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Use of specific phospholipid fatty acids for identifying and quantifying the external hyphae of the arbuscular mycorrhizal fungus Gigaspora rosea
Soil Biology & Biochemistry 36 (2004) 1827–1834 www.elsevier.com/locate/soilbio
Use of specific phospholipid fatty acids for identifying and quantifying the external hyphae of the arbuscular mycorrhizal fungus Gigaspora rosea K. Sakamoto*, T. Iijima, R. Higuchi Faculty of Horticulture, Chiba University, Matsudo, Chiba 271-8510, Japan Received 28 April 2003; received in revised form 18 February 2004; accepted 15 April 2004
Abstract The external hypha of arbuscular mycorrhizal (AM) fungi, extending from roots out into soil, is an important structure in the uptake of phosphate from the depletion zone around each root. In this paper, we analysed some phospholipid fatty acids (PLFAs) derived from external hyphae of four AM fungi (Glomus etunicatum, Glomus clarum, Gigaspora margarita and Gigaspora rosea) to find fatty acids which may be useful as specific markers for identifying and quantify the external hyphae of Gigaspora species. Leek (Allium porrum L.) seedlings inoculated with each AM fungus were grown in river sand. Sand samples were collected and four PLFAs (16:1u5, 18:1u9, 20:1u9 and 20:4) in the sand were analysed. In addition, the hyphal biomass in the sand was determined by the direct microscopic method. PLFAs 18:1u9 and 20:4 were found in all the AM-inoculated and non-inoculated sand samples. PLFA 16:1u5 was detected in the sand inoculated with G. etunicatum, G. clarum and Gi. rosea. PLFA 20:1u9 was detected only in the sand inoculated with Gi. rosea. PLFAs 16:1u5 and 20:1u9 were not found in the sand inoculated with Gi. margarita. The amount of PLFA 20:1u9 was closely correlated with the amount of biomass of external hyphae of Gi. rosea (rZ0.937, P!0.001), whereas no correlation was observed for PLFA 16:1u5. The 20:1u9 content of Gi. rosea was approximately 6.56 nmol mgK1 hyphal biomass. We suggest that PLFA 20:1u9 can be used as a specific marker for identifying and quantifying the external hyphae of Gi. rosea, at least in controlled experimental systems. q 2004 Elsevier Ltd. All rights reserved. Keywords: Arbuscular mycorrhizal fungi; External hyphae; Gigaspora; Glomus; Phospholipid fatty acid; Specific marker
1. Introduction Arbuscular mycorrhizal (AM) fungi in the order Glomales form obligate associations with up to 80% of vascular plants and show positive effects on plant growth. Plant growth promotion by mycorrhiza is mainly due to the increase in the uptake of phosphate, whereas the plants provide the fungi with carbohydrates (Smith and Read, 1997). The external hypha of AM fungi extending from roots out into soil is an important organ to take up phosphate away from the depletion zone around each root. Sanders et al. (1977) observed that enhancement of plant growth is related * Corresponding author. Tel./fax: C81-47-308-8817. E-mail address: [email protected] (K. Sakamoto). 0038-0717/$ - see front matter q 2004 Elsevier Ltd. All rights reserved. doi:10.1016/j.soilbio.2004.04.037
to the development of external hyphae of AM fungi, and suggested that the capacity of AM fungi to promote plant growth is not a function of the degree to which they colonize the roots. Graham et al. (1982) also found that the extent of plant growth enhancement was proportional to the biomass of external hyphae that had developed as estimated by the weight of soil they had bound into aggregates. Therefore, the development of the soil hyphal network appears to be an important factor determining the capacity of AM fungi to enhance plant growth. However, it is difficult to determine hyphal biomass and distribution of AM fungi in soil. Several methods for determining the external hyphal biomass of AM fungi have been established; most of them are based on microscopy by methods similar to those for soil saprophytic fungi (Hanssen et al., 1974). However, microscopic methods are time-consuming and do not
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allow for separation of different fungal hyphae, or a reliable separation of the dead and live fractions of the fungal biomass (Slyvia, 1992). By using biochemical markers for quantifying fungal biomass, we can overcome some of these drawbacks. The chitin and ergosterol contents have been used as such a marker for fungal hyphae. Each marker, however, has some problems for AM fungi. Chitin persists after the death of the fungus and its suitability in estimating living or recently dead hyphae must be questioned. Furthermore, it is also present in other organisms such as insects and arthropods sometimes leading to high background values in field soil (Slyvia, 1992; Olsson, 1999). AM fungi contain little or no detectable ergosterol (Beilby and Kidby, 1980; Beilby, 1980; Nordby et al., 1981; Schmitz et al., 1991; Frey et al., 1992, 1994; Grandmougin-Ferjani et al., 1999; Fontaine et al., 2001; Olsson et al., 2003). Two important types of lipids found in organisms are phospholipids (membrane constituents) and neutral lipids (energy storage in eukaryotes). These types of lipids contain fatty acids connected to a glycerol backbone. Phospholipid fatty acids (PLFAs) can be used for detecting changes in the microbial community structure in soil since organisms differ in the fatty acids they contain and since the total amount of phospholipids in cells is considered to be well correlated to biomass (Tunlid and White, 1992; Zelles, 1999). Fatty acids derived from phospholipids (located in hyphal cell) and from neutral lipids (located in spores) of AM fungi are potentially useful for detecting or identifying AM fungi (Graham et al., 1995; Bentivenga and Morton, 1996; Jansa et al., 1999; Olsson, 1999). Some fatty acids have been known as specific markers for AM fungi. In the case of Glomus species, fatty acids 16:1u5, 18:1u7, 18:1u9, 20:3, 20:4 and 20:5 have been detected in AM spores and the roots of plants colonized by AM fungi (Beilby and Kidby, 1980; Jabaji-Hare, 1988; Pacovsky and Fuller, 1988; Pacovsky, 1989; Graham et al., 1995; Jansa et al., 1999; Madan et al., 2002). A similar fatty acid composition was found in the spores of Acaulospora laevis by Beilby (1980). In addition, Olsson et al. (1995, 1997, 1998) estimated the biomass of the external hyphae of several Glomus species using the PLFA 16:1u5. In Gigaspora species, fatty acids 18:1u9 and 20:1u9 have been found in their spores as specific markers (Bentivenga and Morton, 1994, 1996; Graham et al., 1995; Madan et al., 2002). Graham et al. (1995) found that the sudan grass roots colonized by Gigaspora rosea contained 20:1u9, but nonmycorrhizal roots did not. However, specific PLFAs present in external hyphae of Gigaspora species are unknown to date. In this study, we analysed some PLFAs derived from the external hyphae of four AM fungi (Glomus etunicatum, Glomus clarum, Gigaspora margarita, and Gi. rosea) to find the fatty acids useful as specific markers for identifying and quantifying the external hyphae of Gigaspora species.
2. Materials and methods 2.1. AM fungal species and host plant Two experiments were carried out with the same growth system. In experiment 1, we used four AM fungal inocula: G. etunicatum Becker and Gerdemann, G. clarum Nicolson and Schenck, Gi. margarita Becker and Hall, and Gi. rosea Nicolson and Schenck. In experiment 2, two AM fungal inocula were used: G. etunicatum and Gi. rosea. The cultures of Gi. margarita and Gi. rosea are registered in the Ministry of Agriculture, Forestry and Fisheries of Japan as MAFF 520052 and MAFF 520062. Seeds of leek (Allium porrum L.) were sterilized in a sodium hypochlorite solution (0.25% available chlorine), rinsed with tap water, and sown in the river sand, autoclaved for 30 min, in plastic pots (250 ml in volume, two plants per pot). Three replicate pots were inoculated with each individual species of AM fungi by addition of 200 spores of the species to each pot prior to sowing of leek seeds. The inoculated and uninoculated plants were cultured in a growth chamber (140 mmol mK2 sK1 fluorescent light, 16 h photoperiod, 25 8C). The liquid fertilizer mentioned below was applied every 4 days, and tap water was supplied properly to wash out the salts. The fertilizer consisted of 20:5:30 (N:P2O5:K2O) PETERS soluble fertilizer (W.R. Grace and Co., Fogelsville, PA, USA) to reach a final phosphorus (P) concentrations of 5.5 mg lK1 (50 mg N lK1) was used for the first week after germination. From the second week, the non-P fertilizer consisted of 20:0:25 PETERS soluble fertilizer with the same nitrogen concentration was used. Plants were harvested 80 days after sowing (DAS) in experiment 1, and 60, 80, and 100 days in experiment 2. 2.2. Root colonization of AM fungi The colonized roots were cut into 1 cm segments, cleared and stained according to the method described by Phillips and Hayman (1970) with trypan blue. After staining, colonized root lengths (%) were estimated using the grid– line intersection method of Giovannetti and Mosse (1980). 2.3. Hyphal biomass in sand sample The hyphal biomass in the sand samples were determined by a direct microscopic method (Sakamoto and Oba, 1994). Sand samples were mixed in sterilized water with a homogenizer (at 18,000 rpm) for 5 min and aliquots of the resulting suspension were filtered through a 0.45 mm Millipore filter. Fungal hyphae were stained on the filter with phenol aniline blue (Hanssen et al., 1974). The length and diameter of the fungal hyphae were measured with an optical microscope at !400 magnification. Hyphal length was estimated by the grid–line intersect method of Nishio (1983). A density of 1.1 g cmK3 and a dry weight of 20%
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were adopted for the fungal biomass calculation (Shields et al., 1973). 2.4. Analysis of phospholipid fatty acids in sand sample The phospholipids were analysed by a modified onephase extraction procedure (Bligh and Dyer, 1959; White et al., 1979). Ten grams (wet wt) of each sand samples were extracted in 27 ml of the one-phase chloroform:methanol (1:2, v/v). After centrifugation, the pellets were washed with 27 ml of the one-phase mixture and 7 ml of water for three times and the supernatants were combined. After splitting the extract into two phases by adding 36 ml each of chloroform and water, the lipid extract was evaporated to about 3 ml and applied to SPE-SI column (Varian, Harbor, USA). The column was eluted with 6 ml of chloroform, 6 ml of acetone and then 6 ml of methanol. Each column eluant was regarded as neutral lipids, glycolipids and phospholipids fraction, respectively. The phospholipids were dried under nitrogen and methyl nonadecanoate (fatty acid methyl ester 19:0) was added as an internal standard. Then 1 ml of boron trifluoride methanol complex solution (Wako Pure Chemical Industries Ltd, Japan) was added and the mixture was heated at 60 8C for 1 h to transform the fatty acids in the phospholipids into free fatty acid methyl esters. These were analysed using a gas chromatography equipped with a flame ionisation detector (Shimazu GC-14B) and a 30 m TC-5 capillary column (GL Sciences Inc., Japan). The fatty acids 16:1u5, 18:1u9, 20:1u9 and 20:4, which were specific markers for AM fungi confirmed by Graham et al. (1995) and Olsson (1999), were identified using the retention times previously determined using GC/MS. Nomenclature of fatty acids follows that used by Tunlid and White (1992). 2.5. Numbers of fungi and bacteria in sand samples Fungal and bacterial colony forming units (CFUs) were counted using the dilution plate technique. Each sand sample (10 g wet wt) was dispersed in a reciprocal shaker with 90 ml sterile water for 10 min. The soil suspensions were then diluted in sterile water and spread on rose bengal streptomycin glucose agar for the fungi or yeast extract agar for the bacteria. Fungal colonies were counted after 5 days at 25 8C and bacterial colonies after 14 days at 30 8C.
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2.6. Statistical analysis Differences in the amounts of hyphal biomass and PLFAs among sand samples were analysed by ANOVA (F-ratio) and if significant differences were found, pairs of samples were compared by Tukey’s test. The relationship between the hyphal biomass and the specific PLFAs was assessed by linear regression. Statistical analysis was performed with SYSTAT ver. 8.0 for Windows (SPSS Inc.).
3. Results In the two experiments, root colonization by AM fungi was not observed in non-inoculated plants. However, about 40–50% of the root length at harvest was colonized in the plant inoculated with G. etunicatum and Gi. margarita and about 70–80% of the root length was colonized in the plant inoculated with G. clarum and Gi. rosea. Table 1 shows the sizes of hyphal biomasses and PLFAs in the sand samples inoculated with the four AM fungi from the first experiment. The hyphal biomass was small (1.05 mg kgK1) in the sand not inoculated with AM fungi (control). Since hyphae resembling AM fungal hyphae were not found in the control, the fungi observed likely are the saprophytic fungi present in the surrounding environment. Hyphal biomass in the AM-inoculated sand was significantly higher than those in the control. The increase of hyphal biomass in the AM-inoculated sand compared with the control may be regarded as the biomass of the external hyphae of AM fungi. PLFAs 18:1u9 and 20:4 were found in all the AMinoculated sand and control samples. The contents of PLFAs 18:1u9 and 20:4 in the control were 115 and 34 nmol kgK1, respectively. The content of 18:1u9 in the AM-inoculated sand was not different from that in the control, whereas the content of 20:4 was higher than that in the control. The presence of PLFAs 18:1u9 in the control may be due to fungal contaminants and the presence of 20:4 may also originate from eukaryote. PLFAs 16:1u5 and 20:1u9 were not detectable in the control, but PLFA 16:1u5 was found in the sand samples inoculated with G. etunicatum, G. clarum and Gi. rosea. This is in accordance with the report of Olsson et al. (1995), who found 16:1u5 in the soil
Table 1 Results from experiment 1. Hyphal biomass and the amounts of various PLFAs in the sand samples inoculated with AM fungi at 80 days after sowing the seed of leek (nZ3). Values in columns followed by the same letter are not significantly different by Tukey’s test (P!0.05). ND, Not detected Hyphal biomass (mg kgK1 dry sand) Control (no inoculum) G. etunicatum G. clarum Gi. margarita Gi. rosea
1.05 a 7.18 b 8.13 b 4.63 ab 8.18 b
PLFAs (nmol kgK1 dry sand) 16:1u5
18:1u9
20:1u9
20:4
ND 45 a 77 a ND 28 a
115 ab 75 a 86 ab 97 ab 194 b
ND ND ND ND 47
34 a 57 a 148 ab 131 ab 255 b
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Fig. 1. Results from experiment 2. The hyphal biomass (mg kgK1 dry sand, nZ3) in the sand samples not inoculated (control) and inoculated with G. etunicatum or Gi. rosea. In the respective periods after sowing, bars with the same letter are not significantly different from Tukey’s test (P!0.05).
inoculated with Glomus species. PLFA 20:1u9 was found only in the sand inoculated with Gi. rosea. These PLFAs likely derived from the external hyphae of inoculated AM fungi. However, PLFAs 16:1u5 and 20:1u9 were not found in the sand inoculated with Gi. margarita. Fig. 1 shows the hyphal biomass in the sand samples inoculated with two AM fungi (G. etunicatum and Gi. rosea) and the control from the second experiment. A small amount of hyphal biomass (!0.4 mg kgK1) was detected in the control. In the sands inoculated with two AM fungi, the hyphal biomass increased with an increase in plant growth. The hyphal biomass in the sand samples inoculated with G. etunicatum and Gi. rosea at 100 DAS were 8.1 and 24.7 mg kgK1, respectively. In the second experiment, the number of fungi in the sand inoculated with AM fungi and that in the control were less than 104 CFU gK1 at any time during the cultivation period and the number of bacteria in the sand inoculated with AM fungi and that in the control were about 1.5–4.5! 106 CFU gK1 during the cultivation period (Table 2). Fig. 2 shows the amounts of PLFAs 16:1u5, 18:1u9, 20:1u9 and 20:4 in the sand samples not inoculated or inoculated with G. etunicatum and Gi. rosea from the second experiment. In the control, PLFA 16:1u5 was not detected at 60 and 80 DAS, but a trace amount of 16:1u5 was detected at 100 DAS. PLFA 16:1u5 was detected in the sand samples inoculated with G. etunicatum and Gi. rosea at
any time during the cultivation period. The amount of 16:1u5 in the sand inoculated with G. etunicatum was approximately equal to that in the sand inoculated with Gi. rosea during the cultivation period, and was the highest at 80 DAS. PLFA 18:1u9 was found in both inoculated sand samples and the control except for the control at 80 DAS. The amount of 18:1u9 was the highest in the sand inoculated with Gi. rosea followed by that in the sand with G. etunicatum and the lowest in the control at any time during the cultivation period. PLFA 20:1u9 was found only in the sand inoculated with Gi. rosea, where it increased with an increase in plant growth and reached 163 nmol kgK1 at 100 DAS. PLFA 20:4 was detected in the sand inoculated with Gi. rosea at 80 DAS and in all the sand samples at 100 DAS. We examined the correlation of the hyphal biomass with the amounts of PLFAs 16:1u5 in the sand samples inoculated with G. etunicatum and Gi. rosea from the second experiment. In both sands inoculated with two AM fungi, no significant correlation was observed between the hyphal biomass and the amount of PLFA 16:1u5 (Fig. 3). However, we found a high positive correlation between the hyphal biomass and the amount of PLFA 20:1u9 in the sand inoculated with Gi. rosea (YZ5.66XC9.86, rZ0.937, P!0.001, Fig. 4).
4. Discussion In this study, we examined the possibility of using specific PLFA to identify and quantify the external hyphae of Gigaspora species. PLFA 20:1u9 was found only in the sand inoculated with Gi. rosea and not in the sands from non-inoculated and inoculated with other AM fungi. Fatty acid 20:1u9 comprised 13.0–16.1% of the total fatty acid profile in the sand inoculated with Gi. rosea (Table 1 and Fig. 2). A highly positive correlation was observed between the biomass of external hyphae determined by the direct microscopic method and the content of 20:1u9 in the sand inoculated with Gi. rosea. This is the first report to show presence of 20:1u9 in the external hyphae of Gi. rosea and the close relationship between the biomass of external hyphae and the content of 20:1u9. PLFA 20:1u9 has been found to date in the spores of Gigaspora albida, Gigaspora decipients, Gigaspora gigantia, Gi. margarita and Gi. rosea (Bentivenga and Morton,
Table 2 Results from experiment 2. The number of fungi and bacteria (log CFU gK1 dry sand, nZ3) in the sand samples not inoculated (control) and that inoculated with G. etunicatum and Gi. rosea Days after sowing
60 80 100
Fungi
Bacteria
Control
G. etunicatum
Gi. rosea
Control
G. etunicatum
Gi. rosea
3.54 3.34 3.19
3.80 3.61 3.39
1.52 3.29 3.67
6.65 6.63 6.49
6.72 6.59 6.19
6.58 6.65 6.63
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Fig. 2. Results from experiment 2. Amounts of PLFAs (nmol kgK1 dry sand, nZ3) in the sand samples not inoculated (control) and inoculated with G. etunicatum or Gi. rosea. DAS, days after sowing. tr., trace amount of 16:1u5 was detected. In the respective PLFAs, bars with the same letter are not significantly different by Tukey’s test (P!0.05).
Fig. 3. Result from experiment 2. Correlation between the hyphal biomass and the amount of PLFA 16:1u5 in the sand inoculated with G. etunicatum (a) or Gi. rosea (b).
1994, 1996; Graham et al., 1995; Madan et al., 2002) and in the sudan grass root colonized by Gi. rosea (Graham et al., 1995). Thus, 20:1u9 seems to be distributed in the spores of various Gigaspora species and in the internal and external hyphae of Gi. rosea. Generally, long chain fatty acids (S20-C) are uncommon in saprophytic fungi (Stahl and Klug, 1996) and bacteria (Tunlid and White, 1992). However, Zelles (1997) reported that 20:1u9 was present in small amounts (!1% of total fatty acid presents) in Rhodococcus ruber (Actinomycetes), Lepista nuda (Basidiomycetes) and Sinapsis alba (Plants). Ruess et al. (2002) also found that some fungalfeeding nematodes contain 20:1u9 (1.3–4.5% of total fatty acid present). As not many soil animals have been investigated for fatty acids yet, likely more will be found to contain 20:1u9. Therefore, the use of PLFA 20:1u9 we propose as a specific marker for identifying and quantifying the external hyphae of Gi. rosea may be limited to wellcontrolled experimental systems like this study. Further information regarding the distribution and biomass of soil organisms which contain 20:1u9 will be needed before we can use PLFA 20:1u9 as a signature for detecting the external hyphae of Gi. rosea in natural soil environments.
In the present study, we detected no PLFA 20:1u9 in the external hyphae of Gi. margarita (Table 1). Fatty acid 20:1u9 has been found in the spores of Gi. margarita (Bentivenga and Morton, 1994, 1996; Graham et al., 1995; Madan et al., 2002), whereas Graham et al. (1995) reported that sudan grass roots colonized by Gi. margarita contained no 20:1u9. Thus, Gi. margarita appears to contain 20:1u9 only in its spores. Graham et al. (1995) stated that the profiles of the fatty acids in the spores of AM fungi were not compatible with those in their mycorrhizal roots. Further investigations need to be carried out to elucidate the presence of 20:1u9 in the external and internal hypha of Gigaspora species other than Gi. rosea and Gi. margarita. In contrast to 20:1u9, no significant correlation was found between the hyphal biomass and the amount of PLFA 16:1u5 in the sand inoculated with either G. etunicatum or Gi. rosea (Fig. 3). This results is not consistent with the reports of Olsson et al. (1995, 1997), who observed a significant correlation between the biomass of external hyphae of Glomus species and the amount of PLFA 16:1u5. In order to examine the difference between 20:1u9 and 16:1u5, we calculated the contents of 20:1u9 and 16:1u5 per unit hyphal biomass of Gi. rosea (Table 3). The 20:1u9
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Fig. 4. Result from experiment 2. Correlation between the hyphal biomass and the amount of PLFA 20:1u9 in the sand inoculated with Gi. rosea.
content of Gi. rosea was 6.56 nmol mgK1 hyphal biomass and was approximately constant, whereas the 16:1u5 content per unit biomass significantly decreased during the cultivation period. This indicates that the aging of AM fungal hyphae resulted in a decrease in PLFA 16:1u5 content per unit hyphal biomass. Biochemical components to be used as signatures for estimating microbial biomass in soil should exist in fairly uniform concentration in the cell (Tunlid and White, 1992). Thus, PLFA 16:1u5 would not be useful as a specific marker for quantifying the external hyphae of AM fungi. Fatty acid 16:1u5 was detected in the spore and the external hyphae of the Glomus species (Jabaji-Hare, 1988; Graham et al., 1995; Olsson et al., 1995; Jansa et al., 1999), and in the spore of A. laevis (Beilby, 1980). It was also found in the spores of Gi. albida, Gi. decipients, Gi. gigantia, Gi. margarita and Gi. rosea (Graham et al., 1995; Bentivenga and Morton, 1996; Madan et al., 2002), and in the external hyphae of Gi. rosea in the present study. Graham et al. (1995) reported that 16:1u5 was not detected in the sudan grass roots colonized by Gi. margarita. The external and internal hyphae of Gi. margarita probably contain no PLFA 16:1u5. Though 16:1u5 was not detected in the non-inoculated sand (control) in this study, it is present in small amounts Table 3 Results from experiment 2. PLFA 20:1u9 and 16:1u5 content per unit hyphal biomass of Gi. rosea (nmol mgK1 dry weight, nZ3). Values in columns followed by the same letter are not significantly different by Tukey’s test (P!0.05) Days after sowing
20:1u9
16:1u5
60 80 100 Average
7.86 b 5.20 a 6.60 ab 6.56
14.77 b 9.33 ab 6.43 a 10.18
(!5% of total fatty acid presents) in a few saprophytic fungi, e.g. Mucor spp. (Jansa et al., 1999; Olsson et al., 1999). It was also found in some bacteria such as Vitreoscilla, Flexibacter (Nichols et al., 1986) and Cytophaga (Walker, 1969) and is proposed as a signature PLFA for Eubacteria (Tunlid and White, 1992). Some different studies revealed that 30–60% of PLFA 16:1u5 in soil was originated from the microorganisms other than AM fungi (Olsson et al., 1995, 1998; Green et al., 1999). This would be another important factor limiting the use of PLFA 16:1u5 as a biomass indicator of AM fungi (Graham et al., 1995; Olsson, 1999). PLFAs 18:1u9 and 20:4 are not recommended as specific markers for identifying the external hyphae of AM fungi, because significant amounts of the two PLFAs were detected in the control (Table 1). Many authors have reported the presence of 18:1u9 in the spore of Glomus, Gigaspora, Scutellospora and Acaulospora species (Graham et al., 1995; Bentivenga and Morton, 1996; Madan et al., 2002), and in the soybean root colonized by Glomus fasciculatum (Pacovsky and Fuller, 1988). Fatty acid 20:4 was also found in the spore of Glomus, Gigaspora, Scutellospora and Acaulospora species (Graham et al., 1995; Bentivenga and Morton, 1996; Madan et al., 2002), in the plant roots colonized by Glomus species (Nordby et al., 1981; Pacovsky and Fuller, 1988; Pacovsky, 1989), and in the external hyphae of two Glomus species (Olsson et al., 1995). However, these two fatty acids were detected in large quantities (20–40% of total fatty acid present) in a range of saprophytic fungi (Stahl and Klug, 1996; Ruess et al., 2002). Fungal contamination may explain the high amounts of PLFAs 18:1u9 and 20:4 in the non-inoculated sand. From pure culture studies it is well established that membrane lipids are greatly affected by environmental factors. Studies of bacterial monocultures have revealed that the concentration and composition of PLFA are affected by growth conditions, such as the temperature, pH, and the nutrient composition of the medium (Tunlid and White, 1992). It has also been reported that the fatty acid composition of fungal cell is susceptible to growth rate, culture age, oxygen availability, temperature, pH, and the composition of the growth medium (Kock and Botha, 1998). The environmental factor that most directly affects fatty acid composition is temperature because of its impact on membrane function (Petersen and Klug, 1994; Zelles, 1997). Petersen and Klug (1994) examined the effect of incubation temperature on the PLFA profile of a soil microbial community and found a decrease in the degree of unsaturation fatty acid and a decrease in the total amount of PLFA with an increase in incubation temperature. However, there is still no information about the significance of variability caused by environmental factors on PLFA profiles of AM fungi. Detailed investigation regarding the impact of environmental factors, such as temperature on the concentration and composition of PLFA of AM fungi need to be carried out in order to identify and quantify AM fungi
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in a natural soil environment using PLFA as a specific marker. We conclude that PLFA 20:1u9 can be used as a specific marker for identifying and quantifying the external hyphae of Gi. rosea in soil, at least in controlled experimental systems. Although Gi. rosea is not widely distributed in various field soils, the use of PLFA 20:1u9 as a specific marker of Gi. rosea would be convenient for elucidating the various ecological and functional features of Gigaspora species as compared with those of Glomus species: such as the distribution and biomass of the external hyphae of Gigaspora species colonizing various kinds of plant, the relationship between the extent of plant growth enhancement and the biomass of external hyphae, the competition for the host plant between Gigaspora and other AM fungal species, and the interaction of Gigaspora species with other soil microorganisms. Further examination regarding the distribution and biomass of the soil organisms that contain 20:1u9 and the effect of environmental factors on the 20:1u9 concentration in Gi. rosea are needed in order to establish PLFA 20:1u9 as a specific marker for identifying and quantifying the external hyphae of Gi. rosea in natural soil environment. Finally, in this study we found no specific PLFA in the external hyphae of Gi. margarita. Graham et al. (1995) also detected no specific PLFA in the internal hyphae of this species. These results suggest that it is difficult to identify and quantify the hyphae of Gi. margarita in the soil and plant roots using specific PLFA.
Acknowledgements We thank Dr Kazuhiro Takagi, National Institute for Agro-Environmental Sciences and Dr Tomoyoshi Murata, National Institute for Environmental Studies for the technical support. We thank Dr Kazuyuki Inubushi for his valuable suggestions. This work was supported in part by a Grant-in-aid for Scientific Research from the Ministry of Education, Culture, Sports, Science and Technology of Japan.
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 and germinated spores of the vesicular–arbuscular mycorrhizal fungus, Glomus caledonius. Lipids 15, 375–378. Bentivenga, S.P., Morton, J.B., 1994. Stability and heritability of fatty acid methyl ester profiles of glomalean endomycorrhizal fungi. Mycological Research 98, 1419–1426. Bentivenga, S.P., Morton, J.B., 1996. Congruence of fatty acid methyl ester profiles and morphological characters of arbuscular mycorrhizal fungi in Gigasporaceae. Proceedings of the National Academy of Sciences of the United States of America 93, 5659–5662.
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Bligh, E.G., Dyer, W.J., 1959. A rapid method of total lipid extraction and purification. Canadian Journal of Biochemistry and Physiology 37, 911–917. Fontaine, J., Grandmougin-Ferjani, A., Hartmann, M., Sancholle, M., 2001. Sterol biosynthesis by the arbuscular mycorrhizal fungus Glomus intraradices. Lipids 36, 1357–1363. Frey, B., Buser, H.-R., Schu¨epp, H., 1992. Identification of ergosterol in vesicular–arbuscular mycorrhizae. Biology and Fertility of Soils 13, 229–234. Frey, B., Vilarin˜o, A., Schu¨epp, H., Arines, J., 1994. Chitin and ergosterol content of extraradical and intraradical mycelium of the vesicular– arbuscular mycorrhizal fungus Glomus intraradices. Soil Biology & Biochemistry 26, 711–717. Giovannetti, M., Mosse, B., 1980. An evaluation of techniques for measuring vesicular–arbuscular mycorrhizal infection in roots. New Phytologist 84, 489–500. Graham, J.H., Linderman, R.G., Menge, J.A., 1982. Development of external hyphae by different isolates of mycorrhizal Glomus spp. in relation to root colonization and growth of troyer citrange. New Phytologist 91, 183–189. Graham, J.H., 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. Grandmougin-Ferjani, A., Dalpe, Y., Hartmann, M., Laruelle, F., Sancholle, M., 1999. Sterol distribution in arbuscular mycorrhizal fungi. Phytochemistry 50, 1027–1031. Green, H., Larsen, J., Olsson, P.A., Jensen, D.F., Jakobsen, I., 1999. Suppression of the biocontrol agent Trichoderma harzianumby mycelium of the arbuscular mycorrhizal fungus Glomus intraradicesin roof-free soil. Applied and Environmental Microbiology 65, 1428–1434. Hanssen, J.F., Thingstad, T.F., Goksøyr, J., 1974. Evaluation of hyphal lengths and fungal biomass in soil by a membrane filter technique. Oikos 25, 102–107. Jabaji-Hare, S., 1988. Lipid and fatty acid profiles of some vesicular– arbuscular mycorrhizal fungi: contribution to taxonomy. Mycologia 80, 622–629. Jansa, J., Gryndler, M., Matucha, M., 1999. Comparison of the lipid profiles of arbuscular mycorrhizal (AM) fungi and soil saprophytic fungi. Symbiosis 26, 247–264. Kock, J.K.L., Botha, A., 1998. Fatty acids in fungal taxonomy, in: Frisvad, J.C., Bridge, P.D., Arora, D.K. (Eds.), Chemical Fungal Taxonomy. Marcel Dekker, New York, pp. 219–246. Madan, R., Pankhurst, C., Hawke, B., Smith, S., 2002. Use of fatty acids for identification of AM fungi and estimation of the biomass of AM spores in soil. Soil Biology & Biochemistry 34, 125–128. Nichols, P.D., Stulp, B.K., Jones, J.G., White, D.C., 1986. Comparison of fatty acid content and DNA homology of the filamentous gliding bacteria Vitreoscilla, Flexibacter and Filibacter. Archives of Microbiology 146, 1–6. Nishio, M., 1983. Direct-count estimation of microbial biomassin soil applied with compost. Biological Agriculture and Horticulture 1, 109–125. Nordby, H.E., Nemec, S., Nagy, S., 1981. Fatty acids and sterols associated with citrus root mycorrhizae. Journal of Agricultural and Food chemistry 29, 396–401. 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 phospholipids and neutral lipid fatty acids to estimate biomass of arbuscular mycorrhizal fungi in soil. Mycological Research 99, 623–629. Olsson, P.A., Ba˚a˚th, E., Jakobsen, I., 1997. Phosphorus effects on the mycelium and storage structures of an arbuscular mycorrhizal fungus as studied in the soil and roots by analysis of fatty acid signatures. Applied and Environmental Microbiology 63, 3531–3538.
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Olsson, P.A., Francis, R., Read, D.J., So¨derstro¨m, B., 1998. Growth of arbuscular mycorrhizal mycelium in calcareous dune sand and its interaction with other soil microorganisms as estimated by measurement of specific fatty acids. Plant and Soil 201, 9–16. Olsson, P.A., Larsson, L., Bago, B., Wallander, H., van Aarle, I.M., 2003. Ergosterol and fatty acids for biomass estimation of mycorrhizal fungi. New Phytologist 159, 7–10. Pacovsky, R.S., 1989. Metabolic differences in Zea–Glomus–Azospirillum symbioses. Soil Biology & Biochemistry 21, 953–960. Pacovsky, R.S., Fuller, G., 1988. Mineral and lipid composition of Glycine–Glomus–Bradyrhizobium symbioses. Physiologia Plantarum 72, 733–746. Petersen, S.O., Klug, M.J., 1994. Effects of sieving, storage, and incubation temperature on the phospholipid fatty acid profile of a soil microbial community. Applied and Environmental Microbiology 60, 2421–2430. Phillips, J.M., Hayman, D.S., 1970. Improved procedures for clearing roots and staining parasitic and vesicular–arbuscular fungi for rapid assessment of infection. Transactions of the British Mycological Society 55, 158–161. Ruess, L., Ha¨ggblom, M.M., Garcı´a Zapata, E.J., Dighton, J., 2002. Fatty acids of fungi and nematodes—possible biomarkers in the soil food chain. Soil Biology & Biochemistry 34, 745–756. Sakamoto, K., Oba, Y., 1994. Effect of fungal to bacterial biomass ratio on the relationship between CO2 evolution and total soil microbial biomass. Biology and Fertility of Soils 17, 39–44. Sanders, F.E., Tinker, P.B., Black, R.L.B., Palmerley, S.M., 1977. The development of endomycorrhizal root systems. I. Spread of infection and growth-promoting effects with four species of vesicular–arbuscular endophyte. New Phytologist 78, 257–268.
Schmitz, O., Danneberg, G., Hundeshagen, B., Klingner, A., Bothe, H., 1991. Quantification of vesicular–arbuscular mycorrhiza by biochemical parameters. Journal of Plant Physiology 139, 106–114. Shields, J.A., Paul, E.A., Lowe, W.E., 1973. Turnover of microbial tissue in soil under field conditions. Soil Biology & Biochemistry 5, 753–764. Slyvia, D.M., 1992. Quantification of external hyphae of vesicular– arbuscular mycorrhizal fungi, in: Norris, J.R., Read, D.J., Varma, A.K. (Eds.), Methods in Microbiology, vol. 24, Techniques for the Study of Mycorrhiza. Academic Press, London, pp. 513–525. Smith, S.E., Read, D.J., 1997. Mycorrhizal Symbiosis. Academic Press, London p. 587. 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. Tunlid, A., White, D.C., 1992. Biochemical analysis of biomass, community structure, nutritional stetus, and metabolic activity of microbial communities in soil, in: Stotzky, G., Bollag, J.-M. (Eds.), Soil Biochemistry, vol. 7. Dekker, New York, pp. 229–262. Walker, R.W., 1969. Cis-11-hexadecanoic acid from Cytophaga hutchinsonii. Lipids 4, 15–18. White, D.C., Davis, W.M., Nickels, J.S., King, J.D., Bobbie, R.J., 1979. Determination of the sedimentary microbial biomass by extractible lipid phosphate. Oecologia 40, 51–62. Zelles, L., 1997. Phospholipid fatty acid profiles in selected members of soil microbial communities. Chemosphere 35, 275–294. Zelles, L., 1999. Fatty acid patterns of phospholipids and lipopolysaccharides in the characterization of microbial communities in soil: a review. Biology and Fertility of Soils 29, 111–129.
Use of specific phospholipid fatty acids for identifying and quantifying the external hyphae of the arbuscular mycorrhizal fungus Gigaspora rosea K. Sakamoto*, T. Iijima, R. Higuchi Faculty of Horticulture, Chiba University, Matsudo, Chiba 271-8510, Japan Received 28 April 2003; received in revised form 18 February 2004; accepted 15 April 2004
Abstract The external hypha of arbuscular mycorrhizal (AM) fungi, extending from roots out into soil, is an important structure in the uptake of phosphate from the depletion zone around each root. In this paper, we analysed some phospholipid fatty acids (PLFAs) derived from external hyphae of four AM fungi (Glomus etunicatum, Glomus clarum, Gigaspora margarita and Gigaspora rosea) to find fatty acids which may be useful as specific markers for identifying and quantify the external hyphae of Gigaspora species. Leek (Allium porrum L.) seedlings inoculated with each AM fungus were grown in river sand. Sand samples were collected and four PLFAs (16:1u5, 18:1u9, 20:1u9 and 20:4) in the sand were analysed. In addition, the hyphal biomass in the sand was determined by the direct microscopic method. PLFAs 18:1u9 and 20:4 were found in all the AM-inoculated and non-inoculated sand samples. PLFA 16:1u5 was detected in the sand inoculated with G. etunicatum, G. clarum and Gi. rosea. PLFA 20:1u9 was detected only in the sand inoculated with Gi. rosea. PLFAs 16:1u5 and 20:1u9 were not found in the sand inoculated with Gi. margarita. The amount of PLFA 20:1u9 was closely correlated with the amount of biomass of external hyphae of Gi. rosea (rZ0.937, P!0.001), whereas no correlation was observed for PLFA 16:1u5. The 20:1u9 content of Gi. rosea was approximately 6.56 nmol mgK1 hyphal biomass. We suggest that PLFA 20:1u9 can be used as a specific marker for identifying and quantifying the external hyphae of Gi. rosea, at least in controlled experimental systems. q 2004 Elsevier Ltd. All rights reserved. Keywords: Arbuscular mycorrhizal fungi; External hyphae; Gigaspora; Glomus; Phospholipid fatty acid; Specific marker
1. Introduction Arbuscular mycorrhizal (AM) fungi in the order Glomales form obligate associations with up to 80% of vascular plants and show positive effects on plant growth. Plant growth promotion by mycorrhiza is mainly due to the increase in the uptake of phosphate, whereas the plants provide the fungi with carbohydrates (Smith and Read, 1997). The external hypha of AM fungi extending from roots out into soil is an important organ to take up phosphate away from the depletion zone around each root. Sanders et al. (1977) observed that enhancement of plant growth is related * Corresponding author. Tel./fax: C81-47-308-8817. E-mail address: [email protected] (K. Sakamoto). 0038-0717/$ - see front matter q 2004 Elsevier Ltd. All rights reserved. doi:10.1016/j.soilbio.2004.04.037
to the development of external hyphae of AM fungi, and suggested that the capacity of AM fungi to promote plant growth is not a function of the degree to which they colonize the roots. Graham et al. (1982) also found that the extent of plant growth enhancement was proportional to the biomass of external hyphae that had developed as estimated by the weight of soil they had bound into aggregates. Therefore, the development of the soil hyphal network appears to be an important factor determining the capacity of AM fungi to enhance plant growth. However, it is difficult to determine hyphal biomass and distribution of AM fungi in soil. Several methods for determining the external hyphal biomass of AM fungi have been established; most of them are based on microscopy by methods similar to those for soil saprophytic fungi (Hanssen et al., 1974). However, microscopic methods are time-consuming and do not
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allow for separation of different fungal hyphae, or a reliable separation of the dead and live fractions of the fungal biomass (Slyvia, 1992). By using biochemical markers for quantifying fungal biomass, we can overcome some of these drawbacks. The chitin and ergosterol contents have been used as such a marker for fungal hyphae. Each marker, however, has some problems for AM fungi. Chitin persists after the death of the fungus and its suitability in estimating living or recently dead hyphae must be questioned. Furthermore, it is also present in other organisms such as insects and arthropods sometimes leading to high background values in field soil (Slyvia, 1992; Olsson, 1999). AM fungi contain little or no detectable ergosterol (Beilby and Kidby, 1980; Beilby, 1980; Nordby et al., 1981; Schmitz et al., 1991; Frey et al., 1992, 1994; Grandmougin-Ferjani et al., 1999; Fontaine et al., 2001; Olsson et al., 2003). Two important types of lipids found in organisms are phospholipids (membrane constituents) and neutral lipids (energy storage in eukaryotes). These types of lipids contain fatty acids connected to a glycerol backbone. Phospholipid fatty acids (PLFAs) can be used for detecting changes in the microbial community structure in soil since organisms differ in the fatty acids they contain and since the total amount of phospholipids in cells is considered to be well correlated to biomass (Tunlid and White, 1992; Zelles, 1999). Fatty acids derived from phospholipids (located in hyphal cell) and from neutral lipids (located in spores) of AM fungi are potentially useful for detecting or identifying AM fungi (Graham et al., 1995; Bentivenga and Morton, 1996; Jansa et al., 1999; Olsson, 1999). Some fatty acids have been known as specific markers for AM fungi. In the case of Glomus species, fatty acids 16:1u5, 18:1u7, 18:1u9, 20:3, 20:4 and 20:5 have been detected in AM spores and the roots of plants colonized by AM fungi (Beilby and Kidby, 1980; Jabaji-Hare, 1988; Pacovsky and Fuller, 1988; Pacovsky, 1989; Graham et al., 1995; Jansa et al., 1999; Madan et al., 2002). A similar fatty acid composition was found in the spores of Acaulospora laevis by Beilby (1980). In addition, Olsson et al. (1995, 1997, 1998) estimated the biomass of the external hyphae of several Glomus species using the PLFA 16:1u5. In Gigaspora species, fatty acids 18:1u9 and 20:1u9 have been found in their spores as specific markers (Bentivenga and Morton, 1994, 1996; Graham et al., 1995; Madan et al., 2002). Graham et al. (1995) found that the sudan grass roots colonized by Gigaspora rosea contained 20:1u9, but nonmycorrhizal roots did not. However, specific PLFAs present in external hyphae of Gigaspora species are unknown to date. In this study, we analysed some PLFAs derived from the external hyphae of four AM fungi (Glomus etunicatum, Glomus clarum, Gigaspora margarita, and Gi. rosea) to find the fatty acids useful as specific markers for identifying and quantifying the external hyphae of Gigaspora species.
2. Materials and methods 2.1. AM fungal species and host plant Two experiments were carried out with the same growth system. In experiment 1, we used four AM fungal inocula: G. etunicatum Becker and Gerdemann, G. clarum Nicolson and Schenck, Gi. margarita Becker and Hall, and Gi. rosea Nicolson and Schenck. In experiment 2, two AM fungal inocula were used: G. etunicatum and Gi. rosea. The cultures of Gi. margarita and Gi. rosea are registered in the Ministry of Agriculture, Forestry and Fisheries of Japan as MAFF 520052 and MAFF 520062. Seeds of leek (Allium porrum L.) were sterilized in a sodium hypochlorite solution (0.25% available chlorine), rinsed with tap water, and sown in the river sand, autoclaved for 30 min, in plastic pots (250 ml in volume, two plants per pot). Three replicate pots were inoculated with each individual species of AM fungi by addition of 200 spores of the species to each pot prior to sowing of leek seeds. The inoculated and uninoculated plants were cultured in a growth chamber (140 mmol mK2 sK1 fluorescent light, 16 h photoperiod, 25 8C). The liquid fertilizer mentioned below was applied every 4 days, and tap water was supplied properly to wash out the salts. The fertilizer consisted of 20:5:30 (N:P2O5:K2O) PETERS soluble fertilizer (W.R. Grace and Co., Fogelsville, PA, USA) to reach a final phosphorus (P) concentrations of 5.5 mg lK1 (50 mg N lK1) was used for the first week after germination. From the second week, the non-P fertilizer consisted of 20:0:25 PETERS soluble fertilizer with the same nitrogen concentration was used. Plants were harvested 80 days after sowing (DAS) in experiment 1, and 60, 80, and 100 days in experiment 2. 2.2. Root colonization of AM fungi The colonized roots were cut into 1 cm segments, cleared and stained according to the method described by Phillips and Hayman (1970) with trypan blue. After staining, colonized root lengths (%) were estimated using the grid– line intersection method of Giovannetti and Mosse (1980). 2.3. Hyphal biomass in sand sample The hyphal biomass in the sand samples were determined by a direct microscopic method (Sakamoto and Oba, 1994). Sand samples were mixed in sterilized water with a homogenizer (at 18,000 rpm) for 5 min and aliquots of the resulting suspension were filtered through a 0.45 mm Millipore filter. Fungal hyphae were stained on the filter with phenol aniline blue (Hanssen et al., 1974). The length and diameter of the fungal hyphae were measured with an optical microscope at !400 magnification. Hyphal length was estimated by the grid–line intersect method of Nishio (1983). A density of 1.1 g cmK3 and a dry weight of 20%
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were adopted for the fungal biomass calculation (Shields et al., 1973). 2.4. Analysis of phospholipid fatty acids in sand sample The phospholipids were analysed by a modified onephase extraction procedure (Bligh and Dyer, 1959; White et al., 1979). Ten grams (wet wt) of each sand samples were extracted in 27 ml of the one-phase chloroform:methanol (1:2, v/v). After centrifugation, the pellets were washed with 27 ml of the one-phase mixture and 7 ml of water for three times and the supernatants were combined. After splitting the extract into two phases by adding 36 ml each of chloroform and water, the lipid extract was evaporated to about 3 ml and applied to SPE-SI column (Varian, Harbor, USA). The column was eluted with 6 ml of chloroform, 6 ml of acetone and then 6 ml of methanol. Each column eluant was regarded as neutral lipids, glycolipids and phospholipids fraction, respectively. The phospholipids were dried under nitrogen and methyl nonadecanoate (fatty acid methyl ester 19:0) was added as an internal standard. Then 1 ml of boron trifluoride methanol complex solution (Wako Pure Chemical Industries Ltd, Japan) was added and the mixture was heated at 60 8C for 1 h to transform the fatty acids in the phospholipids into free fatty acid methyl esters. These were analysed using a gas chromatography equipped with a flame ionisation detector (Shimazu GC-14B) and a 30 m TC-5 capillary column (GL Sciences Inc., Japan). The fatty acids 16:1u5, 18:1u9, 20:1u9 and 20:4, which were specific markers for AM fungi confirmed by Graham et al. (1995) and Olsson (1999), were identified using the retention times previously determined using GC/MS. Nomenclature of fatty acids follows that used by Tunlid and White (1992). 2.5. Numbers of fungi and bacteria in sand samples Fungal and bacterial colony forming units (CFUs) were counted using the dilution plate technique. Each sand sample (10 g wet wt) was dispersed in a reciprocal shaker with 90 ml sterile water for 10 min. The soil suspensions were then diluted in sterile water and spread on rose bengal streptomycin glucose agar for the fungi or yeast extract agar for the bacteria. Fungal colonies were counted after 5 days at 25 8C and bacterial colonies after 14 days at 30 8C.
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2.6. Statistical analysis Differences in the amounts of hyphal biomass and PLFAs among sand samples were analysed by ANOVA (F-ratio) and if significant differences were found, pairs of samples were compared by Tukey’s test. The relationship between the hyphal biomass and the specific PLFAs was assessed by linear regression. Statistical analysis was performed with SYSTAT ver. 8.0 for Windows (SPSS Inc.).
3. Results In the two experiments, root colonization by AM fungi was not observed in non-inoculated plants. However, about 40–50% of the root length at harvest was colonized in the plant inoculated with G. etunicatum and Gi. margarita and about 70–80% of the root length was colonized in the plant inoculated with G. clarum and Gi. rosea. Table 1 shows the sizes of hyphal biomasses and PLFAs in the sand samples inoculated with the four AM fungi from the first experiment. The hyphal biomass was small (1.05 mg kgK1) in the sand not inoculated with AM fungi (control). Since hyphae resembling AM fungal hyphae were not found in the control, the fungi observed likely are the saprophytic fungi present in the surrounding environment. Hyphal biomass in the AM-inoculated sand was significantly higher than those in the control. The increase of hyphal biomass in the AM-inoculated sand compared with the control may be regarded as the biomass of the external hyphae of AM fungi. PLFAs 18:1u9 and 20:4 were found in all the AMinoculated sand and control samples. The contents of PLFAs 18:1u9 and 20:4 in the control were 115 and 34 nmol kgK1, respectively. The content of 18:1u9 in the AM-inoculated sand was not different from that in the control, whereas the content of 20:4 was higher than that in the control. The presence of PLFAs 18:1u9 in the control may be due to fungal contaminants and the presence of 20:4 may also originate from eukaryote. PLFAs 16:1u5 and 20:1u9 were not detectable in the control, but PLFA 16:1u5 was found in the sand samples inoculated with G. etunicatum, G. clarum and Gi. rosea. This is in accordance with the report of Olsson et al. (1995), who found 16:1u5 in the soil
Table 1 Results from experiment 1. Hyphal biomass and the amounts of various PLFAs in the sand samples inoculated with AM fungi at 80 days after sowing the seed of leek (nZ3). Values in columns followed by the same letter are not significantly different by Tukey’s test (P!0.05). ND, Not detected Hyphal biomass (mg kgK1 dry sand) Control (no inoculum) G. etunicatum G. clarum Gi. margarita Gi. rosea
1.05 a 7.18 b 8.13 b 4.63 ab 8.18 b
PLFAs (nmol kgK1 dry sand) 16:1u5
18:1u9
20:1u9
20:4
ND 45 a 77 a ND 28 a
115 ab 75 a 86 ab 97 ab 194 b
ND ND ND ND 47
34 a 57 a 148 ab 131 ab 255 b
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Fig. 1. Results from experiment 2. The hyphal biomass (mg kgK1 dry sand, nZ3) in the sand samples not inoculated (control) and inoculated with G. etunicatum or Gi. rosea. In the respective periods after sowing, bars with the same letter are not significantly different from Tukey’s test (P!0.05).
inoculated with Glomus species. PLFA 20:1u9 was found only in the sand inoculated with Gi. rosea. These PLFAs likely derived from the external hyphae of inoculated AM fungi. However, PLFAs 16:1u5 and 20:1u9 were not found in the sand inoculated with Gi. margarita. Fig. 1 shows the hyphal biomass in the sand samples inoculated with two AM fungi (G. etunicatum and Gi. rosea) and the control from the second experiment. A small amount of hyphal biomass (!0.4 mg kgK1) was detected in the control. In the sands inoculated with two AM fungi, the hyphal biomass increased with an increase in plant growth. The hyphal biomass in the sand samples inoculated with G. etunicatum and Gi. rosea at 100 DAS were 8.1 and 24.7 mg kgK1, respectively. In the second experiment, the number of fungi in the sand inoculated with AM fungi and that in the control were less than 104 CFU gK1 at any time during the cultivation period and the number of bacteria in the sand inoculated with AM fungi and that in the control were about 1.5–4.5! 106 CFU gK1 during the cultivation period (Table 2). Fig. 2 shows the amounts of PLFAs 16:1u5, 18:1u9, 20:1u9 and 20:4 in the sand samples not inoculated or inoculated with G. etunicatum and Gi. rosea from the second experiment. In the control, PLFA 16:1u5 was not detected at 60 and 80 DAS, but a trace amount of 16:1u5 was detected at 100 DAS. PLFA 16:1u5 was detected in the sand samples inoculated with G. etunicatum and Gi. rosea at
any time during the cultivation period. The amount of 16:1u5 in the sand inoculated with G. etunicatum was approximately equal to that in the sand inoculated with Gi. rosea during the cultivation period, and was the highest at 80 DAS. PLFA 18:1u9 was found in both inoculated sand samples and the control except for the control at 80 DAS. The amount of 18:1u9 was the highest in the sand inoculated with Gi. rosea followed by that in the sand with G. etunicatum and the lowest in the control at any time during the cultivation period. PLFA 20:1u9 was found only in the sand inoculated with Gi. rosea, where it increased with an increase in plant growth and reached 163 nmol kgK1 at 100 DAS. PLFA 20:4 was detected in the sand inoculated with Gi. rosea at 80 DAS and in all the sand samples at 100 DAS. We examined the correlation of the hyphal biomass with the amounts of PLFAs 16:1u5 in the sand samples inoculated with G. etunicatum and Gi. rosea from the second experiment. In both sands inoculated with two AM fungi, no significant correlation was observed between the hyphal biomass and the amount of PLFA 16:1u5 (Fig. 3). However, we found a high positive correlation between the hyphal biomass and the amount of PLFA 20:1u9 in the sand inoculated with Gi. rosea (YZ5.66XC9.86, rZ0.937, P!0.001, Fig. 4).
4. Discussion In this study, we examined the possibility of using specific PLFA to identify and quantify the external hyphae of Gigaspora species. PLFA 20:1u9 was found only in the sand inoculated with Gi. rosea and not in the sands from non-inoculated and inoculated with other AM fungi. Fatty acid 20:1u9 comprised 13.0–16.1% of the total fatty acid profile in the sand inoculated with Gi. rosea (Table 1 and Fig. 2). A highly positive correlation was observed between the biomass of external hyphae determined by the direct microscopic method and the content of 20:1u9 in the sand inoculated with Gi. rosea. This is the first report to show presence of 20:1u9 in the external hyphae of Gi. rosea and the close relationship between the biomass of external hyphae and the content of 20:1u9. PLFA 20:1u9 has been found to date in the spores of Gigaspora albida, Gigaspora decipients, Gigaspora gigantia, Gi. margarita and Gi. rosea (Bentivenga and Morton,
Table 2 Results from experiment 2. The number of fungi and bacteria (log CFU gK1 dry sand, nZ3) in the sand samples not inoculated (control) and that inoculated with G. etunicatum and Gi. rosea Days after sowing
60 80 100
Fungi
Bacteria
Control
G. etunicatum
Gi. rosea
Control
G. etunicatum
Gi. rosea
3.54 3.34 3.19
3.80 3.61 3.39
1.52 3.29 3.67
6.65 6.63 6.49
6.72 6.59 6.19
6.58 6.65 6.63
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Fig. 2. Results from experiment 2. Amounts of PLFAs (nmol kgK1 dry sand, nZ3) in the sand samples not inoculated (control) and inoculated with G. etunicatum or Gi. rosea. DAS, days after sowing. tr., trace amount of 16:1u5 was detected. In the respective PLFAs, bars with the same letter are not significantly different by Tukey’s test (P!0.05).
Fig. 3. Result from experiment 2. Correlation between the hyphal biomass and the amount of PLFA 16:1u5 in the sand inoculated with G. etunicatum (a) or Gi. rosea (b).
1994, 1996; Graham et al., 1995; Madan et al., 2002) and in the sudan grass root colonized by Gi. rosea (Graham et al., 1995). Thus, 20:1u9 seems to be distributed in the spores of various Gigaspora species and in the internal and external hyphae of Gi. rosea. Generally, long chain fatty acids (S20-C) are uncommon in saprophytic fungi (Stahl and Klug, 1996) and bacteria (Tunlid and White, 1992). However, Zelles (1997) reported that 20:1u9 was present in small amounts (!1% of total fatty acid presents) in Rhodococcus ruber (Actinomycetes), Lepista nuda (Basidiomycetes) and Sinapsis alba (Plants). Ruess et al. (2002) also found that some fungalfeeding nematodes contain 20:1u9 (1.3–4.5% of total fatty acid present). As not many soil animals have been investigated for fatty acids yet, likely more will be found to contain 20:1u9. Therefore, the use of PLFA 20:1u9 we propose as a specific marker for identifying and quantifying the external hyphae of Gi. rosea may be limited to wellcontrolled experimental systems like this study. Further information regarding the distribution and biomass of soil organisms which contain 20:1u9 will be needed before we can use PLFA 20:1u9 as a signature for detecting the external hyphae of Gi. rosea in natural soil environments.
In the present study, we detected no PLFA 20:1u9 in the external hyphae of Gi. margarita (Table 1). Fatty acid 20:1u9 has been found in the spores of Gi. margarita (Bentivenga and Morton, 1994, 1996; Graham et al., 1995; Madan et al., 2002), whereas Graham et al. (1995) reported that sudan grass roots colonized by Gi. margarita contained no 20:1u9. Thus, Gi. margarita appears to contain 20:1u9 only in its spores. Graham et al. (1995) stated that the profiles of the fatty acids in the spores of AM fungi were not compatible with those in their mycorrhizal roots. Further investigations need to be carried out to elucidate the presence of 20:1u9 in the external and internal hypha of Gigaspora species other than Gi. rosea and Gi. margarita. In contrast to 20:1u9, no significant correlation was found between the hyphal biomass and the amount of PLFA 16:1u5 in the sand inoculated with either G. etunicatum or Gi. rosea (Fig. 3). This results is not consistent with the reports of Olsson et al. (1995, 1997), who observed a significant correlation between the biomass of external hyphae of Glomus species and the amount of PLFA 16:1u5. In order to examine the difference between 20:1u9 and 16:1u5, we calculated the contents of 20:1u9 and 16:1u5 per unit hyphal biomass of Gi. rosea (Table 3). The 20:1u9
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Fig. 4. Result from experiment 2. Correlation between the hyphal biomass and the amount of PLFA 20:1u9 in the sand inoculated with Gi. rosea.
content of Gi. rosea was 6.56 nmol mgK1 hyphal biomass and was approximately constant, whereas the 16:1u5 content per unit biomass significantly decreased during the cultivation period. This indicates that the aging of AM fungal hyphae resulted in a decrease in PLFA 16:1u5 content per unit hyphal biomass. Biochemical components to be used as signatures for estimating microbial biomass in soil should exist in fairly uniform concentration in the cell (Tunlid and White, 1992). Thus, PLFA 16:1u5 would not be useful as a specific marker for quantifying the external hyphae of AM fungi. Fatty acid 16:1u5 was detected in the spore and the external hyphae of the Glomus species (Jabaji-Hare, 1988; Graham et al., 1995; Olsson et al., 1995; Jansa et al., 1999), and in the spore of A. laevis (Beilby, 1980). It was also found in the spores of Gi. albida, Gi. decipients, Gi. gigantia, Gi. margarita and Gi. rosea (Graham et al., 1995; Bentivenga and Morton, 1996; Madan et al., 2002), and in the external hyphae of Gi. rosea in the present study. Graham et al. (1995) reported that 16:1u5 was not detected in the sudan grass roots colonized by Gi. margarita. The external and internal hyphae of Gi. margarita probably contain no PLFA 16:1u5. Though 16:1u5 was not detected in the non-inoculated sand (control) in this study, it is present in small amounts Table 3 Results from experiment 2. PLFA 20:1u9 and 16:1u5 content per unit hyphal biomass of Gi. rosea (nmol mgK1 dry weight, nZ3). Values in columns followed by the same letter are not significantly different by Tukey’s test (P!0.05) Days after sowing
20:1u9
16:1u5
60 80 100 Average
7.86 b 5.20 a 6.60 ab 6.56
14.77 b 9.33 ab 6.43 a 10.18
(!5% of total fatty acid presents) in a few saprophytic fungi, e.g. Mucor spp. (Jansa et al., 1999; Olsson et al., 1999). It was also found in some bacteria such as Vitreoscilla, Flexibacter (Nichols et al., 1986) and Cytophaga (Walker, 1969) and is proposed as a signature PLFA for Eubacteria (Tunlid and White, 1992). Some different studies revealed that 30–60% of PLFA 16:1u5 in soil was originated from the microorganisms other than AM fungi (Olsson et al., 1995, 1998; Green et al., 1999). This would be another important factor limiting the use of PLFA 16:1u5 as a biomass indicator of AM fungi (Graham et al., 1995; Olsson, 1999). PLFAs 18:1u9 and 20:4 are not recommended as specific markers for identifying the external hyphae of AM fungi, because significant amounts of the two PLFAs were detected in the control (Table 1). Many authors have reported the presence of 18:1u9 in the spore of Glomus, Gigaspora, Scutellospora and Acaulospora species (Graham et al., 1995; Bentivenga and Morton, 1996; Madan et al., 2002), and in the soybean root colonized by Glomus fasciculatum (Pacovsky and Fuller, 1988). Fatty acid 20:4 was also found in the spore of Glomus, Gigaspora, Scutellospora and Acaulospora species (Graham et al., 1995; Bentivenga and Morton, 1996; Madan et al., 2002), in the plant roots colonized by Glomus species (Nordby et al., 1981; Pacovsky and Fuller, 1988; Pacovsky, 1989), and in the external hyphae of two Glomus species (Olsson et al., 1995). However, these two fatty acids were detected in large quantities (20–40% of total fatty acid present) in a range of saprophytic fungi (Stahl and Klug, 1996; Ruess et al., 2002). Fungal contamination may explain the high amounts of PLFAs 18:1u9 and 20:4 in the non-inoculated sand. From pure culture studies it is well established that membrane lipids are greatly affected by environmental factors. Studies of bacterial monocultures have revealed that the concentration and composition of PLFA are affected by growth conditions, such as the temperature, pH, and the nutrient composition of the medium (Tunlid and White, 1992). It has also been reported that the fatty acid composition of fungal cell is susceptible to growth rate, culture age, oxygen availability, temperature, pH, and the composition of the growth medium (Kock and Botha, 1998). The environmental factor that most directly affects fatty acid composition is temperature because of its impact on membrane function (Petersen and Klug, 1994; Zelles, 1997). Petersen and Klug (1994) examined the effect of incubation temperature on the PLFA profile of a soil microbial community and found a decrease in the degree of unsaturation fatty acid and a decrease in the total amount of PLFA with an increase in incubation temperature. However, there is still no information about the significance of variability caused by environmental factors on PLFA profiles of AM fungi. Detailed investigation regarding the impact of environmental factors, such as temperature on the concentration and composition of PLFA of AM fungi need to be carried out in order to identify and quantify AM fungi
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in a natural soil environment using PLFA as a specific marker. We conclude that PLFA 20:1u9 can be used as a specific marker for identifying and quantifying the external hyphae of Gi. rosea in soil, at least in controlled experimental systems. Although Gi. rosea is not widely distributed in various field soils, the use of PLFA 20:1u9 as a specific marker of Gi. rosea would be convenient for elucidating the various ecological and functional features of Gigaspora species as compared with those of Glomus species: such as the distribution and biomass of the external hyphae of Gigaspora species colonizing various kinds of plant, the relationship between the extent of plant growth enhancement and the biomass of external hyphae, the competition for the host plant between Gigaspora and other AM fungal species, and the interaction of Gigaspora species with other soil microorganisms. Further examination regarding the distribution and biomass of the soil organisms that contain 20:1u9 and the effect of environmental factors on the 20:1u9 concentration in Gi. rosea are needed in order to establish PLFA 20:1u9 as a specific marker for identifying and quantifying the external hyphae of Gi. rosea in natural soil environment. Finally, in this study we found no specific PLFA in the external hyphae of Gi. margarita. Graham et al. (1995) also detected no specific PLFA in the internal hyphae of this species. These results suggest that it is difficult to identify and quantify the hyphae of Gi. margarita in the soil and plant roots using specific PLFA.
Acknowledgements We thank Dr Kazuhiro Takagi, National Institute for Agro-Environmental Sciences and Dr Tomoyoshi Murata, National Institute for Environmental Studies for the technical support. We thank Dr Kazuyuki Inubushi for his valuable suggestions. This work was supported in part by a Grant-in-aid for Scientific Research from the Ministry of Education, Culture, Sports, Science and Technology of Japan.
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