European Journal of Soil Biology 53 (2012) 121e125
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Original article
Xylophagous termites: A potential sink for atmospheric nitrous oxide Muhammad Zeeshan Majeed a, d, *, Edouard Miambi b, Alain Robert c, Martial Bernoux a, Alain Brauman a a
Institut de Recherche pour le Développement (IRD), UMR Eco&Sols, 2 Place Pierre Viala, 34060 Montpellier, France UMR BioEMCo, IBIOS, Université Paris Est, Créteil, France c UMR BioEMCo, IBIOS, Centre de recherche d’Île-de-France, Bondy, France d University College of Agriculture, University of Sargodha, Sargodha, Pakistan b
a r t i c l e i n f o
a b s t r a c t
Article history: Received 27 July 2012 Received in revised form 2 October 2012 Accepted 11 October 2012 Available online 25 October 2012 Handling editor: Bryan Griffiths
To provide a better understanding of soileatmosphere gas exchange processes, this study describes the atmospheric nitrous oxide (N2O) uptake by xylophagous termites and the biological process involved. The N2O consumption rates of three xylophagous termite species (Hodotermes mossambicus, Nasutitermes voeltzkowi and Hodotermopsis sjoestedti) were determined in incubation vials with ambient, artificially enhanced N2O concentrations in the headspace. Live individuals of the three termite species significantly decreased N2O concentrations (88%) in the headspace of the vials after 24 h incubation in the dark. The acetylene reduction assay method applied to N. voeltzkowi, a xylophagous termite species, showed a decrease in N2O uptake in acetylene-treated individuals, indicating the potential involvement of termite gut denitrifying microbes. The N2 formed is potentially subjected to assimilation via nitrogenase reductase into termite biomass through biological fixation as demonstrated by the reduction of acetylene to ethylene at an average rate of 18.21 1.34 nmol C2H4 g1 dw d1. Further studies should focus on measurements of N2O-reductase (nosZ) gene activity in termite guts to gain a better understanding of the N2O reduction process in xylophagous termite species. Ó 2012 Published by Elsevier Masson SAS.
Keywords: Xylophagous termites Nitrous oxide nosZ gene Denitrification Acetylene reduction assay (ARA)
1. Introduction Nitrous oxide (N2O) is the third most important climate-forcing tropospheric gas after carbon dioxide (CO2) and methane (CH4) [1]. Soils in both natural and agroecosystems account for more than 60% of atmospheric N2O emissions [1]. Microbe mediated aerobic nitrification and anaerobic denitrification processes are primarily responsible for N2O emissions from soils [2], mainly due to N fertilizers. Many studies have been carried out to quantify these emissions on both temporal and spatial scales [3e5]. Soils are major sources of N2O and have also been reported to act as sinks for atmospheric N2O [6e8]. Complete respiratory denitrification, with a final step of N2O reduction to nitrogen (N2), has been described as the main process involved in N2O uptake and is essentially catalyzed by N2O-reductase, the only denitrification enzyme known to reduce N2O to N2, encoded by the nosZ gene [9,10]. Termites are a prominent feature of soils in all tropical and subtropical biomes of the world and play a major role in C and N
* Corresponding author. Institut de Recherche pour le Développement (IRD), UMR Eco&Sols, 2 Place Pierre Viala, 34060 Montpellier, France. Tel.: þ33 (0)4 99613045; fax: þ33 (0)4 99612119. E-mail address:
[email protected] (M.Z. Majeed). 1164-5563/$ e see front matter Ó 2012 Published by Elsevier Masson SAS. http://dx.doi.org/10.1016/j.ejsobi.2012.10.002
mineralization [11,12]. Soil-feeding and humus-feeding termites usually thrive on N-rich soil substances [13,14] and have recently been shown to emit N2O into the atmosphere [15]. However, xylophagous termites feed on N-deficient food comprising dry, sound wood or grass and plant litter fragments with N content as low as 0.03e0.1% [16,17]. These wood-feeding and grass-feeding termites rely essentially on the symbiotic bacterial communities in their gut to supplement the N through biological fixation [17e 19]. Previous culture-independent molecular studies have demonstrated the presence of a large phylogenic diversity of nitrogenase reductase (nifH) genes in xylophagous termite guts, carried by N2fixing bacteria, dominated by members of Clostridia, Spirochaetes and gram-negative protoeobacteria including members of genera Desulfovibrio, Enterobacter, Rhizobia [17e21]. The termite gut constitutes a specific microhabitat with physical and chemical conditions such as an alkaline pH with oxygen and hydrogen gradients [22], ideal for denitrification activities including reduction of N2O to N2 [2]. A recent molecular quantification analysis revealed the presence of denitrifying microbial genes, including N2O-reductase (nosZ) genes in the guts of xylophagous termites (Brauman et al., unpublished results). It may, therefore, be possible that, in tropical soil profiles, edaphic xylophagous termites may be a potential sink for atmospheric N2O. This could be the result of i) a direct reduction of
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atmospheric N2O to ammonia (NH3) [23], as this seems more efficient and less energy-consuming for a termite system on a thermodynamic basis as reported by Shestakov and Shilov [24] or ii) an indirect pathway involving sequential reduction of N2O to N2 followed by biological assimilation of reduced N2 to cell biomass in the form of ammonia [25,26]. In order to test these hypotheses, the N2O fluxes from three xylophagous termite species, Hodotermes mossambicus, Nasutitermes voeltzkowi and Hodotermopsis sjoestedti, were determined in vitro and acetylene inhibition was used on N. voeltzkowi to assess the denitrification processes in xylophagous termite guts.
each vial. There were three replicates for each setup. The acetylene reduction assay (ARA) was used to determine whether N2O was reduced to N2 in the termite gut. The nitrogenase enzyme responsible for N2-fixation also reduces acetylene (C2H2) to ethylene (C2H4) [27] and so the production of C2H4 can be used to measure the nitrogenase activity. Since the presence of acetylene also blocks the reduction of N2O to N2 by nitrous oxide reductase, it is possible to measure the denitrification (NO 3 / N2O / N2) in the same experiment by measuring the production of N2O. Therefore, N2O emitted in the presence of acetylene represents the total of N2O þ N2 emissions.
2. Material and methods
2.3. Headspace gas sampling
2.1. Termite collection and rearing
The headspace gases (6 mL) were drawn off for analysis using a Luer lock syringe at the beginning of each experiment (T0) and at the end of the incubation period (T24 or T7 depending on the experiment). The gas pressure in the vials was maintained by inserting 6 mL of ambient air into the vials through a sterile filter just after sampling at T0. The sampled headspace gas (6 mL) was transferred to 5.5 mL pre-evacuated ExetainerÒ vials (Labco Ltd, High Wycombe, England), creating overpressure to avoid changes in composition in the gas during storage before further analysis.
Colonies of three xylophagous termite species (H. mossambicus, N. voeltzkowi and H. sjoestedti) were collected from different tropical biomes (Table 1) and were transferred to the laboratory where they were reared for many months under controlled, semi-natural conditions at 27 2 C and 80% relative humidity. Dry wood or grass was fed to the termites depending on their feeding habits. 2.2. Experimental design
2.4. N2O analysis
The first experimental setup assessed the N2O uptake rates in these three xylophagous termite species. They were performed using sterilized 60 mL glass vials (Wheaton Inc., Millville, USA) containing 0.5 g of live, healthy workers with 11 H. mossambicus termites, 112 N. voeltzkowi termites and 8 H. sjoestedti termites. The vials were sealed at ambient pressure with air-tight sterile butyl rubber septa and aluminium caps and were incubated in the dark for 24 h. There were four replicates for each termite species studied together with controls (empty vials). The second experimental setup was used to determine the effect of the termite nest materials on the N2O uptake rate and was performed for N. voeltzkowi only. N. voeltzkowi workers (0.5 g) were put into the vials without nest materials and with 5 g of sterilized fragments or 5 g of non-sterilized fragments from their nests. The experiment was also carried out with the nest materials without termites and with dead termites obtained by crushing individual live termite workers using sterilized forceps 30 min before the start of incubation. Three replicates were made for each setup. The third experimental setup was designed to determine the biological processes involved in the uptake of N2O in the termite gut. The experiments were performed in vials as described above. The setups included (i) live termites (0.5 g per vial), (ii) fragments of termite nest materials (5 g) and (iii) a combination of termites and termite nest materials. There were two treatments, with and without acetylene. For the acetylene treatment, 10% of the headspace air was replaced at the beginning of the experiment, using a syringe, by an equal quantity of purified C2H2 gas (Linde, France). To avoid substrate limitation, 50 mL of pure N2O (N48, Air Liquide, France), which corresponded to about 833 ppm N2O, was added to
The N2O concentrations were measured in the headspace gas samples using a gas chromatograph (CP-3800 VARIANÒ STAR; Agilent Technologies, Santa Clara, CA, USA). This apparatus is equipped with a 63Ni (15 mCi) electron capturing detector (ECD), maintained at 300 C. The carrier gas consisted of 90% argon and 10% methane. The machine was calibrated before the analyses with ambient air and commercial standards of helium and N2O (N48, Air Liquide, France) to give an accurate measurement of the N2O concentrations from 0 to 880 ppm. Samples of 200 mL were inserted using 1 mL Luer lock syringe. The chromatograms were processed using STARÔ Workstation Version 6.0 (Agilent Technologies, Santa Clara, CA, USA) to determine the concentration of N2O in the sample. 2.5. C2H4 analysis Headspace gas samples (100 mL) were assayed for ethylene concentration using a gas chromatograph with flame ionization detector (AgilentÒ 6850 Series GC System). Pure nitrogen and hydrogen were used as the carrier and combustion gases, respectively. The injector, detector and oven temperatures were maintained at 90, 120 and 60 C. The retention time for C2H4 was 1.34 min and the runtime for one sample was about 4 min. HPChem (V 4.0.1.1) was used to determine the C2H4 concentration. 2.6. Statistical analysis Statistica V 7.1 (StatSoftÒ Inc., Tulsa, OK, USA) was used for the statistical analyses. The differences in N2O influx and ethylene
Table 1 Termite species studied. Species
Feeding group
Family
Origin
Biome
Geographical distribution
Hodotermes mossambicus (Hagen 1853) Nasutitermes voeltzkowi (Wasmann 1911) Hodotermopsis sjoestedti (Holmgren 1911)
Grass-harvesting
Hodotermitidae
South Africa
Dry savanna
Afrotropical
Dry wood-feeding
Termitidae
Mauritius Island
Wet forest
Damp wood-feeding
Termopsidae
Vietnam
Wet forest
Neotropical, Afrotropical, Malagasy, Oriental, Papuan, Australian Indo-Malayan
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efflux values were based on the means of three to four independent replicates for each setup and treatment. Prior to statistical comparison, data normality was checked using the ShapiroeWilk test. The means were compared by one-way analysis of variance (ANOVA) and then tested using Fisher’s least significant difference (LSD) test with standard probability (p ¼ 0.05). In the ARA experiment, the t-test (multiple independent samples) was used to compare the effect of the C2H2 treatment at 95% confidence interval. 3. Results 3.1. N2O consumption by xylophagous termites The three xylophagous termite species reduced the N2O concentration in the headspace compared to the initial ambient air concentration (Fig. 1). The mean negative flux observed for the three species was 0.76 0.11 nmol N-N2O vial1 d1, significantly lower (p < 0.05) than the control value of 0.02 0.07 nmol N-N2O vial1 d1. The grass-harvesting H. mossambicus had the highest uptake (0.93 0.19 nmol N-N2O) and the wood-feeding N. voeltzkowi had the lowest uptake (0.63 0.03 nmol N-N2O), but the differences between the 3 species were not significant (p > 0.05). 3.2. Effect of N. voeltzkowi nest material fragments on N2O fluxes The fluxes of N2O for live and dead worker termites and sterilized and non-sterilized fragments from their nests incubated either separately or in combination, are shown in Fig. 2. A significant N2O uptake rate was observed in all setups with termite workers, whether they were alive or dead. The highest uptake rate was for live termites (0.64 0.11 nmol N-N2O g1 dw (termite dry weight) d1) and the lowest rate (0.38 0.06 nmol NN2O g1 dw d1) was for dead termites. There was no significant difference between the uptake rates of live termites on their own and live termites incubated with sterilized or non-sterilized fragments of nest material. Nest material fragments on their own emitted an average of 0.26 0.06 nmol N-N2O g1 dw d1, which was significantly different from the other setups (p < 0.05). 3.3. Effect of C2H2 on N2O consumption by N. voeltzkowi
Fig. 2. N2O fluxes for Nasutitermes voeltzkowi worker termites and nest fragments. N(ns) ¼ non-sterilized nest fragments, N(s) ¼ sterilized nest fragments, T þ N(ns) ¼ live termites with non-sterilized nest fragments, T þ N(s) ¼ live termites with sterilized nest fragments, T ¼ live termites, DT ¼ dead termites. The bars represent the means (n ¼ 3) SE. Lowercase letters at the top of the bars indicate significant differences between setups (p < 0.05).
Termites significantly decreased the N2O concentration in the headspace whether on their own or in combination with nest material fragments and in the presence or absence of C2H2. The mean N2O uptake values measured for termites were significantly different (p < 0.001): 89.48 8.10 nmol N-N2O g1 dw d1 and 342.10 37.66 and with or without C2H2, respectively. The N2O fluxes from nest materials ranged from 3.30 to 32.08 9.02 nmol NN2O g1 dw d1 with or without C2H2. For all the setups, the fluxes recorded with and without C2H2 were significantly different. Adding acetylene made it possible to assess nitrogenase activity which was determined for live N. voeltzkowi termites and their nonsterilized nest fragments. The average rate of acetylene reduction to ethylene in treatments with live termites was about 18.21 1.34 nmol C2H4 g1 dw d1, while the rates for the nest materials were negligible (Fig. 4). 4. Discussion Although N2O-derived N acquisition through biological fixation has been shown for some phytoplankton populations in oceans
Fig. 3 shows the N2O fluxes for live termites and non-sterilized material from their nests, incubated separately or in combination, with or without the addition of 10% C2H2 in the vial headspaces.
Fig. 1. N2O fluxes for 24 h incubation in the dark, measured for various setups including an empty vial (Ctrl) and vials containing 0.5 g of live Hodotermopsis sjoetedti (Hs), Nasutitermes voeltzkowi (Nv) and Hodotermes mossambicus (Hm) worker termites. The bars represent the means (n ¼ 4) SE. Lowercase letters at the top of the bars indicate significant differences between setups (p < 0.05).
Fig. 3. N2O fluxes for 24 h incubation of Nasutitermes voeltzkowi workers and nest fragments with additional N2O and with (þ) or without () 10% (v/v) acetylene in the headspace. Setups had live termite workers (T), non-sterilized nest fragments (N(ns)) and termite workers with non-sterilized nest fragments (T þ N(ns)). The bars represent the means (n ¼ 3) SE. For treatments with and without acetylene, lowercase letters at the top of the bars indicate significant difference between setups (p < 0.05). For each setup, the flux with or without acetylene was significantly different (p < 0.001).
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Fig. 4. Nitrogenase activity (C2H4 production rate) of Nasutitermes voeltzkowi with 10% (v/v) acetylene in the headspace. Setups had live termite workers (T), non-sterilized nest fragments (N(ns)) and termite workers with non-sterilized nest fragments (T þ N(ns)). The bars represent the means (n ¼ 3) SE. Lowercase letters at the top of the bars indicate significant differences between setups (p < 0.05).
[28], this subject is still poorly documented for soil invertebrates. This study reports for the first time evidence of reduction of N2O by termite-gut denitrifiers and its possible assimilation into termite biomass through biological N fixation processes in terrestrial xylophagous termites. An experiment performed in vitro on two xylophagous termite species and one grass-feeding -termite species revealed a significant, reproducible uptake of N2O as compared to controls. The Nuptake may be explained by the need to compensate for the N-poor diets of these insects. These features may be common to most organisms feeding on substrates with high C to N ratios. Recent research has shown that soils with high C to N ratio have a high affinity for N2O [7]. However, it has also been reported recently that soil-feeding termites emit N2O [15]. This was confirmed by our findings that indicated that the fate of N in termite guts is governed by the feeding guilds (Brauman et al., unpublished results). Karsten and Drake [29] have also reported N2O consumption by earthworms in some of their microcosms. As earthworms do not usually possess any specific gut symbiotic microbial communities [30], to some extent this apparent N2O consumption could be attributed to the parent soil-derived denitrifiers in the earthworms’ guts. This study investigated the possible implication of denitrifying microbial communities in termite guts or nest material in N2O uptake, using N. voeltzkowi species as a model. The results showed clearly that termite gut symbiotic microbiota could have a prominent role in the observed N2O uptake because dead termite mass also exhibited the same magnitude of N2O consumption as live termites. Moreover, nest material (sterilized or non-sterilized) only appeared to play a minor role in the uptake, as they only produced a small amount of N2O when incubated alone. Uptake of N2O usually depends on the prevailing concentrations of N2O and O2 in a given environment [10]. As ambient air N2O concentration is normally low (0.2e0.3 ppmv [1]), to determine N2O uptake while avoiding substrate limitation for the termites and their gut microorganisms, pure N2O was introduced into the vials for the third experimental setup. We assumed that N2O consumption by termites should be increased at high N2O concentrations in the headspace and could be due to a pathway involving either the direct reduction of N2O to NH3 (by nitrogenase) or the indirect reduction to N2 (by termite gut denitrifiers carrying nosZ genes) and subsequent biological fixation of reduced N2 by
N2-fixing symbionts (carrying nifH genes). It has been shown that N2-fixing bacteria, particularly of genus Rhizobium and gramnegative free living soil bacteria, can have the genes involved in both direct (nifH encoding nitrogenase [25,31]) and indirect (nifH and nosZ [32,33]) reduction pathways in the same genome. This enzyme-catalyzed microbial reduction of N2O and fixation of N2 can be determined by acetylene reduction assay (ARA) [34]. ARA is still widely used because it is a highly sensitive, low cost means of quantifying relative nitrogenase enzyme activity, even if, recently, it has been criticized because it may underestimate real N fixation rates [35], but the magnitude of the underestimation remains unknown [36]. The limited diffusion of C2H2 in samples under investigation may result in incomplete inhibition of nitrogenase reductase by C2H2 [36]. The effect would be very small in these experiments as termites have a small diameter which should ensure sufficient diffusion into their gut. Vessey [36] reported that one of the possible causes of underestimation could be prolonged incubation. These experiments were carried out using short incubation times. The main purpose was, however, not to quantify the absolute activity rather to get a relative indication of the prevailing process of the biological N fixation in the termite gut. The third experiment confirmed the substrate limitation because the addition of pure N2O to the vials increased the termites’ N2O consumption by a factor of 400e600 and this consumption was reduced significantly (p < 0.001) by a factor of 3e6 by the addition of C2H2 with a significant rate of N fixation (18.21 nmol C2H4 g1 dw d1) in the vials with termites, a rate very similar to that reported by Ohkuma et al. [19] for Nasutitermes takasagoensis. Sameshima-Saito et al. [37] reported similar results, showing an almost entire N2O take-up by soybean root systems inoculated with a wild symbiotic N2-fixing Bradyrhizobium japonicum bacterium with an introduced nosZ gene. Xylophagous termite species including N. voeltzkowi have been shown to have a dense, diverse community of denitrifying and N2-fixing gut symbionts which play a significant role in the N metabolism of these termites [17,18]. The C2H2-induced inhibition of N2O-reductase (nosZ) and the measurement of N2-fixing nitrogenase (nifH) activities appear to confirm the involvement of an N2O reduction process and possibly the assimilation of the N2 in the termite body tissues by a biological N fixation process. This supports the hypothesis that it is better, from a thermodynamic point of view, for termites to conserve the N derived from N2O reduction in their bodies rather than losing it into the atmosphere in gaseous form as N2 [38]. In tropical savanna and forest ecosystems, xylophagous termites represent a substantial part (41%) of termite assemblages [39] and are spreading, driven by land-use intensification [40]. Based on the trends for termite functional groups and the results of this study, it would appear that xylophagous termites could constitute a potential sink for atmospheric N2O. However, more extensive data on the in situ N2O consumption by xylophagous termites is required to support this hypothesis. Further studies should address the determination of N2O-reducing (nosZ) gene activity and the fate of nitrogen in xylophagous termites using 15N2O tracer techniques [8]. Acknowledgement This study received financial and technical support from UMR Eco&Sols (Montpellier, France), UMR BioEMCo (Bondy, France) and UMR LSTM (Montpellier, France). We are grateful to Mr. Bruno Buatois (CEFE, Montpellier, France) for his technical assistance. References [1] IPCC, Intergovernmental Panel on Climate Change, IPCC Fourth Assessment Report. Climate Change, AR4, Cambridge University Press, Cambridge, 2007.
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