The use of fatty acid signatures to study mycelial interactions between the arbuscular mycorrhizal fungus Glomus intraradices and the saprotrophic fungus Fusarium culmorum in root-free soil

Mycol. Res. 102 (12) : 1491–1496 (1998)

1491

Printed in the United Kingdom

The use of fatty acid signatures to study mycelial interactions between the arbuscular mycorrhizal fungus Glomus intraradices and the saprotrophic fungus Fusarium culmorum in root-free soil J O H N L A R S E N1*, P AH L A X E L O L S S O N2 A N D I V E R J A K O B S E N1 " Plant Biology and Biogeochemistry Department, Risø National Laboratory, DK-4000 Roskilde, Denmark # Department of Microbial Ecology, Lund University, Ecology Building, S-223 62 Lund, Sweden

The saprotrophic fungus Fusarium culmorum, Penicillium hordei, Rhizoctonia solani and Trichoderma harzianum and the arbuscular mycorrhizal fungus Glomus intraradices were examined for content of phospholipid fatty acids (PLFA) and neutral lipid fatty acids (NLFA). The AM fungus differed from the saprotrophic fungi especially by its content of the fatty acid 16 : 1ω5 which was absent in the saprotrophs. The fatty acid 18 : 2ω6,9 was the dominant fatty acid of the saprotrophic fungi while it was negligible in mycelium of G. intraradices. Specificity in content of fatty acids made it possible to quantify G. intraradices. and F. culmorum simultaneously in soil. Furthermore, a compartmented growth system made it possible to study mycelial interactions in the absence of roots. We measured hyphal spread of both fungi, hyphal $$P transport of G. intraradices and sporulation of F. culmorum. The two fungi did not interact according to the parameters used in this study. We conclude that fatty acid signatures may be a valuable tool when studying interactions between AM fungi and other fungi in root-free soil.

Arbuscular mycorrhizal (AM) fungi transport P from the soil to their host plant via an external hyphal network, and this P transport can result in enhanced plant growth (Jakobsen, Joner & Larsen, 1994). Little is known, however, about the interactions between the external mycelium of AM fungi and other micro-organisms which could potentially affect P uptake by the AM fungi (Fitter & Garbaye, 1994). To our knowledge no attempt has been made to study mycelial interactions in soil, which may be related to the lack of appropriate tools to measure fungal biomass of different fungi in soil simultaneously. Microscopic examination of hyphae collected on membrane filters is widely used for the quantification of AM hyphae in soil (Sylvia, 1992), but it is generally accepted that this method has a number of limitations (Dodd, 1994). It is rather difficult to distinguish AM hyphae from hyphae of other fungi and hyphal counts are, therefore, most often corrected for counts obtained from non-mycorrhizal controls. Experienced researchers may be able to distinguish AM hyphae from hyphae of other fungi (Miller & Jastrow, 1990), but the use of specific chemical signatures would provide a more objective quantification of AM fungi. Chemical constitutents like chitin (Bethlenfalway & Ames, 1987), ergosterol (Nylund & Wallander, 1992), and fatty acids (Frostega/ rd & Ba/ a/ th, 1996) have been used to quantify fungi in soil. Among these chemical markers, however, only fatty acids provide the

* Present address of corresponding author : Department of Plant Pathology and Pest Management, Danish Institute of Agricultural Sciences, Ministry of Food, Agriculture and Fisheries, Flakkebjerg, DK-4200 Slagelse, Denmark.

possibility to quantify different fungi co-existing in the same soil as fungi are known to differ in their fatty acid profile (Mu$ ller et al., 1994). The fatty acid 18 : 2ω6,9 is dominating in most fungi (Mu$ ller et al., 1994) but not in AM fungi (Johansen, Finlay & Olsson, 1996). AM fungi on the other hand contain the fatty acid 16 : 1ω5 (Beilby, 1980 ; Beilby & Kidby, 1980) which is absent in most other fungi. This specificity in fatty acid profiles made it possible to quantify AM fungi in soil (Olsson et al., 1995) and these specific fatty acids may prove valuable for distinguishing AM fungi and saprotrophic fungi in soil. In addition to the hyphal growth interactions it is also important to develop methods for studying the fungal interactions with respect to their nutrient uptake. Radioactive and stable isotopes are widely used to study nutrient transport by AM fungal mycelium (Jacobsen et al., 1992 ; Johansen, Jakobsen & Jensen, 1993) and these methods are also valuable for studying effects of saprotrophic fungi on AM fungal nutrient transport. The objective of this work was to determine whether fatty acid signatures could be used to measure hyphal interactions between an AM fungus and a saprotrophic fungus. MATERIALS AND METHODS Experiment 1 Fungi. Fusarium culmorum W. G. Sm. (isolate 119-95-2-7, Dr S. Elmholt), Penicillium hordei Stolk (isolate K18, Dr S. Elmholt), Rhizoctonia solani J. G. Ku$ hn (AG5, Dr D. Funck Jensen) and Trichoderma harzianum Rifai (isolate T3, Dr D. Funch Jensen)

Mycelial interactions in root-free soil were maintained on potato dextrose agar at room temperature while Glomus intraradices N. C. Schenck & G. S. Sm. (isolate 28A, Dr S. Rosendahl) was grown in symbiosis with Trifolium subterraneum L. in a compartmented growth system with a root-free sand compartment as described by Johansen et al. (1996). Production of mycelium for fatty acid analysis. Mycelium of the saprotrophic fungi was produced on potato dextrose agar covered by a cellophane sheet. The mycelia were harvested from the cellophane sheets after 4 d. Mycelium of G. intraradices was extracted from sand from a root-free sand compartment (units 4 wk old) of an AM pot culture as described by Johansen et al. (1996). Freeze-dried mycelium (5 mg) of the different fungi was homogenized by placing two steel mill balls in teflon test tubes containing the mycelium and shaking the test tubes in a rotary shaker for 15 min. The homogenate of each fungus was subjected to lipid extraction according to the method used by Frostega/ rd, Tunlid & Ba/ a/ th (1991). The extracted lipids were fractionated on silicic acid columns (see Olsson et al., 1995) and fatty acid residue phospholipids and neutral lipids were transformed into free fatty acid methyl esters by a mild alkaline methanolysis (Dowling, Widdel & White, 1986). Finally, the free fatty acid methyl esters were analysed on a Hewlett Packard 5890 gas chromatograph with a flame ionization detector and a 50 m HP5 capillary column (Frostega/ rd, Tunlid & Ba/ a/ th, 1993). Identification was carried out using the relative retention times compared to an internal standard on a chromatogram with PLFAs previously determined using GC\MS. Nomenclature of fatty acids follows Tunlid & White (1992). Experiment 2 Experimental design. Glomus intraradices was grown in symbiosis with Cucumis sativus L. in a compartmented growth system with or without the saprotrophic fungus F. culmorum. The growth system was made from PVC tubes (4n5-cm internal diam.) and consisted of a central root compartment (RC) (32n5 cm) separated from two 7-cm root-free soil compartments (RFSC) by means of a 37 µm nylon mesh (Larsen & Jakobsen, 1996). F. culmorum was added to the soil at the distal end of the RFSC which was accessible to the external mycelium of the AM fungus. Non-AM treatments with or without F. culmorum were also included. Four replicate units of each main treatment were harvested after 25, 35 and 45 d. Soil. The soil, which was a 1 : 1 (w\w) mixture of sandy loam and quartz sand containing 8 mg kg−" soil of 0n5  NaHCO $ extractable P (Olsen et al., 1954) and with pH (H O) of 6n1, # was partially sterilized by irradiation (10 kGy, 10 MeV electron beam). The following nutrients were mixed into the soil (mg kg−" soil) : NH NO (86), KH PO (44), K SO (70), % $ # % # % CaCl (70), CuSO i5H O (2n2), ZnSO i7H O (5), # % # % # MnSO i7H O (10), CoSO i7H O (0n33), % # % # NaMoO i2H O (0n2) and MgSO i7H O (20). Ten ml of % # % # an aqueous solution containing 14n4 MBq H $$PO was added $ % to 360 g of quartz sand, allowed to dry and mixed with 3240 g

1492 soil. This resulted in a radioactivity level of 4n0 kBq g−" soil. Each root compartment was filled with 300 g soil in the bottom, a mixture of 100 g inoculum (crude inoculum of G. intraradices containing soil, roots and spores obtained from T. subterraneum pot cultures) and 200 g soil adjacent to the two RFSC and 140 g soil as a top layer. Non-AM treatments received no inoculum. One of the RFSC of each unit was filled with 125 g soil. The other RFSC of each unit was filled with 25 g soil, then 75 g $$P labelled soil and finally 25 g soil as the outermost layer. The soil in each unit was watered to 60 % of field capacity and incubated for 4 d at room temperature. In order to establish similar initial microflora communities in all treatments all units received 10 ml of a soil suspension obtained by wet-sieving (20 µm nylon mesh) of 100 g of inoculum in 1 l water. Ten days after seedling emergence, 2 g of F. culmorum inoculum (sterile wheatbran\sawdust 1 : 1, w\w) : water, 2 : 1 ; w\w was applied to the soil surface in both RFSC in half of both AM and non-AM treatments. Plants and growth conditions. Two pre-germinated seeds of Cucumis sativus (Aminex F1 hybrid) were sown in each RC and thinned to one after seedling emergence. Plants were maintained in a growth chamber with a 16\8 h light\dark cycle at 21\16 mC and Osram daylight lamps provided a photosynthetic active radiation of 500–550 µmol m−# s−". Initially the growth units were randomly arranged and thereafter rearranged daily so that each growth unit had a new position in the growth chamber every day. Each growth unit was watered daily by weight to maintain 60 % of the field capacity. Nitrogen was supplied weekly as a NH NO % $ solution with a total of 125 mg N per plant during the growth period. Harvest and analyses. At each harvest the shoot was dried (24 h at 80m) and weighed. The unlabelled RFSC was dismantled and the soil core was pushed out and the spread of each fungus was directly observed using a stereo microscope. The soil core was divided into five 1 cm sections and stored in a freezer (k18m) until further analysis. The roots in the RC were washed, dried (48 h at 80m) and weighed. A subsample was cleared and stained for measurement of mycorrhiza formation according to Kormanick & McGraw (1982) except that we used trypan blue instead of acid fuchsin. Dried plant materials (shoot and root) were ground and digested in a solution of nitric\perchloric acid (4 : 1, v\v). Three ml of the diluted digest was mixed with 15 ml scintillation fluid and analysed on a Packard TR1900 liquid scintillation counter for content of $$P. At the last harvest (45 d), numbers of F. culmorum conidia in the 4–5 cm soil zone in the RFSC were measured by a membrane filter technique (Jakobsen et al., 1992). Three grams of soil from each of the soil zones in the RFSC of all treatments were subjected to lipid extraction and subsequently the fatty acid content was determined as described for the pure mycelium in experiment 1. The fatty acids 16 : 1ω5 and 18 : 2ω6,9 was used as biomass indicator for G. intraradices and F. culmorum, respectively. Statistics. Levels of significance of main treatments and their interactions were calculated by analysis of variance after

J. Larson, P. A. Olsson and I. Jakobsen

1493

testing for normality and variance homogeneity. Means were compared by LSD n . ! !& RESULTS Experiment 1 Fungal fatty acid profiles. Common fungal fatty acids, both PLFAs and NLFAs, were detected in the pure mycelia of all fungi (Table 1). The fatty acids 16 : 1ω5, 20 : 4 and 20 : 5 were found in G. intraradices mycelium but not in the saprotrophic

fungi. In contrast, fatty acid 18 : 2ω6,9 was present in high amounts in the mycelium of saprotrophic fungi, but only in negligible amounts in G. intraradices. The fatty acids 17 : 1ω8 and 17 : 0 were found only in mycelium of P. hordei. All fungi contained much higher concentration of NLFAs than that of PLFAs. Experiment 2 Plant growth and mycorrhiza formation. Plant dry weights and mycorrhiza formation were unaffected by the presence of

Table 1. Phospholipid (PLFA) and neutral lipid (NLFA) fatty acid content in mycelium of different fungi 15 : 0

16 : 1ω7

16 : 1ω5

16 : 0

17 : 1ω8

17 : 0

18 : 2ω6,9

18 : 1ω7

18 : 1ω9

18 : 0

20 : 4

20 : 5

(nmol mg−" .. mycelium) PLFA Glomus intraradices Fusarium culmorum Trichoderma harzianum Penicillium hordei Rhizoctonia solani NLFA Glomus intraradices Fusarium culmorum Trichoderma harzianum Penicillium hordei Rhizoctonia solani

0n01 0n02 0n03 0n03 0n05

0n06 0n18 0n04 0n04 0n08

0n15 — — — —

0n33 2n32 0n59 0n43 0n47

— — — 0n01 —

— — — 0n02 —

0n02 6n71 1n03 0n76 0n75

0n26 — — — 0n05

0n05 1n57 0n36 0n22 0n26

0n01 0n21 0n06 0n07 0n04

0n02 — — — —

0n05 — — — —

0n04 1n01 0n94 1n91 0n48

0n06 8n17 1n22 3n13 0n81

4n80 — — — —

2n92 147n5 33n86 36n87 4n29

— — — 1n05 —

— — — 0n89 —

0n07 144n9 59n9 42n7 7n07

0n43 — — — —

0n38 252n3 43n19 32n05 3n21

0n09 35n8 3n91 4n85 0n30

0n01 — — — —

0n06 — — — —

Table 2. Content of PLFAs 16 : 1ω5 and 18 : 2ω6,9 in the root-free soil compartment of mycorrhizal and non-mycorrhizal units at different distances from the rooting compartment (0–1, 1–2, 2–3, 3–4 and 4–5 cm) with and without Fusarium culmorum inoculated wheatbran\sawdust adjacent to the outermost zone PLFA 16 : 1ω5 (nmol g−" soil)

Treatment AM

FC

k k j j

k j k j

0n35 0n34 0n74 0n80

0n27 0n28 0n93 0n90

k

0n35

0n31

0n01 0n81 0n79

0n001 0n93 0n86

LSD

!n!&

AM FC AMiFC

0–1

1–2

2–3 0n28 0n29 1n02 0n74

PLFA 18 : 2ω6,9 (nmol g−" soil) 3–4 0n29 0n48 0n89 0n85

4–5 0n28 1n30 1n03 2n86

0–1

1–2

2–3

3–4

0n14 0n12 0n20 0n16

0n10 0n11 0n23 0n18

0n10 0n13 0n24 0n13

0n18 1n58 0n17 0n59

0n2 16n27 0n18 11n91

ns

1n27

7n39

0n14 0n41 0n18

0n25 0n05 0n26

0n39 0n001 0n39

0n23 0n32 1n18 ns 0n12 P values obtained from two-way analysis of variance 0n001 0n001 0n01 0n31 0n03 0n10 0n49 0n01 0n65 0n53 0n86 0n27 0n32 0n84 0n46

4–5

Table 3. Content of NLFAs 16 : 1ω5 and 18 : 2ω6,9 in the root-free soil compartment of mycorrhizal and non-mycorrhizal units at different distances from the rooting compartment (0–1, 1–2, 2–3, 3–4 and 4–5 cm) with and without Fusarium culmorum inoculated wheatbran\sawdust adjacent to the outermost zone NLFA 16 : 1ω5 (nmol g−" soil)

Treatment

NLFA 18 : 2ω6,9 (nmol g−" soil)

AM

FC

0–1

1–2

2–3

3–4

4–5

k k j j

k j k j

0n41 0n34 35n13 115n25

0n29 0n33 120n95 187n25

0n48 0n82 287n85 117n51

0n41 0n44 107n72 89n55

0n45 0n48 139n01 141n57

k

30n02

55n24

LSD

!n!&

AM FC AMiFC

0n001 0n01 0n001

0n001 0n09 0n09

0–1

235n20 28n49 118n85 P values obtained from two-way analysis 0n02 0n001 0n01 0n29 0n35 0n97 0n29 0n34 0n97

1–2

2–3

3–4

4–5

0n29 0n62 1n17 1n93

0n34 0n79 2n01 2n47

0n65 0n87 5n12 1n56

0n77 2n29 1n84 2n40

1n03 9n88 2n11 8n20

0n46 of variance 0n001 0n01 0n17

0n43

ns

1n15

4n97

0n001 0n01 0n95

0n13 0n31 0n25

0n1 0n02 0n22

0n86 0n01 0n41

Mycelial interactions in root-free soil F. culmorum in the RFSC. The average plant dry weight of both non-AM and AM plants were 5n3, 8n5 and 12n5 g, after 25, 35 and 45 d, respectively. Plants which received no AM inoculum remained non-mycorrhizal while plants which received AM inoculum had 33, 50 and 65 % of their root length colonised after 25, 35 and 45 d, respectively. Visual and microscopic observation on spread of fungi in the root-free soil compartment. Direct observations of hyphal spread of the two fungi was made at the two first harvests. Mycelium of G. intraradices had spread 3 cm and 5 cm from the RC into the RFSC after 25 and 35 d, respectively. AM fungal spores were observed after 35 d in the 4–5 cm zone irrespective of the presence of F. culmorum. Mycelium of F. culmorum had spread 2–3 cm from the wheatbran\sawdust inoculum into the RFSC after 25 d and no further spread was found after 35 d. When both fungi were present in the RFSC it was only possible to follow F. culmorum at the first harvest when the mycelia did not overlap. The numbers of F. culmorum conidia in the 4–5 cm zone in the RFSC were 4n7i10( (.. l 0n7i10() and 5n1i10( (.. l 1n8i10() g−" soil in the absence and presence of G. intraradices, respectively. Conidia were also observed but not counted in soil from the 3–4 cm zone. In the 0–3 cm zone no conidia were found. Content of 16 : 1ω5 (PLFA and NLFA) in the root-free soil compartment. Soil from RFSC in AM units with or without F. culmorum contained significantly higher amounts of both PLFA and NLFA 16 : 1ω5 than measured in the corresponding non-AM units (Tables 2 and 3). Content of both PLFA and NLFA 16 : 1ω5 were in the same range in the different soil sections of each treatment except for an increase in the PLFA 16 : 1ω5 content in soil from the 4–5 cm soil section in both AM and non-AM treatments with F. culmorum inoculated wheatbran\sawdust. The F. culmorum inoculation had no effect on the content of NLFA 16 : 1ω5 except for an increase in soil from the 0–1 cm section in the mycorrhizal treatment only (Table 3). Content of 18 : 2ω6,9 (PLFA and NLFA) in the root-free soil compartment. Soil from section 3–4 cm and 4–5 cm of both non-AM and AM units treated with F. culmorum inoculum contained significantly more PLFA 18 : 2ω6,9 than that of the other three sections in the 0–3 cm zone and more than that of the corresponding zones without F. culmorum inoculum (Table 2). The content of PLFA 18 : 2ω6,9 in RFSC soil were unaffected by the AM treatment except for soil in the 1–2 cm section where the content of PLFA 18 :2ω6,9 were higher in the AM treatment than in the non-AM treatment. Soil from section 4–5 cm of both non-AM and AM units treated with F. culmorum inoculum contained more NLFA 18 : 2ω6,9 than that of the other four sections in the 0–4 cm zone and more than that of the corresponding zones without F. culmorum inoculum (Table 3). In general, RFSC soil from AM treatments contained more NLFA 18 : 2ω6,9 than that of soil from the corresponding non-AM treatment irrespective of the F. culmorum inoculation. Similarly, RFSC soil in treatment with F. culmorum had a higher content of NLFA 18 : 2ω6,9

1494 Table 4. Total uptake of $$P (i10% cpm) of Glomus intraradices at intervals after seedling emergence as affected by Fusarium culmorum. 25 d

35 d

45 d

Without F. culmorum With F. culmorum

0n4 0n4

55n3 76n9

138n3 143n7

ANOVA (P values)

0n90

0n11

0n72

than that of RFSC soil from treatments without F. culmorum irrespective of mycorrhiza inoculation. Uptake of 33P from the root-free soil compartment. NonAM plants did not contain any $$P at any of the three harvests. The content of $$P in AM plants increased with time but were unaffected by the presence of F. culmorum (Table 4). DISCUSSION Previous studies on interactions between mycelia of AM fungi and saprotrophic fungi have been carried out in dual compartment petri dish systems based on cultures of mycorrhizal roots which permit mycelial interactions to be directly observed (St Arnaud et al., 1995). The present work is the first to quantify co-existing mycelia in soil of an AM fungus and a specific species of another fungus by means of fatty acid signatures. The two fungi seemed to influence each other only to a limited extent under the conditions imposed in this experiment, but we cannot outrule possible interaction at other inoculum densities. Fatty acid signatures, however, appear to be a promising tool for future studies on interaction between AM fungi and other fungi. The fatty acid composition of a range of saprotrophic fungi and of G. intraradices suggest that specific fatty acid profiles are a valuable tool to differentiate mycelium of G. intraradices and saprotrophic fungi in root-free soil. Our results that the fatty acid 18 : 2ω6,9 was dominant among the saprotrophic fungi is consistent with other reports (Mu$ ller et al., 1994 ; Stahl & Klug, 1996). Accordingly, the fatty acid composition of G. intraradices was similar to that of other Glomus species (Graham, Hodge & Morton, 1995 ; Olsson et al., 1995) as well as to another isolate of G. intraradices (Johansen et al., 1996). Specific fatty acid profiles should enable us to distinguish between mycelia of AM fungi and of saprotrophic fungi coexisting in soil. The fact that the fatty acid 18 : 2ω6,9 is, however, present in most fungi makes it important to consider the interference from other fungi in our estimation of F. culmorum biomass on the basis of this fatty acid. The inoculum material for F. culmorum might have stimulated the growth of other fungi as well as G. intraradices. From microscopic observations of the F. culmorum inoculum at the end of Experiment 2, however, no other sporulating fungi were observed but the presence of non-sporulating fungi or other Fusarium looking fungi growing from the substrate cannot be ruled out. Consequently, we cannot eliminate the possibility that saprotrophic fungi other than F. culmorum might have been contributing to the content of the fatty acid 18 : 2ω6,9 in the soil. Indeed, fungi other than F. culmorum seem to be responsible for the increase in the NLFA 18 : 2ω6,9 content in

J. Larson, P. A. Olsson and I. Jakobsen soil from the 1–2 and 2–3 cm soil section in non-AM treatments as direct observations, numbers of conidia and PLFA measurements indicates that the F. culmorum mycelium was restricted to the 3–5 cm soil section. The relatively high PLFA 16 : 1ω5 background values might also be a matter of concern but these have also been found in another report (Olsson et al., 1995) and is probably due to the presence of bacteria containing this fatty acid (Walker 1969 ; Nichols et al., 1986). The increase in the PLFA 16 : 1ω5 soil content in non-AM treatments in the 4–5 cm soil section is most likely caused by the presence of bacteria containing this PLFA which responds positively to the presence of the wheatbran\sawdust-based inocolum. This is supported by the absence of a similar response in NLFA 16 : 1ω5 as the content of NLFA in bacteria in general are low (Niedhardt, Ingraham & Schaechter, 1990). The increase in the NLFA 16 : 1ω5 content in the 0–1 cm soil section as a response to F. culmorum inoculation could be related to sporulation as NLFA is mainly found in storage structures like spores and vesicles (Cooper & Lo$ sel, 1978). We did not measure the numbers of spores in the soil but Olsson, Ba/ a/ th & Jakobsen (1997) found a strong correlation between numbers of G. caledonium spores and content of NLFA 16 : 1ω5. F. culmorum mycelium was, however, only present in the 3–5 cm soil section, indicating that an interaction taking place in one part of the mycelium could result in a response in another part of the mycelium. The content of NLFA in pure mycelium of F. culmorum was much higher than that of PLFA, while in the soil F. culmorum contained about the same amount of both NLFA and PLFA. This seems to indicate that high amounts of neutral lipids are not stored in F. culmorum mycelium in soil while AM fungi usually contain high amounts of neutral lipids also in soil (Olsson et al., 1995, 1997). F. culmorum and G. intraradices mycelium did not compete for $$P under the conditions imposed in this experiment. Since F. culmorum mycelium only grew 1 cm into the 3 cm $$P labelled soil strong competition was not, however, expected. In addition, G. intraradices biomass was unaffected by the presence of F. culmorum supporting the lack of competition. In the present experiments we found that the fatty acids 16 : 1ω5 and 18 : 2ω6,9 can be used as biomass indicators of G. intraradices and other fungi, respectively. These biomass indicators revealed that interactions between mycelium of the AM fungus G. intraradices and the saprotrophic fungus F. culmorum in root-free soil are limited at the inoculum densities tested. Overall, fatty acid signatures seems to be a valuable tool in studies on mycelial interactions between AM fungi and saprotrophic fungi in root-free soil.

We wish to thank Anette Olsen (Risø) for excellent technical assistance and Dr Søren Rosendahl (University of Copenhagen, Denmark), Dr Susanne Elmholt (Danish Institute of Agricultural Sciences, Denmark) and Dr Dan Funck Jensen (Royal Veterinary and fungal Agricultural University, Denmark) for providing us with isolates. Furthermore, Dr Morten Reerslev (University of Copenhagen, Denmark) is thanked for advice concerning production of pure mycelium for fatty acid

1495 analysis. This work was supported by a grant from the Danish Veterinary and Agricultural Research Council. REFERENCES Beilby, J. P. (1980). Fatty acid and sterol composition of ungerminated spores of the vesicular-arbuscular mycorrhizal fungus, Acaulospora laevis. Lipids 15, 949–952. Beilby, J. P. & Kidby, D. K. (1980). Sterol composition of ungerminated spores of the vesicular-arbuscular mycorrhizal fungus, Glomus caledonium. Lipids 15, 375–378. Bethlenfalway, G. J. & Ames, R. N. (1987). Comparison of two methods for quantifying external mycelium of vesicular-arbuscular mycorrhiza. Soil Science Society of America Journal 51, 834–837. Cooper, K. M. & Lo$ sel, D. M. (1978). Lipid physiology of vesicular-arbuscular mycorrhiza. I. Composition of lipids in roots of onion, clover and ryegrass infected with Glomus mosseae. New Phytologist 80, 143–151. Dodd, J. (1994). Approaches to the study of the external mycelium of arbuscular mycorrhizal fungi. In Impact of Arbuscular Mycorrhizas on Sustainable Agriculture and Natural Ecosystems (ed. S Gianinazzi & H. Schu$ epp), pp. 147–166. BirkHa$ user Verlag : Berlin. Dowling, N. J. E., Widdel, F. & White, D. C. (1986). Phospholipid ester-linked fatty acid biomarkers of acetate oxidizing sulphate reducers and other sulphide-forming bacteria. Journal of General Microbiology 132, 1815–1825. Fitter, A. H. & Garbaye, J. (1994). Interactions between mycorrhizal fungi and other soil organisms. Plant and Soil 159, 123–132. Frostega/ rd, AH . & Ba/ a/ th, E. (1996). The use of phospholipid fatty acid analysis to estimate bacterial and fungal biomass in soil. Biology and Fertility of Soils 22, 59–65. Frostega/ rd, AH . Tunlid, A. & Ba/ a/ th, E. (1991). Microbial biomass measured as total lipid phosphate in soils of different organic content. Journal of Microbiological Methods 14, 156–163. Frostega/ rd, AH ., Tunlid, A. & Ba/ a/ th, E. (1993). Phospholipid fatty acid composition, biomass, and activity of microbial communities from two soil types experimentally exposed to different heavy metals. Applied and Environmental Microbiology 59, 3605–3617. Graham, H. J., Hodge, N. C. & Morton, J. B. (1995). Fatty acid methyl ester profiles for characterization of Glomalean fungi and their endomycorrhizae. Applied and Environmental Microbiology 61, 58–64. 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 120, 371–380. Jakobsen, I., Joner, E. & Larsen, J. (1994). Hyphal phosphorous transport, a keystone to mycorrhizal enhancement of plant growth. In Impact of Arbuscular Mycorrhizas on Sustainable Agriculture and Natural Ecosystems (ed. S. Gianinazzi & H. Schu$ epp), pp. 147–166. BirkHa$ user Verlag : Berlin. Johansen, A., Jakobsen, I. & Jensen, E. S. (1993). External hyphae of vesiculararbuscular mycorrhizal fungi associated with Trifolium subterraneum L. 3. Hyphal transport of $#P and "&N. New Phytologist 124, 61–68. Johansen, A., Finlay, R. D. & Olsson, P. A. (1996). Nitrogen metabolism of external hyphae of the arbuscular mycorrhizal fungus Glomus intraradices. New Phytologist, 133, 705–712. Kormanick, P. P. & McGraw, A. C. (1982). Quantification of vesiculararbuscular mycorrhiza in plant roots. In Methods and Principles of Mycorrhizal Research (ed. N. C. Schenck), pp. 37–45. American Phytopathological Society : St. Paul, MN, U.S.A. Larsen, J. & Jakobsen, I. (1996). Interactions between a mycophagous Collembola, dry yeast and an arbuscular mycorrhizal fungus. Mycorrhiza 6, 259–264. Miller, R. M. & Jastrow, J. D. (1990). Hierarchy of root and mycorrhizal fungal interactions with soil aggregation. Soil Biology and Biochemistry 22, 579–584. Mu$ ller, M. M., Kantola, R. & Kitunen, V. (1994). Combining sterol and fatty acid profiles for the characterization of fungi. Mycological Research 98, 593–603. Nichols, P. D., Stulp, B. K., Jones, J. G. & White, D. G. (1986). Comparison of fatty acid content and DNA homology of the filamentous gliding bacteria Vitreoscilla, Flexibacter and Filibacter. Archives of Microbiology 141, 1–6.

Mycelial interactions in root-free soil Niedhardt, F. C., Ingraham, J. L. & Schaetcher, M. (1990). Physiology of the Bacterial Cell. A Molecular Approach. Sinauer Associates : Sunderland, Massachusetts, U.S.A. Nylund, J. E. & Wallander, H. (1992). Ergosterol analysis as a mean of quantifying mycorrhizal biomass. Methods in Microbiology 24, 77–88. Olsen, S. R., Cole, C. V., Watanabe, F. S. & Dean, L. P. (1954). Estimation of available phosphorous in soils by extraction with sodium bicarbonate. Circular No 939, United States Department of Agriculture. Olsson, P. A., Ba/ a/ th, E. & Jakobsen, I. (1997). Distribution of an arbuscular mycorrhizal fungus between soil and roots, and between mycelial and storage structures as response to P application studied by fatty acid signatures. Applied and Environmental Microbiology (in press). Olsson, P. A., Ba/ a/ th, E., Jakobsen, I. & So$ derstro$ m, B. (1995). The use of phospholipid and neutral lipid fatty acids to estimate biomass of arbuscular mycorrhizal fungi in soil. Mycological Research 99, 623–629. (Accepted 9 February 1998)

1496 St Arnaud, M., Hamel, C., Vimard, B., Caron, M. & Fortin, J. A. (1995). Altered growth of Fusarium oxysporum f. sp. chrysanthemi in an in vitro dual culture system with the vesicular arbuscular mycorrhizal fungus Glomus intraradices growing on Daucus carota transformed roots. Mycorrhiza 5, 431–438. Stahl, P. D. & Klug, M. J. (1996). Characterization and differentiation of filamentous fungi based on fatty acid composition. Applied and Environmental Microbiology 62, 4136–4146. Sylvia, D. M. (1992). Quantification of external hyphae of vesicular-arbuscular mycorrhizal fungi. Methods in Microbiology 24, 53–65. Tunlid, A. & White, D. C. (1992). Biochemical analysis of biomass, community structure, nutritional status, and metabolic activity of microbial communities in soil. In Soil Biochemistry, Vol. 7 (ed. G. Stotzky & J.-M. Bollag), pp. 229– 262. Marcel Dekker : New York. Walker, R. W. (1969). Cis-1-hexadecanoic acid from Cytophaga hutchisionii lipids. Lipids 4, 15–18.