Ergosterol and microbial biomass in the rhizosphere of grassland
soils
Rainer Georg Joergensen*
Department of Soil Biology, University of Kassel, Nordbahnhofstr. 1a, 37213 Witzenhausen, Germany
Accepted 2 October 1999
Abstract
Fungal and microbial biomass were determined by ergosterol and fumigation±extraction, respectively, in bulk grassland soil (<2 mm), rhizosphere and root material. The aim was to quantify the contribution of these three microbial fractions to the total soil microbial biomass and to soil organic matter. In the bulk soil, the average concentration was 3.37 mg gÿ1 for ergosterol, 860mg gÿ1for microbial biomass C, and 30.4 mg gÿ1for organic C. In the rhizosphere material, the corresponding
concentrations exceeded those of the bulk soil by 80, 80 and 50%, respectively. The large average ergosterol concentration of 74.2 mg gÿ1root revealed a strong fungal colonisation of the root material. About 75% of the total ergosterol was found in the bulk soil fraction, 11% in the rhizosphere and 14% in the root material. In one soil nearly half of total ergosterol was found in the root material. However, the average ergosterol-to-biomass ratio in the root material was less than a third of the bulk soil or the rhizosphere soil, indicating that approximately two-thirds of CHCl3-labile C are presumably root-derived. 72000 Elsevier
Science Ltd. All rights reserved.
Keywords:Ergosterol; Fungal biomass; Microbial biomass C; Rhizosphere; Fumigation extraction; CHCl3-labile root material
1. Introduction
The development and use of new methods more than 20 yr ago for quantifying microbial biomass in soil such as fumigation±incubation (Jenkinson and Powlson, 1976) and substrate-induced respiration (Anderson and Domsch, 1978) was one of the major advances in soil microbial ecology. The next important step was the development of the fumigation±extraction method, giving the possibility of measuring microbial biomass in the presence of actively decomposing sub-strates (Vance et al., 1987; Ocio and Brookes, 1990). However, the fumigation±extraction method suers from the problem that results are aected by the pre-sence of intact living roots, leading to overestimations of the actual microbial biomass (Mueller et al., 1992).
It was shown repeatedly that organic carbon is ren-dered extractable by CHCl3 fumigation (Martin and
Foster, 1985; Sparling et al., 1985). Consequently, soil samples freshly taken in the ®eld need a pre-treatment before the microbial biomass can be measured. Soils must be sieved to remove roots or preincubated to condition the micro¯ora (Jenkinson, 1988). However, this pre-treatment is always a point of critical discus-sion, especially if the microbial biomass is measured in ®eld experiments (Joergensen et al., 1994; Ohlfs and Scherer, 1996).
An unknown amount of microbial biomass in the rhizosphere soil attached to the roots remains on the sieve, i.e. soil material with much higher microbial bio-mass and carbon availability than the bulk soil (Nor-ton et al., 1990; Cheng et al., 1996). For this reason, the risk of underestimating microbial biomass is es-pecially large in densely rooted grassland soils. Never-theless, it is possible to detect at least the fungal part of the microbial biomass by measuring the
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brane component ergosterol, which is a speci®c bio-marker for fungi. In contrast to the methods men-tioned above, ergosterol is not aected by the presence of roots and can be easily detected by a variety of methods in solid substrates (West et al., 1987; Padgett and Posey, 1993; Stahl and Parkin, 1996). For these reasons, ergosterol has been successfully used to dier-entiate between fungal and plant tissue (Seitz et al., 1977; Newell, 1992, 1996; Nylund and Wallander, 1992; Wallander et al., 1997). However, the conversion of ergosterol data to fungal biomass is hampered by the large variability of the ergosterol concentration in dierent fungal species as reviewed by Djajakirana et al. (1996). The conversion of carbon rendered extracta-ble by CHCl3 fumigation is less variable (Joergensen,
1996).
Consequently, fumigation±extraction and ergosterol data were combined in the present study in an attempt to quantify the contribution of rhizosphere microor-ganisms to the total soil microbial biomass. For this purpose, soils were sampled at several grassland sites and divided into the sample fractions bulk soil (<2 mm), rhizosphere and root material by combining ordinary sieving of ®eld moist soil with the wet-sieving procedure of Mueller et al. (1992).
2. Materials and methods
2.1. Soil sampling and preparation
Soils were sampled on 13 Mondays from October to January in periods without frost at grassland sites around GoÈttingen, south lower Saxony, Germany. Four 125-cm3 cores were sampled at each site for the determination of bulk density. Samples of between 4 and 5 kg were taken to a depth of 10 cm using a spade. The samples were transferred to the laboratory and divided into four parts. First of all, each quarter was weighed and then it was sieved (2 mm) to remove roots. The sieved fraction of the sample was called the bulk soil. The soil attached to the roots, called rhizo-sphere material, was washed away on a 2 mm sieve and collected in a bucket. The soil slurry in the bucket was transferred in 500-ml polyethylene bottles and cen-trifuged for 20 min at 500gto separate rhizosphere soil and water. Then, subsamples were taken from the wet rhizosphere material for fumigation±extraction and ergosterol measurement. The remaining rhizosphere material was dried at 1058C for the determination of dry weight. From the cleaned and thus soil-free root material, also moist subsamples for fumigation±extrac-tion and ergosterol were taken and the remaining root material was dried at 1058C for the determination of dry weight and then ashed at 7008C for the determi-nation of organic matter. All samples were analysed
separately within 4 d for ergosterol and microbial bio-mass.
2.2. Analysis
Microbial biomass C was measured by fumigation extraction (Vance et al., 1987). Moist bulk soil (20 g) was extracted for 45 min by oscillating shaking at 250 rev minÿ1with 80 ml 10 mM CaCl2for the fumigated
and for the nonfumigated treatment and ®ltered through a folded paper ®lter (Schleicher & Schuell, 595 1/2). Moist rhizosphere material (4 g) was extracted for 45 min by oscillating shaking at 250 rev minÿ1 with 16 ml 10 mM CaCl2 for the fumigated and for
the nonfumigated treatment and centrifuged at 2000g. The organic C concentration in ®ltered and centrifuged extracts is not signi®cantly dierent. However, ®l-tration is less laborious and centrifugation gives larger amounts of extract which is important using small samples sizes. Moist root material (2 g) was extracted for 45 min by oscillating shaking at 250 rev minÿ1 with 30 ml 10 mM CaCl2 for the fumigated and for
the nonfumigated treatment and ®ltered through a folded paper ®lter (Schleicher & Schuell, 595 1/2). Or-ganic C in the extracts was measured by catalytic oven oxidation at 8008C using a Maihak Tocor 2 automatic analyser. Soil microbial biomass C was estimated from the relationship: biomass CEC=kEC, whereEC is
(or-ganic C extracted from fumigated soil) minus (or(or-ganic C extracted from nonfumigated soil) and kEC 0:45
(Joergensen, 1995). Ergosterol was measured according to Djajakirana et al. (1996). Moist soil of 1 g dry weight was extracted with 100 ml ethanol for 30 min by oscillating shaking at 250 rev minÿ1. Ergosterol was measured by reversed-phase HPLC analysis at 258C using a column of 12.5 cm Spherisorb ODS II S5 with a mobile phase of 97 vol% methanol/3 vol% water and detection at 282 nm.
Subsamples of dried bulk soil and rhizosphere ma-terial were homogenised in a ball mill. Total C was determined after dry combustion at 12008C using an elemental analyser (Carlo Erba). Organic C was measured as total C minus carbonate C which was measured gas-volumetrically after the addition of 4 M HCl. Organic C in the root material was from the for-mula: 0.5555 ash-free organic matter (Joergensen, 1991). Ash concentration of root material was gravi-metrically determined after combustion of a 2 g air-dried subsample at 7008C for 5 h. Soil pH was measured in water using a soil-to-water ratio of 1-to-2.5. Clay content was determined by sedimentation after pre-treatment with H2O2, Na-citrate and
Na2S2O4 to remove organic matter and iron oxides
means and are given on an oven-dry basis (1058C, 24 h).
3. Results
Soil pH ranged from 4.3 to 7.9, bulk density from 0.99 to 1.54 kg dmÿ3and clay content from 8 to 27% re¯ecting the dierent parent materials (Table 1). The sieved bulk soil (<2 mm) represented on average 91.9% of the total sample dry weight
sum of bulk soil, rhizosphere and root material), the
rhizosphere material 7.4% and the root material only 0.7%, ranging from 0.3 to 2.1%. The average concen-trations were 0.86 mg gÿ1 for microbial biomass C (Table 2), 3.37mg gÿ1for ergosterol (Table 3) and 30.4 mg gÿ1for organic C (Table 2) in the bulk soil. There, microbial biomass C showed signi®cant correlations (P < 0.01) with the ergosterol r0:80 and organic C
r0:82). In the rhizosphere material, the correspond-ing concentrations exceeded those of the bulk soil by 80, 80 and 50%, respectively. This means that the bio-mass C-to-organic C ratios were usually larger in the rhizosphere material than in the bulk soil (Table 2) and that the ergosterol-to-biomass C ratios were almost identical in these two sample fractions (Table 3). In the root material, the average ergosterol concen-tration was 74.2 mg gÿ1 dry weight, but the average ergosterol-to-biomass C ratio was only 0.13%, less than a third of that in the bulk soil or in the rhizo-sphere material (Table 3). The ergosterol-to-biomass C ratios of bulk soil and rhizosphere material were highly signi®cantly correlated r0:83,P< 0.01)
indi-cating a stable relationship between these two mi-crobial indices within a soil. About 75% of total ergosterol was found in the bulk soil fraction, 11% in the rhizosphere and 14% in the root material (Table 3). In soil No. 7 nearly half of the total sum was found in the root material, exceeding the value of the bulk soil.
4. Discussion
The large ergosterol concentrations in the soil-free root material revealed a strong fungal colonisation. Similar concentrations of 52 and 72 mg ergosterol gÿ1 dry material were determined in the roots of Zea mays and Trifolium alexandrinum, respectively, infected with the vesicular±arbuscular endomycorrhizal fungus Glo-mus intraradices after 80 d of growth (Frey et al., 1992). In the bulk soil, the concentrations of ergosterol and microbial biomass C are in the range of those reported for grassland soils, i.e. relatively low in com-parison to that of arable or forest soils (Djajakirana et al., 1996). However, the extremely low ergosterol-to-biomass C ratio in the root material indicates that a signi®cant percentage of CHCl3-labile C is probably
root-derived, although the amount of CHCl3-labile C
is much lower than those reported by Mueller et al. (1992) who extracted young intact wheat roots.
The ergosterol-to-biomass C ratios in all soils were almost identical in soil and rhizosphere material. Assuming that the same is true for root colonising microorganisms, microbial derived CHCl3-labile C in
the root fraction can be calculated as follows:
DC= AB=2100,
Table 1
Soil classi®cation according to the Food and Agricultural Organization (FAO) system, soil properties, dominating grass species, distribution of the sample on the fractions sieved bulk soil (<2 mm), rhizosphere material, and root material
No. Soil pH-H2O Bulk density
(kg dmÿ3)
(1) Gleyic Chernozem 7.9 0.99 24.4 96.7 2.7 0.6 Arrhenatherum elatius,Poa trivialis
(2) Orthic Luvisol 7.8 1.54 19.8 94.4 5.3 0.3 Festuca rubra,P. trivialis
(3) Orthic Luvisol 7.7 1.36 22.7 93.6 5.9 0.6 P. trivialis,F. pratensis
(4) Orthic Luvisol 7.7 1.48 18.7 90.8 8.6 0.7 A. elatius,P. trivialis
(5) Orthic Luvisol 6.5 1.21 24.9 92.4 7.3 0.3 Holcus lanatus,Phleum pratense
(6) Orthic Luvisol 7.7 1.20 19.0 86.3 12.7 1.0 Dactylis glomerata,P. trivialis
(7) Dystric Luvisol 5.1 1.46 16.6 86.3 11.7 2.1 Agrostis tenuis,F. rubra
(8) Dystric Cambisol 5.1 1.31 9.7 96.8 2.4 0.8 A. elatius,P. pratensis
(9) Dystric Cambisol 4.3 1.00 8.2 94.6 4.8 0.6 Festuca altissima,H. lanatus
(10) Eutric Cambisol 6.5 1.18 5.7 90.8 8.5 0.7 D. glomerata,A. tenuis
(11) Eutric Fluvisol 7.4 1.36 12.8 95.1 3.9 0.9 Alopecurus pratensis,P. trivialis
(12) Eutric Fluvisol 7.8 1.12 16.3 91.7 7.7 0.6 P. trivialis,D. glomerata
(13) Eutric Fluvisol 7.2 1.21 42.7 84.8 14.7 0.5 A. elatius,Lolium perenne
Mean 6.8 1.26 91.9 7.4 0.7
LSDa 0.1 0.15 6.7 6.4 0.5
b
Total sample weight=sum of bulk soil, rhizosphere and root material.
a
where A is the ergosterol-to-biomass C ratio in the bulk soil, B the ergosterol-to-biomass C ratio in the rhizosphere material, Cthe ergosterol concentration in the root material and D the true microbial biomass concentration in the root material. According to this calculation, on average 33% of CHCl3-labile C was
microbially derived and 66% was non-microbial, root-derived organic C (Table 4). The corrected microbial biomass C concentrations in the root material were still very large compared to the grassland soils, ranging from 7.1 to 36.7 mg gÿ1, but were similar to those values measured in fresh leaf litter of forest ¯oors
(Joergensen and Scheu, 1999). This is still less than the fungal biomass concentration of between 13 and 26% estimated by Salmanowicsz and Nylund (1988) in bulk root material from Scots pine infected with the ecto-mycorrhizal fungus Laccaria laccata, converting ergos-terol data to fungal biomass.
The basic assumption of my work is that the ergos-terol-to-biomass C ratio is identical throughout the dierent soil compartments. This assumption might be wrong. Root colonising microorganisms may have a lower ergosterol-to-biomass C ratio because bacteria contribute a larger percentage of biomass to
rhizo-Table 2
Biomass C, organic C and the biomass C-to-organic C ratio in the sample fractions bulk soil (<2 mm), rhizosphere material, and root material
No. Biomass C (mg gÿ1) Organic C (mg gÿ1) Biomass C-to-organic C (%)
Soil Rhizosphere Root Soil Rhizosphere Soil Rhizosphere Root
1 1.71 2.03 63.6 44.2 51.1 3.9 3.9 4.7
2 1.04 1.96 109.7 34.0 45.8 3.1 4.3 6.6
3 1.19 1.95 49.4 34.1 37.1 3.5 5.3 2.3
4 0.87 1.18 81.8 26.5 31.1 3.3 3.3 4.3
5 0.58 1.08 79.8 22.2 34.1 2.6 3.2 8.2
6 0.70 1.38 142.5 30.3 53.4 2.3 2.6 12.2
7 0.43 1.19 63.5 16.6 19.5 2.6 6.1 7.2
8 0.51 1.48 95.6 18.4 27.7 2.8 3.3 3.7
9 0.56 1.95 40.9 39.3 96.8 1.4 2.0 3.4
10 0.44 0.81 38.6 13.7 27.1 3.2 3.0 2.6
11 0.79 1.62 18.7 35.0 43.7 2.3 3.7 1.1
12 0.80 1.39 73.6 31.3 36.4 2.5 3.8 6.9
13 1.58 2.10 91.5 49.8 58.0 3.2 3.6 7.6
Mean 0.86 1.55 73.0 30.4 43.2 2.8 3.8 5.4
LSDa 0.06 0.50 21.8 4.5 5.0 0.7 1.2 1.1
a
LSD=least signi®cant dierence. Minimum and maximum values are in bold.
Table 3
Ergosterol inmg gÿ1
soil, in % of the total sum and the ergosterol-to-biomass C ratio in the sample fractions bulk soil (<2 mm), rhizosphere material, and root material
No. Ergosterol (mg gÿ1) Ergosterol (% sum) Ergosterol-to-biomass C (%)
Soil Rhizosphere Root Soil Rhizosphere Root Soil Rhizosphere Root
(1) 6.88 8.96 66.5 91.3 3.3 5.5 0.40 0.44 0.11
(2) 2.44 6.98 63.0 81.2 13.0 5.8 0.24 0.36 0.06
(3) 5.32 7.57 99.1 83.0 7.4 9.6 0.45 0.39 0.20
(4) 4.47 5.90 75.5 80.1 10.0 9.9 0.52 0.50 0.09
(5) 1.57 2.33 23.3 86.1 10.1 3.8 0.27 0.22 0.03
(6) 3.24 6.88 34.3 69.6 21.7 8.7 0.46 0.50 0.02
(7) 1.99 2.87 85.3 45.0 8.8 46.2 0.47 0.24 0.13
(8) 2.49 5.22 81.0 76.0 4.0 20.0 0.48 0.35 0.09
(9) 3.37 12.98 95.5 72.8 14.2 13.0 0.60 0.66 0.23 (10) 2.98 5.55 124.1 67.8 11.8 20.3 0.68 0.69 0.32
(11) 2.05 3.72 34.3 80.8 6.1 13.1 0.26 0.23 0.18
(12) 2.23 3.63 78.4 73.6 10.1 16.3 0.28 0.26 0.11
(13) 4.72 5.68 104.5 74.1 15.5 10.3 0.30 0.27 0.11
Mean 3.37 6.02 74.2 75.5 10.5 14.0 0.42 0.39 0.13
LSDa 0.68 1.89 22.2 15.5 9.9 7.2 0.13 0.13 0.18
a
sphere organisms than do fungi (Vancura and Kunc, 1977). Another important component of rhizosphere organisms are mycorrhizal fungi which in grassland soil consist almost exclusively of the vesicular±arbuscu-lar type (Frey et al., 1994). They estimated an ergos-terol concentration of 0.2 mg gÿ1 dry weight in living vesicular±arbuscular mycorrhizal mycelia, 10 times less than in dierent ectomycorrhizal fungi (Salmanowicsz and Nylund, 1988). However, Frey et al. (1994) measured roughly ®ve times smaller ergosterol concen-trations in root material than in another experiment published earlier (Frey et al., 1992). Other ergosterol concentrations in vesicular±arbuscular mycorrhizal fungi have rarely been reported and are in most cases hampered by an insucient sample preparation such as drying (Beilby, 1980; Schmitz et al., 1991). More work needs to be done to identify the origin of the ergosterol found in roots and in the rhizosphere of grassland soils. However, the combination of fumi-gation extraction and ergosterol determination clearly reveals that rhizosphere and root colonising fungi and other microorganisms contribute a signi®cant percen-tage to the total microbial biomass of grassland eco-systems, an aspect which should not be neglected in the future.
Acknowledgements
This study was carried out at the Institute of Soil Science in GoÈttingen. I would like to thank Ingrid Ostermeyer, Karin Schmidt and especially Ulrike Hill for their help with the analyses.
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