Fungal N₂O production in an arable peat soil in Central Kalimantan, Indonesia

Abstract

To clarify the microbiological factors that explain high N₂O emission in an arable peat soil in Central Kalimantan, Indonesia, a substrate-induced respiration-inhibition experiment was conducted for N₂O production. The N₂O emission rate decreased by 31% with the addition of streptomycin, whereas it decreased by 81% with the addition of cycloheximide, compared with a non-antibiotic-added control. This result revealed a greater contribution of the fungal community than bacterial community to the production of N₂O in the soil. The population density of fungi in the soil, determined using the dilution plate method, was 5.5 log c.f.u. g⁻¹ soil and 4.9 log c.f.u. g⁻¹ soil in the non-selective medium (rose bengal) and the selective medium for Fusarium, respectively. The N₂O-producing potential was randomly examined in each of these isolates by inoculation onto Czapek agar medium (pH 4.3) and incubation at 28°C for 14 days. Significant N₂O-producing potential was found in six out of 19 strains and in five out of seven strains isolated from the non-selective and selective media, respectively. Twenty-three out of 26 strains produced more than 20% CO₂ during the 14-day incubation period, suggesting the presence of facultative fungi in the soil. These strains were identified to be Fusarium oxysporum and Neocosmospora vasinfecta based on the sequence of 18S rDNA, irrespective of the N₂O-producing potential and the growth potential in conditions of low O₂ concentration.

Key words:

• dilution plate method
• N₂O emission
• substrate-induced respiration-inhibition
• tropical peat

INTRODUCTION

N₂O is an important greenhouse gas produced in soil (Bouwman 1990). It has a destructive potential in the stratospheric ozone layer (Crutzen 1981). N₂O is produced as a byproduct of nitrification and as an intermediate product of denitrification, in which produced NO^- ₃ is respired in conditions of low O₂ availability as an alternative final electron acceptor.

In an arable land in Central Kalimantan, Indonesia, the highest N₂O emission is recorded among the values reported from cultivated tropical, temperate and boreal organic soils (Takakai et al. 2006). To clarify the reason for such high N₂O emission, isolation of the denitrifying bacteria has been carried out as a source of N₂O production (Hashidoko et al.). Although N₂O-producing microorganisms have long been regarded to be bacteria (Zumft 1997), recent focus has been on the contribution of fungi. Shoun and Tanimoto (1991) found potent denitrifying activity in the hyphomycetous fungus Fusarium oxysporum, and Shoun et al. (1992) reported that a range of fungi (Fusarium and its teleomorphs) had the relevant enzyme P-450_nor and N₂O-producing activity in conditions of low O₂ concentration. This result was supported by genome projects (Zhang et al. 2001) and a laboratory experiment that revealed the requirement of a minimal amount of O₂ for the induction of the denitrifying enzyme of fungi (Zhou et al. 2001). These results suggest active N₂O production by a variety of fungi in upland conditions of soil rather than in flooded conditions, which have previously been believed to be sufficient conditions for denitrification.

Recently, a high contribution of fungi to N₂O production in arable soil was demonstrated. Laughlin and Stevens (2002) demonstrated the dominance of fungal N₂O production in a grassland soil, which was evaluated using the substrate-induced respiration-inhibition (SIRIN) method, which was originally developed for the estimation of soil microbial biomass (Anderson and Domsch 1975). Streptomycin (C₂₁H₃₉N₇O₁₂) was used as the bacterial inhibitor and cycloheximide (C₁₅H₂₃NO₄) was used as the fungal inhibitor, and cycloheximide decreased the N₂O emission rate by 89%, whereas streptomycin decreased the rate by 23% (Laughlin and Stevens 2002), indicating a greater contribution of fungal N₂O production in soil. Spokas et al. (2006) showed stimulation of N₂O emissions from soil in a grassland after the application of chloropicrin, and the addition of a bacterial inhibitor (streptomycin and tetracycline: C₂₂H₂₄N₂O₈) suppressed N₂O production by 20%, whereas the addition of a fungal inhibitor (cycloheximide and benomyl: C₁₄H₁₈N₄O₃) suppressed it by 70%, indicating a greater contribution of fungal N₂O production in soil. However, the presence of denitrifying fungi was not confirmed in these studies.

In the present study, the N₂O-producing microbial community was characterized by an inhibition experiment, and the relevant microorganisms were isolated from an air-dried soil sample collected from peatland in Central Kalimantan, Indonesia.

MATERIALS AND METHODS

Soil samples

Surface (0–10 cm depth) soil samples were collected from two croplands (referred to as CL-A and CL-C) and adjacent natural forest (referred to as NF) at Kalampangan village, near Palangka Raya (2°S, 114°E) in Central Kalimantan, Indonesia, in 2003 and 2005. Physicochemical properties of the tropical peat soils and general site information are shown in Takakai et al. (2006). Fresh and large plant debris was removed and the soil samples were stored at room temperature until use.

Nitrification and denitrification potential

Nitrification potential was determined by NO^- ₃ accumulation in the peat soil during 10 days of incubation at 28°C after the addition of 200 µg N g⁻¹soil of (NH₄)₂SO₄, and expressed as the net NO^- ₃ production rate. Air-dried peat soil from CL-A, CL-C and NF (equivalent to 1 g of dried soil, gravimetric water content was 0.82, 0.61 and 1.81 g g⁻¹ dry soil, respectively) was placed in 50 mL of plastic tube and a substrate solution was added to adjust the water content to 4 g g⁻¹ dry soil. The NO^- ₃-N content in the soil was determined as previously described (Yanai et al. 2004). The denitrification potential was determined using the method described in Yanai et al. (in press), with a slight modification in the concentration of KNO₃ in 8 g nutrient broth per liter from 5 mmol L⁻¹ to 50 mmol L⁻¹, because of originally high NO₃-N content in the soil at CL-A (236 µg N g⁻¹ dry soil; Takakai et al. 2006). The air-dried peat soil (equivalent to 1 g of dried soil) was placed in a 30 mL grass vial and 5 mL of the liquid medium was added. After sealing with a butyl rubber septum in the ambient condition, 100% C₂H₂ was added to make the headspace concentration of C₂H₂ 10% to suppress the N₂O-reducing activity of denitrifiers (Yoshinari et al. 1977; referred to as +C₂H₂) and the total pressure in the vial was set to the atmospheric pressure. To estimate N₂O-reducing activity, vials without C₂H₂ addition were also prepared, and the samples were incubated at 30°C. N₂O concentration in the headspace was periodically measured using a gas chromatograph (GC-8A; Shimadzu, Kyoto, Japan) equipped with a thermal conductivity detector (TCD). The running condition is described in Wada and Toyota (2007).

Substrate-induced respiration-inhibition experiment

Air-dried peat soil from CL-A (20 g of air-dried material, gravimetric water content was 0.21 g g⁻¹ dry soil) was placed into nine Petri dishes (1.3 cm height and 8.5 cm diameter) without compaction. A glucose solution (final concentration: 5 mg glucose g⁻¹ dry soil), including each antibiotic (streptomycin or cycloheximide), was added into the soil to adjust its water content to 1.1 g g⁻¹ dry soil (58% of water-filled pore space [WFPS]). The concentrations of the antibiotics were 3 mg streptomycin g⁻¹ dry soil and 15 mg cycloheximide g⁻¹ dry soil (Laughlin and Stevens 2002). In this condition, CO₂ production was significantly decreased (23 and 26%) by the addition of streptomycin and cycloheximide, respectively (P < 0.01). After the addition of the substrate solution, peat soils were incubated for 6 h in a laboratory (18–23°C) and the N₂O emission rate was determined using the closed chamber method as shown in Yanai et al. (2007).

Enumeration and isolation of filamentous fungi in soils

Based on the result of the SIRIN experiment, colony-forming units of culturable fungi were estimated using the dilution plate method with rose bengal medium as a non-selective cultivation and GMBP medium (Takehara et al. 2003) as a selective cultivation for Fusarium. After inoculation of the soil dilution, the media were incubated at 28°C for 4 days. Part of the colonies was randomly collected, purified and a freeze stock (in 20% glycerol solution) was prepared for further analysis.

Determination of the N₂O-producing potential of isolated fungi

Modified Czapek medium (30 g sucrose, 0.5 g MgSO₄ 7H₂O, 1 g KH₂PO₄, 0.5 g KCl, 2 g NaNO₃, 0.01 g Fe₂(SO₄)₃ in 1 L of distilled water, pH 4.3) with 50 µg mL⁻¹ streptomycin was solidified with 2% (w/v) agar and was used to test the N₂O-producing potential of the fungal isolate. The isolate was aseptically inoculated in the Czapek agar medium in a 100 mL Erlenmeyer flask sealed with a rubber stopper in the ambient condition, and incubated at 28°C. After 14 days of incubation, 0.5 mL of the headspace gas was withdrawn from the headspace of the flask through a rubber stopper and the CO₂ and N₂O concentrations were analyzed using the TCD gas chromatograph. When the N₂O concentration was less than 200 p.p.m.v, an additional 5 mL of headspace gas was withdrawn and the N₂O concentration was analyzed using a gas chromatograph (GC-14A: Shimadzu, Kyoto, Japan) equipped with a ⁶³Ni-electron capture detector (ECD-GC). The running condition of the ECD-GC is shown in Yanai et al. (2007). In the present study, Fusarium oxysporum (Fusarium oxysporum f. sp. lactucae) was examined as a positive control (denitrifying fungi that can grow under conditions of low O₂ concentration and produce N₂O) and Pythium (Pythium ultimum Trow var ultimum OPU407) and Rhizoctonia sp. were examined as negative controls (fungal strains that can grow only under oxic conditions and do not have N₂O-producing activity). After determination of the headspace gas, BTB (C₂₇H₂₈O₅Br₂S) solution was added to estimate the pH of the agar medium.

Fungal DNA extraction and analysis based on 18S rDNA

Pure cultures of filamentous fungi were grown in the modified Czapek medium for 3–10 days at 30°C prior to DNA extraction. Fungal mycelia were collected from liquid culture and 400–800 µL (bulk volume) was added to 1.5 mL tubes. Mycelium material was washed twice with TE buffer and transferred to 2 mL tubes containing 0.75 g of 0.1-mm diameter zirconium beads and 0.25 g of 0.5-mm diameter glass beads. The mycelium material was suspended in 1,200 µL of a lysis buffer containing 0.1 mol L⁻¹ Tris-HCl (pH 8.0), 0.3 mol L⁻¹ ethylenediaminetetraacetic acid (pH 8.0) and 0.5% sodium dodecyl sulfate, and then bead beating was carried out using a Bead Smash 12 (BS-12; Wakenyaku, Kyoto, Japan) twice for 1 min at 5,000 rev min⁻¹. Then, the tubes were centrifuged for 5 min at 12,000 g and 700 µL of the supernatant was transferred into new 2 mL tubes. Each supernatant was suspended in 377 µL of 5 mol L⁻¹ NaCl and 270 µL of 10% CTAB (cetyltrimethylammonium bromide), and incubated at 60°C for 10 min. After cooling at room temperature, 500 µL of chloroform was added to each sample, mixed vigorously using a vortex for 15 s and then centrifuged for 20 min at 15,000 g. Then, 400 µL of the aqueous layer was transferred to a new 1.5 mL tube, 1 µL of 10 mg mL⁻¹ RNase was added and incubated at 37°C for 30 min. After the incubation, 40 µL of 3 mol L⁻¹ CH₃COONa and 400 µL of isopropanol were added to each sample for the precipitation of DNA. The tubes were placed at room temperature for 30 min and centrifuged at 4°C for 5 min at 20,630 g. All supernatant was removed and centrifuged again at 4°C for 5 min at 20,630 g after the addition of 500 µL of 70% ethanol to wash the remaining pellet. The pellets were air-dried and re-suspended in 20–50 µL of sterile TE buffer. The subsequent DNA yields and quality were assessed using standard electrophoresis through a 1% (w/v) agarose gel followed by ethidium bromide staining.

The forward primer NS1-seq (TGG AAA ACG GCC AGT GTA GTC ATA TGC TTG TCT C) and the reverse primer FR1-GC (CCC CCG CCG CGC GCG GCG GGC GGG GCG GGG GCA CGG GCC GAI CCA TTC AAT CGG TAI T) were used for polymerase chain reaction (PCR) amplification of the 18S rDNA gene (Vainio and Hautula 2000). The reaction was carried out in a 25 µL volume that contained approximately 15 ng DNA, 10 mmol L⁻¹ Mg-free buffer, 0.2 mmol L⁻¹ deoxynucleoside triphosphates, 3.75 mmol L⁻¹ MgCl₂, 0.4 µg µL⁻¹ BSA (Bovine Serum Albumin), 0.16 µmol L⁻¹ of each primer and 2 U µL^_1 of Taq DNA polymerase (TaKaRa Ex Taq; TaKaRa Bio, Shiga, Japan). The samples were subjected to 30 cycles of 94°C for 30 s, 48°C for 45 s and 72°C for 180 s for denaturation, annealing and elongation steps, respectively, using a thermal cycler (Dice mini; TaKaRa Bio). An initial denaturation step (94°C for 5 min) was used to ensure complete denaturation of the template DNA. The PCR products were first analyzed using electrophoresis in 0.75% (w/v) agarose gels and ethidium bromide staining. Direct sequencing of the PCR products derived from various isolates was done by Hitachi Science Systems (Tokyo, Japan). The sequences obtained have been submitted to the DDBJ (DNA Data Bank of Japan) under accession numbers AB302197 to AB302214, and were examined using the DDBJ homology search system BLAST (http://blast.ddbj.nig.ac.jp/top-j.html).

RESULTS AND DISCUSSION

The CL-A and CL-C peat soils showed higher activity of nitrification and denitrification compared with the NF peat soil (Figs 1,2). These results are comparable with the amount of N₂O emission from these fields (Takakai et al. 2006), suggesting that biochemical N

Figure 1  Nitrification potential of the peat soils collected from arable land (CL-A, CL-C) and adjacent natural forest (NF). Values shown are mean ± standard deviation (n = 2).

Figure 2  Denitrification potential of the peat soils collected from arable land (CL-A, CL-C) and adjacent natural forest (NF). Values shown are mean ± standard deviation (n = 2). After a 70-h incubation period, N2O production of the CL-A soil sample incubated with and without C2H2 was significantly different according to an unpaired t-test (P < 0.05). +, the incubation of soil with 10% C2H2 in the headspace; –, the incubation of soil without 10% C2H2 in the headspace.

transformation was promoted in these croplands. In particular, the CL-A peat soil had the highest activity of nitrification and denitrification potential among the peat soils (Figs 1,2). N₂O production in the CL-A peat soil was significantly lower in the incubation with C₂H₂ than in the incubation without C₂H₂ (70 h of incubation period [P < 0.05]; Fig. 2). This result does not correspond to the principal of the C₂H₂ inhibition method (Yoshinari et al. 1977), that is, N₂O-reducing activity is suppressed by the addition of 10% C₂H₂ and total denitrification activity (N₂O + N₂ production) is estimated through the observation of N₂O production. Although the presence of some specific denitrifying microorganisms whose growth is significantly inhibited by C₂H₂ was pointed out by Tiedje (1994), we did not encounter this type of

Figure 3  Effect of the addition of antibiotics on N2O production in the peat soil (CL-A). Values shown are mean ± standard deviation (n = 3). NS, not significant (P = 0.121); **P < 0.01 (Dunnett's test results).

soil in the previous study (Yanai et al. in press), that is, N₂O production was lower in the soil incubated with C₂H₂ than in the soil without C₂H₂. Therefore, we speculated that the CL-A peat soil might harbor a unique denitrifier and the presence of N₂O-producing microorganisms in the CL-A peat soil was explored in the present study.

The N₂O emission rates from the CL-A peat soil were 33 ± 6.3, 23 ± 7.2 and 6.5 ± 3.1 ng N₂O-N g⁻¹ dry soil per hour in the non-antibiotic control, the streptomycin addition and the cycloheximide addition, respectively (mean ± standard deviation [SD], n = 3; Fig. 3). The cycloheximide addition decreased the N₂O emission rate by 81% (P < 0.01), whereas streptomycin decreased the rate by 31% (P = 0.12), although the bacterial and fungal biomass ratio was approximately 1:1. As a high N₂O emission rate (approximately 8,000 µg N m⁻² h⁻¹) was recorded at the CL-A site at 60% WFPS (Takakai et al. 2006), and the moisture condition in the SIRIN experiment was similar (58% WFPS), this result suggests that fungal N₂O production may have a considerably high contribution at the CL-A site. In this sense, the unusual pattern of N₂O production of soil incubated with or without C₂H₂ (Fig. 2) can be interpreted as fungistasis caused by trace C₂H₄ in the C₂H₂ (Liebman and Epstein 1994), although we did not test this experimentally.

The population density of culturable fungi in the CL-A peat soil was 5.5 log c.f.u. g⁻¹ dry soil (n = 2) and 4.9 log c.f.u. g⁻¹ dry soil (n = 3) in non-selective and selective media, respectively, and six out of 19 strains from the non-selective medium and five out of seven strains from the selective medium had N₂O-producing potential (Table 1). Irrespective of the N₂O-producing potential, some strains showed unexpectedly high CO₂ production, that is, the CO₂ concentration in the headspace exceeded 20% during the 14-day incubation period, suggesting significant growth even after the consumption of O₂ in the ambient air (Table 1). The nearest species of these strains were identified as Fusarium oxysporum and Neocosmospora vasinfecta (Table 1). In general, eukaryotic organisms, including fungus, only use O₂ as a final electron acceptor. In fact, the Pythium did not produce N₂O and the CO₂ concentration did not exceed 20% even after a 14-day incubation (data not shown). In contrast, the Rhizoctonia did not produce N₂O, but the CO₂ concentration in the

Table 1 Isolation of fungi from peat soil in a crop land (CL-A) and their potential denitrification activity (14-day incubation period)

Source	Strain or accession number	Facultative growth^†	N2O production^‡	Nearest relative (accession number)	Homology (%)	BTB test
Fusarium oxysporum	AB302203	++	+++	Fusarium oxysporum (DQ916150)	98.5	Yellow
Pythium ultimum	OPU407	–	–	n.d.	n.d.	Yellow
Rhizoctonia sp.	RS	+	–	n.d.	n.d.	Yellow
Selective medium	AB302200	+	+	Neocosmospora vasinfecta (U32414)	99.6	Yellow
	AB302207	+	+	Neocosmospora vasinfecta (U32414)	99.8	Yellow
	43	+	++	n.d.	n.d.	Yellow
	AB302212	+	++	Neocosmospora vasinfecta (U32414)	99.8	Green
	AB302214	+	++	Fusarium oxysporum (AB110910)	100	Yellow
	AB302201	++	++	Fusarium oxysporum (AB110910)	99.9	Green
	AB302206	++	+++	Fusarium oxysporum (AB110910)	100	Yellow
Non-selective medium	A-15	–	+	n.d.	n.d.	Yellow
	AB302211	–	+	Fusarium oxysporum (AB110910)	100	Green
	A-14	+	–	n.d.	n.d.	Yellow
	AB302204	+	–	Fusarium oxysporum (AB110910)	99.9	Yellow
	A-16	+	–	n.d.	n.d.	Blue
	A-12	+	+	n.d.	n.d.	Yellow
	AB302199	+	+	Neocosmospora vasinfecta (U32414)	99.9	Yellow
	5	+	+	n.d.	n.d.	Blue
	AB302205	+	+	Neocosmospora vasinfecta (U32414)	98.4	Green
	17	+	+	n.d.	n.d.	Yellow
	3	+	+	n.d.	n.d.	Yellow
	AB302202	+	+	Fusarium oxysporum (DQ916150)	98.9	Yellow
	AB302208	+	+	Neocosmospora vasinfecta (U32414)	99.9	Blue
	AB302197	+	++	Neocosmospora vasinfecta (U32414)	99.5	Yellow
	AB302210	+	+++	Neocosmospora vasinfecta (U32414)	99.6	Yellow
	AB302198	++	++	Neocosmospora vasinfecta (U32414)	99.5	Blue
	AB302209	++	++	Fusarium oxysporum (AB110910)	99.8	Yellow
	19	++	+++	n.d.	n.d.	Yellow
	AB302213	++	+++	Fusarium oxysporum (AB110910)	100	Yellow
^†CO2 accumulation in the headspace less than 20% (–), more than 20% (+) or more than 30% (++).
^‡N2O accumulation in the headspace less than 2 p.p.m.v (–), less than 10 p.p.m.v (+), more than 10 p.p.m.v (++) or more than 200 p.p.m.v (+++).BTB, C27H28O5Br2S; n.d., not determined.

headspace exceeded 20% during the 14-day incubation period (Table 1), suggesting the presence of fungi that have the potential to grow in conditions of low O₂ concentration (Lockhart et al. 2006; Shoun et al. 1992).

In addition, despite high CO₂ accumulation in the flask and the use of acidified agar medium (pH 4.3), eight out of 31 cultures showed neutral or alkali pH after the 14-day incubation period (Table 1), suggesting ammonia fermentation by fungi (Zhou et al. 2002). Denitrification by fungi is an ATP-producing reaction in mitochondria at low O₂ conditions that results in the production of N₂O, whereas ammonia fermentation by fungi is an ATP-producing reaction at cytosol at low O₂ conditions that does not result in the production of N₂O. Therefore, this study experimentally suggested that a variety of fungi in soil can grow under conditions of low O₂ concentration and a some of these fungi are involved in N₂O production in the soil.

Takakai et al. (2006) suggested that changing land use from forestry to agriculture increases N₂O production in the soil. Although there is no experimental evidence for this suggestion, such changes in land use can affect microbial community structure (enrichment of N₂O-producing microorganisms), resulting in the stimulation of N₂O production in the soil. Further study is expected to clarify the contribution of fungal communities to N₂O emissions from peat soil, the diversity of the fungal community that produces N₂O in the soil, and the relationship between N₂O emission from soil and soil-borne plant disease because the fungi obtained in this study are Fusarium oxysporum and Neocosmospora vasinfecta, which are known to be plant-pathogenic fungi (O'Donnell and Gray 1995).

ACKNOWLEDGMENTS

We thank Mr S. Yoshida (Nagano Chusin Agricultural Experiment Station) for providing a fungal strain of Fusarium oxysporumf. sp. lactucaeand Dr M. Tojo (Osaka Prefecture University) for providing a fungal strain of Pythium ultimum Trowvar ultimumOPU407. We also thank Mr M. Amari, Ms E. Sato, Mr T. Shirakashi and Ms A. Enomoto for their technical assistance. We are grateful to Professor M. Okazaki for fruitful discussions. This study was financially supported by the Japan Society for the Promotion of Science (JSPS) Research Fellowships for Young Scientists (17-6518), a Japanese Grant-in-Aid for Science Research from the Ministry of Education, Culture, Sports, Science and Technology (13574012), and the JSPS-LIPI (Indonesian Institute of Sciences) Core University Program.