Introduction

Steaming of soil is a procedure often practiced in horticultural greenhouse cultures and pot experiments (Luvisi et al. 2008), especially for reducing weeds, soil-borne plant pathogenic fungi, and nematodes (van Loenen et al. 2003), but also for reducing biotrophic arbuscular mycorrhizal fungi (Endlweber and Scheu 2006). Heat treatments reduce the microbial biomass, but also mobilize some soil organic matter (Jenkinson 1966; Powlson and Jenkinson 1976). Without application of substrate, the microbial biomass remains at a low level for long periods after heat-induced disturbance (Banning and Murphy 2008). The regrowth of a reduced microbial biomass could be induced by a simple carbohydrate such as sucrose (Engelking et al. 2008) or by rhizodeposition of a growing plant, which has a more complex chemical composition and which is closer to the natural conditions (Wichern et al. 2008).

Sucrose has been repeatedly added to soil in field experiments to investigate the effects of a simple easily available C source on the turnover of nutrients, mainly N (Wheatley et al. 1991; Ritz et al. 1992; Evans et al. 1998), but also P (Joergensen and Scheu 1999). Sucrose addition to soil was useful for analyzing the relationship between C availability and the relative abundance of bacteria (Fierer et al. 2007). Sucrose has also been applied to soil for increasing the solubility of heavy metals to facilitate their uptake by hyperaccumulating plants (Gramss et al. 2003) and for enhancing plant tolerance to xenobiotics (Sulmon et al. 2007). If sugarcane-derived sucrose is used, it is possible to monitor the changes in substrate-derived microbial biomass and residues simply using the differences in the δ13C values (Engelking et al. 2007a, 2008). Sugarcane is a C4 plant with significantly higher δ13C values than the soil organic matter in Central Europe, mainly derived from C3 plants (Ryan and Aravena 1994; Potthoff et al. 2003).

Microbial residues have often been claimed to be an important intermediate storage pool for plant available nutrients (Mueller et al. 1998; Mayer et al. 2004), which should be released after a substrate is fully decomposed (Engelking et al. 2007a). The first objective of the present pot experiment was to monitor the recovery of a steaming-reduced microbial biomass (C, N, and P) by sucrose addition. A special focus was on the changes in fungal ergosterol. This cell membrane component specifically indicates saprotrophic fungi in agricultural soils (Joergensen and Wichern 2008). Fungi are reckoned to be especially sensitive against heat treatments (Bååth et al. 1995). The second objective was to investigate the recovery of a steaming-reduced microbial biomass by a growing plant (white mustard, Sinapis alba) and its interactions with microbial residues, freshly formed from sucrose addition. Nonmycorrhizal mustard was used due to its efficiency in taking up P from soil without interference by biotrophic fungi. This efficiency may lead to a decline in microbial biomass P.

Materials and methods

Soil and steaming

The soil used for the experiment was sampled in March 2007 at 0–30 cm depth from the arable site Saurasen, Neu-Eichenberg, in the north of Hessia close to Gottingen, Germany (Quintern et al. 2006). The soil was derived from eroded loess overlying clayey sandstone and classified as Stagnic Luvisol, with low water conductivity according to the World Reference Base/Food and Agriculture Organization of the United Nations classification system. The soil contained 9.9 mg organic C, 0.99 mg total N, and 0.41 mg total P per gram soil as well as 20% clay, 72% silt, and 8% sand. Soil pH in water was 6.6. The soil was sieved (<2 mm), and an aliquot for the pot experiment was heated for 12 h between 70°C and 80°C using a soil steamer (Sterilo 1K, MAFAC/Harter, Alpirsbach, Germany) and then stored for 30 days at ambient temperatures in polyethylene buckets before the experiment started. The δ13C value of the bulk soil was −26.6 ± 0.1‰.

Pot experiment

A pot experiment with four replicates each was conducted with the following four treatments: (1) control, (2) addition of 2 mg sucrose C g-1 soil, (3) mustard cultivation, (4) addition of 2 mg sucrose C g-1 soil and mustard cultivation. Each 4 L pot contained 3 kg (on an oven-dry basis) soil adjusted to 50% water holding capacity. All treatments were amended with 50 µg (NH4)2SO4 and 50 µg NaH2PO4 per gram soil. Six plants of white mustard (S. alba L., variety Tango) were sown 5 days after the pot experiment started and thinned to four after emergence. Soil samples for the determination of microbial biomass and ergosterol at day 0 and 5 were taken from the pots before addition of nutrients and sucrose or sowing, respectively. The pots were kept in temperature-controlled greenhouse cabinets (24ºC for 16 h during the day and 18ºC for 8 h during the night) at 60% air humidity without supplementary lighting from 26 April to 18 June 2007. The pots were watered twice a week to make up the water loss and harvested during flowering 47 days after sowing. At harvest, all aboveground plant biomass was removed, weighed, and milled for elemental analysis. The soil used for further soil chemical and soil biological analysis was passed through a sieve (2 mm) to remove roots. The soil attached to the roots was washed away. A moist subsample of the soil-free root material was taken for measuring ergosterol and amino sugars. The rest of the root material was dried for at least 72 h at 60°C and milled.

Analytical procedures

Microbial biomass C and microbial biomass N were estimated by the fumigation–extraction method (Brookes et al. 1985; Vance et al. 1987). At day 52 of the pot experiment, the pre-extraction procedure of Mueller et al. (1992) as described by Muhammad et al. (2007) was used to remove interfering roots. Fumigated and nonfumigated portions of 10 g moist soil were extracted for 30 min by oscillating shaking at 200 rpm with 40 ml, using 0.05 M K2SO4 instead of 0.5 M K2SO4 (Potthoff et al. 2003) and filtered (hw3, Sartorius Stedim Biotech, Göttingen, Germany). Organic C and total N in the extracts was measured after combustion at 850ºC using a Dimatoc 100 + Dima-N automatic analyzer (Dimatec, Essen, Germany). Microbial biomass C was calculated as E C/k EC, where E C = (organic C extracted from fumigated soils) − (organic C extracted from nonfumigated soils) and k EC = 0.45 (Wu et al. 1990). Microbial biomass N was calculated as E N/k EN, where E N = (total N extracted from fumigated soils) − (total N extracted from nonfumigated soils) and k EN = 0.54 (Brookes et al. 1985). Soil microbial biomass P was also measured by fumigation–extraction (Brookes et al. 1982) as described by Joergensen et al. (1995). Microbial biomass P was calculated as E P/k EP/recovery, where E P = (PO 3−4 -P extracted from fumigated soil) − (PO 3−4 -P extracted from nonfumigated soil) and k EP = 0.40 (Brookes et al. 1982). Recovery of added P (25 µg g−1 soil) to account for P adsorption during extraction was calculated as follows: 1 − ((PO 3−4 -P extracted from nonfumigated and spiked soil) − (PO4 3−-P extracted from nonfumigated soil))/25.

The fungal cell membrane component ergosterol was extracted from 2 g of moist soil samples or from 1 g of moist root samples with 100 ml ethanol (Djajakirana et al. 1996). Then, ergosterol was determined by reversed-phase HPLC with 100% methanol as the mobile phase and detected at a wavelength of 282 nm. The fungal and bacterial cell wall components glucosamine and muramic acid were determined according to Appuhn and Joergensen (2006) in 500 mg moist root material after 3-h hydrolysis with 6 M HCl. Fluorometric emission of amino sugar ortho-phthaldialdehyde derivatives was measured at a wavelength of 445 nm with 340 nm as the excitation wavelength (Agilent 1100, Palo Alto, USA). Fungal glucosamine was recalculated into fungal C and muramic acid into bacterial C using the procedure and conversion values proposed by Appuhn and Joergensen (2006) and Engelking et al. (2007b).

Total C and total N in plant material were analyzed gas chromatographically after combustion using a Vario Max CN analyzer (Elementar, Hanau); δ13C was measured on a Delta plus isotope ratio mass spectrometer (Finnigan, Bremen) after combustion using a Carlo Erba NA 1,500 gas chromatograph. Concentrations of P, S, K, Ca, and Mg were determined by HNO3 pressure digestion as described by Chander et al. (2008), and measured by inductively coupled plasma atomic emission spectroscopy (Spectro Analytical Instruments/Kleve).

Calculations and statistical analysis

The isotopic composition of a sample was calculated according to Balesdent and Mariotti (1996) relative to the V-PDB standard. The C content of the fumigated extracts (Cf) was the sum of the C content of the nonfumigated (control) extracts (Cc) and the additionally extracted C from cell lyses by chloroform fumigation (chloroform-labile C, Cb). Then, the δ13C of the fumigated extracts was (Ryan and Aravena 1994):

$$ \left( {{\delta^{13}}{{\text{C}}_{\text{f}}} \times {{\text{C}}_{\text{f}}}} \right) = \left( {{\delta^{13}}{{\text{C}}_{\text{c}}} \times {{\text{C}}_{\text{c}}}} \right) + \left( {{\delta^{13}}{{\text{C}}_{\text{b}}} \times {{\text{C}}_{\text{b}}}} \right) $$
(1)

The chloroform-labile fraction was converted into microbial biomass C, assuming that the extractable and nonextractable fractions of the microbial biomass have the same δ13C value. The part of sugarcane sucrose-derived C (C 4-Csample) was calculated for each single replicate of all treatments from the δ13C data by the following equation (Balesdent and Mariotti 1996):

$$ {{\text{C}}_4} - {\text{Csample}} = {{\text{C}}_{\text{T}}}{\text{sample}} \times \frac{{{\delta^{13}}{\text{Csample}} - {\delta^{13}}{\text{Ccontrol}}}}{{{\delta^{13}}{\text{Csubstrate}} - {\delta^{13}}{\text{Ccontrol}}}} $$
(2)
$$ {{\text{C}}_3} - {\text{Csample}} = {{\text{C}}_{\text{T}}}{\text{sample}} - {{\text{C}}_4} - {\text{Csample}} $$
(3)

where CTsample represented microbial biomass C; δ 13 Csubstrate was the δ13C value of the sugarcane sucrose. The results presented in the tables are arithmetic means and expressed on an oven-dry basis (about 24 h at 105°C).

The significance of sucrose addition and mustard cultivation in the pot experiment at day 52 was analyzed by a two-way analysis of variance (ANOVA). The significance of experimental effects of steaming and sucrose addition was analyzed by a one-way ANOVA using the Scheffé post hoc test, which is robust against the violation of normality and homogeneity of variances (StatView Reference Manual, SAS Institute Inc.). Application of the χ 2 test and the F test showed that the data violated both assumptions without further improvement by log or arcsin transformation. All statistical calculations were performed using JMP 7.0 (SAS Institute Inc.).

Results

Effects of steaming

Steaming led to long-term increase in extractable C, which remained 30 µg g−1 soil higher than before steaming after a storage period of 30 days (Fig. 1a), followed by a decrease in 10 µg C per gram soil until day 52. The addition of sucrose increased the content of extractable C by roughly 100 µg g−1 soil. However, only 40 µg g−1 soil of this increase was sucrose-derived C, and 60 µg g−1 soil were soil organic matter-derived. Both fractions declined considerably until day 52. Steaming had no significant effects on the content of extractable P (Fig. 1b). The application of P fertilizer led to an increase of 6 μg P per gram soil in the control treatment at day 5, equivalent to 12% of the added amount, which decreased by 4 μg P per gram soil until day 52. The application of sucrose led to an immediate decrease in extractable P, which continued until day 52.

Fig. 1
figure 1

Contents of (a) microbial biomass C3-C and C4-C and (b) 0.5 M K2SO4 extractable C3-C and C4-C in pots without white mustard (S. alba) before steaming, immediately before sucrose addition, at day 5 immediately after sowing of mustard, and at day 52 immediately after harvest of mustard; vertical bars show ± one standard deviation; different letters above the bars indicate a significant difference (Scheffé test, P < 0.05)

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Thirty days after steaming, the soil microbial biomass C (Fig. 2) and N (results not shown) was still significantly reduced by 80%, leading to a rather constant microbial biomass C/N ratio around 7 throughout the experiment (Fig. 3a). The steaming-induced decreases of microbial biomass P and ergosterol were only roughly 50% (results not shown), leading to a decrease in the microbial biomass C/P ratio from 11 to values around 4 (Fig. 3b) and an increased ergosterol-to-microbial biomass C ratio from 0.24% to 0.77% at the start of the pot experiment (Fig. 3c). The microbial biomass C/P ratio remained roughly constant throughout the experiment and the ergosterol-to-microbial biomass ratio declined by 45% until day 52. Sucrose addition resulted in a strong increase in the microbial biomass C/P ratio up to 22 at day 5, followed by a decline below 10 at day 52. The ergosterol-to-microbial biomass C ratio was roughly 25% lower at day 5 and 52 in the sucrose treatment without mustard cultivation than in the control treatment.

Fig. 2
figure 2

Contents of microbial biomass C3-C and C4-C in pots without white mustard (Sinapis alba) before steaming, immediately before sucrose addition, at day 5 immediately after sowing of mustard, and at day 52 day immediately after harvest of mustard; vertical bars show ± one standard deviation; different letters above the bars indicate a significant difference (Scheffé-text, P < 0.05)

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Fig. 3
figure 3

Ratios of (a) microbial biomass C/N, (b) microbial biomass C/P, and ergosterol-to-microbial biomass C in pots without white mustard (S. alba) before steaming, immediately before sucrose addition, at day 5 immediately after sowing of mustard, and at day 52 immediately after harvest of mustard; vertical bars show ± one standard deviation; different letters above the bars indicate a significant difference (Scheffé test, s < 0.05)

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Effects of mustard cultivation

Mustard cultivation had significant positive effects on microbial biomass C, N, P, and ergosterol, but the effects were smaller than those of sucrose addition (Table 1). The microbial biomass C/N and C/P ratios were significantly increased by mustard cultivation. Significant second-order interactions were caused by strong increases in ergosterol and the ergosterol-to-microbial biomass C ratio in the combined sucrose and mustard treatment. In contrast, the microbial biomass C/N ratio was significantly decreased by this combination in comparison with the pure mustard treatment.

Table 1 Contents and ratios of microbial biomass indices in pots without and with white mustard (S. alba) after 0, 5, and 52 days
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Approximately 30% of the sucrose C added at day 0 were measured as soil organic C4-C in the soil at day 52 (Table 2). Cultivating mustard had no significant effects on the C loss or on the incorporation of sucrose C into the microbial biomass. In contrast, the application of sucrose led to a significant decrease in the mustard shoot biomass and especially in the mustard root biomass (Table 3). This led to a significant increase in the ratio of shoot C to root C from 18 to 37. Sucrose application only had a significant further depressive effect on the Mg concentration in the mustard shoots, but led to a marked 40% to 60% reduction in the concentration of all nutrients analyzed in the mustard roots (Table 3). Sucrose addition had no effects on the concentrations of ergosterol, glucosamine, or muramic acid, or on the ratio of fungal C-to-bacterial C in the roots (Table 4).

Table 2 Sucrose-derived soil organic C4-C, 0.05 M K2SO4 extractable C4-C, and microbial biomass C4-C in comparison with soil organic matter C3-C, 0.05 M K2SO4 extractable C3-C, and microbial biomass C3-C in pots without and with white mustard (S. alba) at day 52
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Table 3 Dry matter and C yield of the aboveground shoot biomass and the belowground root biomass of white mustard (S. alba) and their nutrient concentrations
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Table 4 Concentrations of ergosterol, glucosamine, and muramic acid, the ratio of fungal C-to-bacterial C in the root samples at day 52
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Discussion

Steaming and microbial indices

During the storage period after steaming and before the pot experiment started, the strong increase in the ergosterol-to-microbial biomass C ratio accompanied by a strong decline in the microbial biomass C/P suggests a considerable restructuring of the microbial community toward saprotrophic fungi. A disproportionately high sensitivity of this type of fungi was not observed, contrasting the view stated in the introduction. In arable and grassland soils, ergosterol is a specific indicator for saprotrophic fungi (Joergensen and Wichern 2008), because biotrophic arbuscular mycorrhizal fungi do not contain ergosterol (Olsson et al. 2003), and ectomycorrhizal fungi are not present. Especially high ergosterol contents can be found in yeasts (Park et al. 1990), which may be responsible for the rapid decomposition of the easily available organic substrates released by steaming. Such heat treatment of soil mobilizes organic C by killing soil microorganisms and by hydrolyzing nonbiomass organic C (Jenkinson 1966; Powlson and Jenkinson 1976). A similar mobilizing effect on P was not observed in the present experiment, probably due to the immediate absorption of heat-mobilized P during extraction (Serrasolsas and Khanna 1995; Endlweber and Scheu 2006).

Substrate-induced recovery of microbial indices

Without further substrate input, the microbial biomass remained more or less at the constant low level obtained by steaming throughout the further experiment. Adding sucrose and cultivating mustard led to a content of microbial biomass C nearly identical to the level before steaming. This result contrasts experiments in which plant growth and the resulting rhizodeposition had no further effect on microbial biomass (Chen et al. 2003; Muhammad et al. 2007). A certain amount of easily available energy is necessary to fill the biological space of a soil (Nannipieri et al. 1983). After a certain percentage of the autochthonous microbial biomass was killed by a treatment, the application of easily available substrate apparently led to a rapid and probably stable increase of the microbial biomass. Above this threshold, new microsites (Wu et al. 1993) or excessive amounts of substrate have to be added to increase the microbial biomass above this biological space for a longer period. This might be the reason why the effects of sucrose application and mustard cultivation were additive without any significant interactions, leading to the maximum content of the total microbial biomass. However, more frequent sampling would help to detect stable conditions.

Within 5 days after application, sucrose is fully decomposed to CO2, microbial biomass, and mainly microbial residues, which comprise exoenzymes, mucous substances, secondary microbial metabolites, and dead tissue (Engelking et al. 2007a). In the sucrose treatments, the lower ergosterol-to-microbial biomass C ratio suggests that the contribution of saprotrophic fungi to the microbial biomass was reduced in favor of bacteria. This shift has been observed by Engelking et al. (2007b) and may be due to the ability of many bacteria to transport sucrose directly through special porins into the cytoplasm (Schmidt et al. 1991). In contrast, fungi have to cleave sucrose by extracellular invertase for uptake (Nehls et al. 2001). Another possibility for the lower ergosterol-to-microbial biomass C ratio in the presence of sucrose would be the incorporation of neutral lipid fatty acids into the biomass of fungi (Rinnan and Bååth 2009). This would result in minor differences in ergosterol, while fungal biomass C would increase. However, in this case, also the microbial biomass C/N should increase concomitantly. As the decline in the ergosterol-to-microbial biomass C ratio was also observed in treatments with sucrose addition, a certain part of this decline might be due to the decomposition of ergosterol in dead fungal tissue accumulated during a period of rapid growth (Mille-Lindblom et al. 2004; Zhao et al. 2005; Engelking et al. 2008).

Cultivating mustard had no effect on the further decomposition of nonbiomass microbial residues. The 40 µg g−1 soil of sucrose-derived extractable organic C observed at day 5 might be the soluble fraction of the microbial residues formed during the early period of rapid microbial turnover. This is certainly also true for the fraction of sucrose-derived extractable organic C detected at the end of the 52-day pot experiment. The strong mobilization of soil organic matter by sucrose application has not been observed before and might be due to desorption and exchange processes from soil colloids. Indications for the existence of these processes are the mobilization of heavy metals after sucrose addition (Gramss et al. 2003), but also the priming effect, i.e., an increased mineralization of soil organic matter-derived C (Kuzyakov et al. 2000; Blagodatskaya and Kuzyakov 2008), after the application of sucrose (Engelking et al. 2007a, b).

The microbial biomass C/P ratio was in most samples below the average of 11.4 in 44 German agricultural soils (Joergensen and Emmerling 2006) and varied in a much larger range than the microbial biomass C/N ratio. This observation contrasts the view of a stable microbial biomass C/P ratio reported by Cleveland and Liptzin (2007). The microbial biomass P turnover is apparently less dependent on the microbial biomass C dynamics. Low microbial biomass C/P ratios may be caused by the storage of excess P in polyphosphates (Gächter and Meyer 1993; Nielsen et al. 2002) and teichoic acids (Grant 1979; Oberson and Joner 2005). High microbial biomass C/P ratios in the present experiment were caused by microbial growth after sucrose addition and by plant uptake during mustard cultivation. A large variability of the microbial biomass C/P ratio points to the possibility that the biomass of soil microorganisms may act as a sink and source for plant-available P (Joergensen and Emmerling 2006).

Sucrose addition and plant growth

In contrast to the absence of mustard effects on decomposing microbial residues, their presence strongly depressed plant growth, especially that of the roots. The reasons for the negative effects cannot be explained by the present data. In the combined sucrose and mustard treatment, the microbial biomass C/N ratio was significantly lower than in the sole mustard treatment and only moderately higher than the ratio of 6.7, given by Jenkinson (1988) as a mean value for microbial biomass in agricultural soils without N limitation. This suggests that the soil microbial biomass in the present experiment had no further N demand, which could compete with mustard for inorganic N. More likely would be a P limitation of the mustard due to P immobilization by soil microorganisms. This is suggested by the high microbial biomass C/P ratio observed at day 5 after sucrose application. However, the P content of the mustard shoots was not specifically reduced.

In contrast, the concentration of all nutrients was reduced in the roots, indicating that not only growth but also nutrient uptake was restricted after sucrose addition. The fungal and bacterial colonization of nonmycorrhizal mustard roots was similar with and without sucrose addition in terms of the ergosterol, glucosamine, and muramic acid concentrations, which were in the medium range observed in a range of root samples from different plants grown in pots with nonsteamed soil (Appuhn and Joergensen 2006). In contrast to these plants, the microbial colonizers of the present mustard roots grown in pots with steamed soil were dominated by bacteria.

Sucrose addition led to a 40% reduction in the shoot weight of Arabidopsis thaliana, but increased its biomass markedly in the presence of atrazine, while Nieminen (2009) also observed an 88% reduction in grass biomass (Deschampsia flexuosa and Calamagrostis epigejos), but no effects on the growth of Norway spruce (Picea abies) seedlings. Wheatley et al. (1991) also found no effects on potato (Solanum tuberosum) yields. The effects of sucrose addition to soil on plant yields may differ depending on plant species and soil properties.

Conclusions

Thirty days after steaming, all microbial biomass indices were still significantly reduced, but no specific effects on saprotrophic fungi were observed. The addition of sugarcane sucrose and cultivation of white mustard led to a recovery of microbial biomass C and N to the initial value. Mustard led to an increased microbial biomass C/P ratio but did not affect the decomposition of microbial residues formed by sucrose addition. However, the presence of these microbial residues had a strong negative effect on plant yield for unknown reasons. Sucrose addition also led to an increase in soil organic matter-derived K2SO4 extractable C.