Day, P.R. (2000) Plant^Fungal Pathogen Interaction. A Classical and Molecular View. Springer Verlag, Berlin. [18] Burdon, J.J. (1993) The structure and pathogen populations in natural plant communities. Annu. Rev. Phytopathol. 31, 305^323. [19] Vandenkoornhuyse, P., Baldauf, S.L., Leyval, C., Stracek, J. and

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170

[20] [21] [22] [23] [24]

[25]

[26]

[27] [28] [29]

[30]

[31]

[32]

[33]

[34] [35]

F. Hadacek, G.F. Kraus / FEMS Microbiology Ecology 41 (2002) 161^170 Young, J.P.W. (2002) Extensive fungal diversity in plant roots. Science 295, 2051. Harborne, J.B. (2000) Arsenal for survival : secondary plant products. Taxon 49, 435^449. Hartmann, T. (1996) Diversity and variability of plant secondary metabolism: a mechanistic view. Entomol. Gen. Appl. 80, 177^188. Harborne, J.B. (1993) Introduction to Ecological Biochemistry, 4th edn. Academic Press, Oxford. Sheen, J., Zhou, L. and Jang, J.-C. (1999) Sugars as signaling molecules. Curr. Opin. Plant Biol. 2, 410^418. Thellung, A. (1925^1926) Peucedanum. In: Ilustrierte Flora von Mitteleuropa. Band V/2 (Hegi, G., Ed.), pp. 1363^1404. J.F. Lehmanns Verlag, Munich. Morse, C.C., Yevdokimov, I.V. and DeLuca, T. (2000) In situ extraction of rhizosphere organic compounds from contrasting plant communities. Commun. Soil Sci. Plant Anal. 31, 725^742. Kantona, Z.F., Sass, P. and Molna¤r-Perl, I. (1999) Simultanous determination of sugars, sugar alcohols, acids and amino acids in apricots by gas chromatography-mass spectrometry. J. Chromatogr. Ser. A 847, 91^102. http://www.mpimp-golm.mpg.de/mms-library/index-e.html. Warcup, J.H. (1950) The soil^plate method. Nature 170, 166^167. Gams, W. and Domsch, K.H. (1967) Beitra«ge zur Anwendung der Bodenwaschtechnik fu«r die Isolierung von Bodenpilzen. Arch. Mikrobiol. 58, 134^144. Gams, W. (1992) The analysis of communities of saprophytic microfungi with special reference to soil fungi. In: Fungi in Vegetation Science (Winterho¡, W., Ed.), pp. 183^224. Kluwer Academic, Dordrecht. Gams, W. and Van Laar, W. (1982) The use of solacol (validamycin) as a growth retardant in the isolation of soil fungi. Neth. J. Plant Pathol. 88, 39^45. Holdenrieder, O. and Sieber, T.N. (1992) Fungal association of serially washed non-mycorrhizal roots of Picea abies. Mycol. Res. 96, 151^156. Schulz, B., Wanke, U., Draeger, S. and Aust, H.J. (1993) Endophytes from herbaceous plant and shrubs: e¡ectiveness of surface sterilisation methods. Mycol. Res. 97, 1447^1450. Gams, W., Domsch, K.H. and Anderson, T.-H. (1980) Compendium of Soil Fungi, Vol. 1. IHW Verlag, Eching. Christensen, M., Frisvad, J.C. and Tuthill, D. (1999) Taxonomy of the Penicillium myczynskii group based on morphology and secondary metabolites. Mycol. Res. 103, 527^541.

[36] http://www.scioncorp.com. [37] Norusis, M.J. (1999) SPSS Advanced Models 9.0. SPSS, Chicago, IL. [38] Turner, J.R. and Thayer, J.F. (2001) Introduction to Analysis of Variance. Sage Publications, Thousand Oaks, CA. [39] Nehls, U., Wiese, J. and Hampp, R. (2000) External sugar concentration as a signal controling ectomycorrhizal fungal gene expression. In: Current Advances in Mycorrhizal Research (Poldila, G. and Douds, D.D., Jr., Eds.), pp. 1^26. APS Press, St. Paul, MN. [40] Coruzzi, G.M. and Zhou, L. (2001) Carbon and nitrogen sensing and signaling in plants: emerging matrix e¡ects. Curr. Opin. Plant Biol. 4, 247^253. [41] Sequerra, J., Capellano, A., Gianinazzi-Pearson, V. and Moiroud, A. (1995) Ultrastructure of cortical root cells of Alnus incana infected by Penicillium nodositatum. New Phytol. 130, 545^555. [42] Collado, J., Platas, G., Gonzalez, I. and Pelaez, F. (2000) Geographical and seasonal in£uences on the distribution of fungal endophytes in Quercus ilex. New Phytol. 144, 525^532. [43] Stierle, A.A. and Stierle, D.B. (2000) Bioactive compounds from four endophytic Penicillium sp. of a Northwest Paci¢c yew tree. In: Studies in Natural Product Chemistry, Vol. 24: Bioactive Natural Products, Part E (Atta-ur-Rahman, Ed.), pp. 933^977. Elsevier, Amsterdam. [44] Sinclair, J.B. and Cerkauskas, R.F. (1996) Latent infection vs. endophytic colonization by fungi. In: Endophytic Fungi in Grasses and Woody Plants: Systematics, Ecology, and Evolution (Redlin, S.C. and Carris, L.M., Eds.), pp. 3^29. APS Press, St. Paul, MN. [45] Clay, K. (1992) Fungal endophytes of plants: biological and chemical diversity. Nat. Toxins 1, 147^149. [46] Hallmann, J. and Sikora, R.A. (1996) Toxicity of fungal endophyte secondary metabolites to plant parasitic nematodes and soil-borne plant pathogenic fungi. Eur. J. Plant Pathol. 102, 155^162. [47] Hammerschmidt, R. and Nicholson, R.L. (1999) A survey of plant defenses to pathogens. In: Induced Plant Defenses against Pathogens and Herbivores (Agrawal, A.A., Tuzun, S. and Bent, E., Eds.), pp. 55^72. APS Press, St. Paul, MN. [48] Buyer, J.S., Roberts, D.P., Millner, P. and Russek-Cohen, E. (2001) Analysis of fungal communities by sole carbon source utilization pro¢les. J. Microbiol. Methods 45, 53^60. [49] Werner, D. (2001) Organic signals between plants and microorganisms. In: The Rhizosphere. Biochemistry and Organic Substances at the Soil^Plant Interface (Pinton, R., Varanini, Z. and Nannipieri, P., Eds.), pp. 197^222. Marcel Dekker, New York.

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