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Chemical formation of hybrid di-nitrogen calls fungal codenitrification into question

Scientific Reports volume 6, Article number: 39077 (2016) | Download Citation

  • A Corrigendum to this article was published on 22 December 2017

Abstract

Removal of excess nitrogen (N) can best be achieved through denitrification processes that transform N in water and terrestrial ecosystems to di-nitrogen (N2) gas. The greenhouse gas nitrous oxide (N2O) is considered an intermediate or end-product in denitrification pathways. Both abiotic and biotic denitrification processes use a single N source to form N2O. However, N2 can be formed from two distinct N sources (known as hybrid N2) through biologically mediated processes of anammox and codenitrification. We questioned if hybrid N2 produced during fungal incubation at neutral pH could be attributed to abiotic nitrosation and if N2O was consumed during N2 formation. Experiments with gas chromatography indicated N2 was formed in the presence of live and dead fungi and in the absence of fungi, while N2O steadily increased. We used isotope pairing techniques and confirmed abiotic production of hybrid N2 under both anoxic and 20% O2 atmosphere conditions. Our findings question the assumptions that (1) N2O is an intermediate required for N2 formation, (2) production of N2 and N2O requires anaerobiosis, and (3) hybrid N2 is evidence of codenitrification and/or anammox. The N cycle framework should include abiotic production of N2.

Introduction

The nitrogen (N) removal pathway known as denitrification is typically considered a biological process, where nitrate (NO3) or nitrite (NO2) is sequentially reduced by bacteria and archaea to nitric oxide (NO), nitrous oxide (N2O) and finally inert di-nitrogen (N2)1. Incomplete denitrification results in emission of N2O, an important greenhouse gas now playing a primary role in stratospheric ozone depletion2. While prokaryotes are well known to denitrify3, new findings indicate denitrification by eukaryotes, such as fungi, may be widespread4,5,6. Unlike prokaryotes, fungi do not have the genes encoding nitrous oxide reductase, which reduces N2O to N2, so fungal denitrification terminates at N2O7,8. Denitrification occurs when a single N source is used to produce N2O, such as NO2 or NO3. Codenitrification occurs when individual atoms of the N2O or N2 molecules are derived from two distinct N sources9,10, resulting in hybrid N28. Formation of hybrid N2 is widely reported as evidence of anammox11 or codenitrification12,13,14. Previously, isotope pairing experiments revealed chemodenitrification15,16 and denitrification by the fungi Bipolaris sorokiniana used NO2 as the sole source for N2O formation16. Fungal denitrification rates of NO2 to N2O were similar under both anaerobic and microaerophilic conditions, contrary to classical denitrification, suggesting O2 may not, in this case, be a strong regulator16.

Incubation experiments with pure cultures and soils report a number of fungi may play a significant role in soil N trace gas production5,6,17,18. Fungi not only denitrify to N2O but may also codenitrify to form N28,9. When O2 is not available, some fungi (Fusarium oxysporum) reportedly co-metabolise organic forms of N to reduce NO2 or NO3 and form hybrid N27,9,10. This has been demonstrated in soils by tracing 29N2 and 30N2 following application of antibiotics to selectively inhibit bacteria or fungi12,14. Codenitrification is widely viewed as an anaerobic, enzymatically-mediated nitrosation process requiring low (<−1) formal oxidation state of the nucleophilic N8. Figure 1 illustrates how anammox, like codenitrification, also forms hybrid N2, although anammox uses two forms of inorganic N, NO2 and ammonium (NH4+)11,19.

Figure 1: Schematic of codenitrification, anammox and known chemical denitrification pathways.
Figure 1

Selected processes potentially leading to N2O and N2 formation, involved N compounds, their reaction pathways as well as their oxidation states are shown. Closed circles are biotic and open circles are abiotic reactions. Terms are defined as follows: Norg/R-NH2, monomeric organically bound N forms; NH4+, ammonium; NH3, ammonia; NH2OH, hydroxylamine; NO2, nitrite; NO3, nitrate; NO, nitric oxide; N2O, nitrous oxide; N2, molecular dinitrogen. The last process, abiotic N2 formation, was observed in this study. Figure 1 is a truncated adaptation from Butterbach-Bahl et al.23, which is licensed under a Creative Commons Attribution License 3.0 (https://creativecommons.org/licenses/by/3.0/).

The specific codenitrification pathway is unknown, but fungi reportedly reduce NO2 to NO using NO2 reductase (encoded by the nirK gene) and then reduce NO to N2O using nitric oxide reductase (P450nor)7. The role of N2O in the codenitrification process and in N2 formation is not clear20. While utilisation of N2O to form N2 has been suggested as a plausible codenitrification pathway8, reports of N2O consumption during fungal production of N2 (commonly observed during bacterial denitrification)21 are lacking. Potentially bypassing reduction of N2O to form N2, in addition to formation of hybrid N2, sets codenitrification and anammox apart from classical denitrification22.

Laboratory studies commonly report evidence of fungal denitrification or codenitrification when pure cultures are incubated under anaerobic or microaerophilic conditions with sterile media, consisting of carbon, NO2 and mineral salts5,6,9,10,16. However, reduced metals in the medium, such as Fe(II), could provide electrons required for abiotic reduction of NO2 to N2O, commonly known as chemodenitrification15,23,24. Chemodenitrification occurs through nitrosylation when reduced forms of inorganic N react with a metal centre to form N2O in the absence of oxygen15,24. Nitrosylation may also drive abiotic formation of N2, given high concentrations of metal and NO225. Wullstein and Gilmour25 mixed 10,000 ppm N as potassium nitrite (KNO2) with 5,000 ppm ferrous sulfate in an abiotic, anoxic reactor and recovered 15% of added N as N2 within 3 d. In this case, the N2 formed would have been denitrified from a single N source, KNO2. Alternatively, chemical formation of hybrid N2 through nitrosation is often ignored by biologists. Abiotic nitrosation of organic matter by NO2 in soil was first suggested by Nelson and Bremner26 when they recovered over 20% of added N (5 mmols NO2 g−1 soil) as N2 for sterile soil at neutral pH in helium (He) and heliox (20% O2, 80% He) atmospheres. The isotopic composition of N2 was not reported, but they indicated soil organic matter was an important factor26. Trimmer and Prudy27 showed that deep seawater samples amended with 15NO2 and 14NH4 produced more 29N2 when organic N (allylthiourea) was added. They suggested an alternative metabolic pathway to anammox but did not address the possibility of abiotic 29N2 formation. Babbin et al.28 also found organic N enhances anammox N2 production. A common thread among chemical, anammox and fungal denitrification studies is NO2. Thus, NO2 is a pivot-point for divergence in biological and chemical N trace gas production29 (Fig. 1).

Here, we pursue open questions raised by this early work regarding abiotic N trace gas production recently reviewed by Heil et al.30. We aimed to investigate if previously unexplored sources of N2O and N2 could be contributing to reactive N removal and if N2O was an intermediate in the abiotic N2-production pathway. We questioned whether N2 reportedly due to fungal codenitrification in pure culture experiments was formed abiotically in the presence and absence of O2, given diverse inorganic and organic sources of N. New knowledge of abiotic N2O and N2 production would advance environmental N-removal research and applications and perhaps explain some mass balance discrepancies found in isotopic pairing studies27,28.

To address these questions, we used a ubiquitous soil fungus, Bipolaris sorokiniana (Sacc.) Shoemaker [telemorph: Cochliobolus sativus (S. Ito & Kurib.) Drechsler ex Dastur], as our model. Previously, we found that B. sorokiniana used NO2 as the sole source for denitrification to N2O under anaerobic and microaerophilic conditions16. We also found NO2 as the sole source for chemodenitrification, which accounted for 6–8% of total N2O production16. These results prompted further inquiry regarding fungal and chemical trace gas production of N2. We aimed to more explicitly evaluate abiotic and biotic N2O and N2 production by comparing live fungi with fungal necromass incubated under strictly sterile conditions, with and without O2. Pure culture experiments have demonstrated fungal and chemical N2O production, but reports are lacking that indicate abiotic nitrosation of organic compounds and formation of hybrid N2. We include necromass in the design because amino acids and other nucleophilic compounds from necromass could potentially, in the absence of live fungi, react with NO2− to form N2. Further, necromass would provide more surface area for chemical decomposition of NO2 to N2O24 (Fig. 1). This would serve as a test for abiotic formation of N2O and N2 in the presence of decomposing organic material.

We aimed to assess biotic and abiotic N2O and N2 production using established microbiological laboratory incubation methods (Fig. 2). We exposed live fungi and necromass to both inorganic and organic N sources [0.25 mmol N as sodium nitrite (NaNO2) and 0.25 mmol N as glutamine (C5H10N2O3); concentration of 20 mmol L−1 each] under two O2 conditions (anaerobic and 20% O2). For each treatment, there were replicate sets where N was not added to the media (No N control). We also included sterile media-only control vessels. Oxygen status was tightly controlled with airtight laboratory incubation vessels, where headspace was filled with either helium (anaerobic) or heliox (aerobic). Accumulation of headspace O2, CO2, N2O, and N2 were measured approximately every 6 h with a customised, robotic gas chromatography system21. Headspace O2 was monitored to confirmed anaerobiosis was maintained during the 30 h incubation. Slopes of the linear increases in N2O and N2 in the headspace of each vessel were calculated to determine production rates. We used ANOVAs to test if production of N2O and N2 by live fungi was similar to necromass, and if both groups responded similarly to O2. Production rates were calculated in units of μmol N as N2O and N2 g fungal biomass−1 h−1 and in units of μmol N as N2O and N2 h−1. We performed a second incubation (illustration of this incubation design not shown) using isotope pairing techniques to determine if 29N2O or 30N2O were produced abiotically from sterile medium amended with glutamine and NO2 in the presence or absence of O2. We used equal amounts of unlabelled glutamine and 15N-labelled NO2 to achieve 0.5 mmol N and 1.0 mmol N, as well as a medium that was not amended with N.

Figure 2: Experimental set-up to test how live fungi, necromass and media only incubated in an anoxic and 20% O2 atmosphere affects production of N2O, CO2, O2, and N2 following addition of both organic and inorganic forms of N to pure cultures under aseptic conditions.
Figure 2

Terms are defined as: Glut, glutamine; NaNO2, sodium nitrite; He, helium; HeOx heliox.

Results and Discussion

We found both fungal states (live fungi and necromass) produced N2O and N2 but only for those samples amended with N. Linear production of N2O and N2, following N amendment, occurred quickly under completely anaerobic conditions and in a 20% O2 atmosphere (Fig. 3). Average [(standard deviation (SD)] rate of N2O produced by necromass was 0.012 (<0.001) μmol N2O g biomass−1 h−1 under both aerobic and anaerobic conditions. Average N2O production by live fungi was 0.017 (<0.001) μmol N2O g biomass−1 h−1 and 0.028 (0.003) μmol N2O g biomass−1 h−1, respectively, under aerobic and anaerobic conditions (Fig. 3a). Rates of N2O production were significantly greater for live fungi incubated anaerobically (O2 × fungal state interaction; p < 0.001), indicating biological production of N2O in the absence of O2. In a 20% O2 atmosphere, rates of N2O production were similar to necromass. Our previous study indicated B. sorokiniana produced N2O at similar rates when incubated anaerobically and in a 0.4% O2 atmosphere16. Here, we used 20% O2, and we found no evidence of microbial denitrification under these high-O2 conditions.

Figure 3
Figure 3

Kinetics of (a) N2O and (b) N2 production per g biomass over time following N addition at each time point as gases in the headspace accumulated. Average for each treatment are slightly staggered in time due to robotized measurement system, which is why comparisons were based on slopes over the entire incubation. Production rates of N2O and N2 were strongly affected by O2 status [O2 × fungal state (live or dead); p < 0.001] for those samples amended with N. Boxes represent average data; error bars, s.d.; n = 4.

Figure 4a provides a basis of comparison for N2O recovered in the headspace of sterile media using comparable units (μmol N2O h−1). These data were not normalised per g of fungal biomass but illustrate accumulation of N2O in sterile medium relative to necromass and live fungi. Nitrous oxide production rates were greater for necromass, as compared to media only (Fig. 4a). Necromass produced an average of 0.0034 μmol N2O h−1 at both O2 levels, as compared to 0.0011 μmol N2O h−1 for sterile media. Greater N2O for necromass suggests the presence of decaying biomass provided greater surface area for chemodenitrification24 (Fig. 4a).

Figure 4
Figure 4

Abiotic and biotic contributions to rates of (a) N2O and (b) N2 accumulated per hour following N addition under both aerobic and anaerobic conditions for sterile medium, necromass and live fungi. Boxes represent the 90th percentile data; error bars, s.d.; n = 4. Median values are the lines horizontally bisecting each box.

Rates of fungal N2O production observed here are lower than other reports in the literature. Maeda et al.6 found fungi carrying the nirK gene produced from 0.1 to 3.2 μmol N2O g biomass−1 h−1, and Rohe et al.31 reported fungal N2O production rates from 0.05 to 14 μmols N2O h−1. In both cases, denitrification varied with fungal species and N amendment. Here, B. sorokiniana does not carry the nirK gene, which may influence NO2 reduction to NO. Data on chemical formation of N2O under oxic (20% O2) conditions are lacking, so these first results challenge the paradigm that chemodenitrification requires anoxia15. Contrary to chemostat studies, we included fungal necromass, which contributed to abiotic N2O. Results call into question if all N2O produced in pure culture experiments6,18,31 is enzymatically mediated if cultures include live and dead fungal or bacterial biomass.

We found no evidence of biological N2 production in the presence or absence of necromass (Fig. 3b). Lowest rates of N2 production were observed for live fungi incubated in a 20% O2 atmosphere (O2 × fungal state interaction; p < 0.001). Rates of N2 produced in the headspace of live fungi were 0.243 (0.039) μmol N2 g biomass−1 h−1 and 0.424 (0.049) μmol N2 g biomass−1 h−1 under aerobic and anaerobic conditions, respectively. Rates of N2 produced in the headspace of necromass were 0.415 (0.041) μmol N2 g biomass−1 h−1 and 0.356 (0.047) μmol N2 g biomass−1 h−1 under aerobic and anaerobic conditions, respectively. Our results indicate there is a strong abiotic component to measured rates of N2 production and challenge the assumption that fungal N2 production7 requires anoxic or microaerophilic conditions8.

Most of the N2 accumulated in the headspace for live B. sorokiniana was commensurately found in the headspace of B. sorokiniana necromass (Fig. 3b), which points to chemical N2 formation. We show chemical production of N2 (μmol h−1) for sterile medium relative to necromass and live fungi at both O2 levels in Fig. 4b. Overall average rates of N2 formation were 0.12 (0.039) μmol N2 h−1 for sterile medium, 0.11 (0.024) μmol N2 h−1 for necromass, and 0.08 (0.018) μmol N2 h−1 for live fungi. Others have reported much higher rates of anoxic chemical N2 formation (7–32 μmol N2 h−1) at neutral pH when high concentrations of reduced metals were reacted with high concentrations of NO225 and when high NO2 solutions (20 M) were added to autoclaved soil26, but these did not report if hybrid N2 was formed or not. Comparable rates of abiotic N2 production for necromass and sterile medium caused us to question if abiotic N2, like N2O, could result from nitrosylation of inorganic N only or from nitrosation to form hybrid N2. Organic N has been found to increase biological production of 29N2 in oceanic studies32, but our results (Fig. 4b) indicated there may also be abiotic processes that contribute strongly to 29N2 production. Consequently, we determined if both inorganic and organic N were used to produce N2 abiotically in a separate, isotope pairing experiment.

The headspace above sterile media incubated under oxic and anoxic conditions with 15N-labelled NaNO2 and unlabelled C5H10N2O3 indicated abiotic, formation of hybrid N2. Almost all (99.9%) of the N2 produced was 29N2 (Table 1). The 29N2 production was proportional to the mass of added N and approximately 3–4% of the total N added was transformed to 29N2 under both oxic and anoxic conditions and at both levels of NO2 addition. Results demonstrate hybrid formation of N2 is not necessarily enzymatically mediated and does not required anoxia. Formation of abiotic, hybrid N2 by two distinct inorganic N molecules has not been ruled out here. Previous bodies of work that use formation of 29N2 as evidence of anammox and/or codenitrification12,14,32,33 need to be reviewed with respect to abiotic N2 production.

Table 1: Average (SD) of 29N2 and 30N2 recovered in the headspace following aerobic and anaerobic incubation of sterile media at three levels of N addition, where added N comprised 50% N from unlabelled C5H10N2O3 and 50% N from labelled 15N-NaNO2 (n = 5).

Linear increases in cumulative N2O and N2 over time under aerobic and anaerobic conditions for both live fungi and necromass are shown in Fig. 3. These data suggest that N2O was not consumed during the incubation and both gases were produced independently. This finding contrasts with bacterial denitrification22, where a sharp rise in microbial production of N2O is followed by a rise in biological reduction of N2O to N2, and a drop in cumulative N2O production21. Bipolaris sorokiniana denitrifies NO2 only and does not use glutamine to form N2O16. If the pathway to N2 were through the intermediate N2O, we would expect not only N2O consumption but also accumulation of 30N2 in the isotopic pairing experiment. Like anammox, our abiotic N2 production results indicate N2O formation was bypassed in the pathway to N2 (Fig. 1).

We show compelling evidence that formation of 29N2 does not result from solely biotic nitrosation but also abiotic nitrosation. Abiotic nitrosation occurred both in the presence and absence of O2, and abiotic N2 was formed exclusively through hybridisation of inorganic and organic N sources and did not require the N2O intermediary. Experimental protocols, including N concentrations, were in accordance with fungal denitrification6,16 and codenitrification10 studies but in the absence of soils and sediments. In soil, NO2− accumulates when excess free NH3 inhibits bacterial NO2 oxidation, which is often a consequence of urea hydrolysis30. Venterea et al.34 recovered 3–60% of added urea N as NO2, suggesting high soil NO2− is likely following urea addition (depending upon conditions and application rate). Based on these data34, a 1-kg bovine urine addition to soil (N concentration of 0.4 mmol kg−1)35 could result in up to 0.24 mmol NO2. Here, we added 0.25 mmol NO2 to 10 ml of fungal culture, which is on the high end of this scale. While we would expect abiotic formation of hybrid N2 via nitrosation of organic N to be more likely in grazed or fertilised agroecosystems, further N2 investigations with soil and at lower NO2 levels are needed to bridge the gap between pure culture and environmental applications.

Other areas where NO2 could accumulate include laboratory incubations where antibiotic inhibitors are added to soil and used to partition N2 and N2O production into fungal and bacterial contributions12,14. These data may be subject to artefacts if antibiotics repress NO2 oxidation, leading to NO2 accumulation and potential abiotic formation of N2 and/or N2O. Our report also calls into question if codenitrification alone accounts for N2 and N2O emissions following high doses of N as urea (approximately 9 mmol g−1 soil)13 and/or NO2 (166 mmol L−1)10. If abiotic, hybrid N2 is formed under these conditions, our understanding of soil fungal codenitrification and N trace gas emissions would need to be re-examined. We offer new insight into chemodenitrification of NO2 to N2 and present an alternative ‘N2O bypass pathway’. Finally, we conclude the potential for aerobic, abiotic removal of excess reactive N in terrestrial and aquatic ecosystems presents environmental mitigation research opportunities, particularly where transitory or chronic NO2 accumulation occurs.

Methods

Fungal incubations were conducted as described in Phillips et al. (2016) with the same culture ICMP 6809 isolated as a pathogen of Hordeum distichon in New Zealand. (https://scd.landcareresearch.co.nz/Specimen/ICMP_6809) GenBank: KU194490. We used laboratory incubations and controlled O2 status to evaluate if rates of N2O, N2 and CO2 production rates vary with aerobic conditions, as commonly reported for bacteria20. Experimental design was similar to previous work16, with 4 replicates for each treatment (N with and without O2; no N with or without O2). Cultures were grown in medium similar to other fungal denitrification studies9,10,31, consisting of 1% glucose, 0.2% peptone and inorganic salts5,36. The only N source in this medium used for fungal growth was peptone. One day prior to the experiment, the growth medium was washed off cultures and replaced with the same medium that was identical except it was free of peptone and so contained no N. Using the peptone-free medium herein, two amendment solutions were prepared for fungal inoculation: (a) media without N and (b) 0.25 mmol N as NaNO2 and 0.25 mmol N as C5H10N2O3. Nitrogen concentration was 0.04 mmol N L−1. Necromass was obtained by heating live B. sorokiniana at 60 °C for 48 h. Cell death was confirmed by (a) microscopy and (b) lack of CO2 respiration over a 24 h period under aerobic and anaerobic conditions. Approximately 10 ml live fungi or fungal necromass were blindly pipetted by a second independent scientist into 0.125 L serum bottles that were then amended with either media that included 0.25 mmol N as NaNO2 and 0.25 mmol N as C5H10N2O3 or media without N and mixed gently. Additional bottles containing sterile media solutions (with and without N) only and without fungi were also prepared. Oxygen status was controlled with airtight laboratory incubation vessels for all necromass, live fungi and media only samples. Headspace of each sample was evacuated and filled with either He (anaerobic) or heliox [aerobic (80% He, 20% O2)] within 2 hr following inoculation18,20. Headspace gases were quantified approximately every 6 h at 19 °C using a robotic gas chromatograph (GC) fitted with electron capture and thermal conductivity detectors according to McMillan et al.21. Helium or heliox blank standards were included in each GC run. Cumulative production rates were calculated as slopes of the masses (μmol) of N2O or N2 measured over time (h) per g of fungal biomass. Vessels incubated in He remained anoxic with the exception of necromass, where we observed 0.05% O2. Vessels incubated in heliox remained above 19% O2 with the exception of live fungi incubated without N, where 5% of the headspace O2 was consumed. As reported previously, addition of NO2 inhibited O2 consumption and CO2 respiration by live fungi16. Fungal biomass was determined as the difference in mass with and without media after air-drying each vessel post-incubation16. For comparisons with sterile media, rates were also calculated as slopes of the masses (μmol) of N2O and N2 measured over time. Sterility was maintained and conditions remained constant, including pH (6.2–6.9). Sterility was tested at the end of gas sampling by pipetting 200 μL medium used in the experiment onto blood agar plates and incubating under aerobic, anaerobic (AnaeroGenTM, Thermo Scientific), and enriched CO2 (3.5–9%; CO2GenTM, Thermo Scientific) conditions. Further, medium was also pipetted onto yeast nutrient agar, brain heart infusion agar, and potato dextrose agar plates under aerobic conditions. No evidence of fungal or bacterial growth was observed following 3, 7, and 10 d incubations at 26 °C. Data were analysed to test for effects of O2 and live fungi on either N2O or N2 production rates with a generalised linear model. We analysed only those samples amended with N because samples without N did not produce N2O or N2. Log transformations were employed when data did not meet the assumptions of normality. Treatments variances met assumptions of homoscedasticity. All interactions were tested and remained in the model if significant.

Di-nitrogen isotopes were evaluated in a separate experiment to assess sources of N used in abiotic production of N2 by adding Na15NO2 and C5H10N2O3 to sterile medium at neutral pH, as described above. We aimed to determine if N2 would be produced through combination of 15NO2 only (thus forming 30N2) or through combination of both C5H10N2O3 and 15NO2 (thus forming the hybrid 29N2) for N-enriched medium relative to no-N medium only. In this experiment, we used five replicates at three levels of N: (a) no-N, (b) 0.25 mmol NaNO2 and 0.25 mmol C5H10N2O3, and (c) 0.5 mmol NaNO2 and 0.5 mmol C5H10N2O3. We used the same concentration of N in each N treatment but doubled the mass of N added. Vials were prepared under aerobic and anaerobic conditions as described previously and accompanied by He blanks. The aerobic experiment was conducted separately from the anaerobic experiment, which obviated testing for effects of O2 on masses of 29N2 and 30N2. Headspace 29N2 and 30N2 were measured on a continuous-flow isotope ratio mass spectrometer (Thermo Finnigan Delta V, Thermo Scientific) in line with an automated gas bench interface (Thermo Gas Bench II). Precision of the isotopic analysis was <0.001atom%.

Additional Information

How to cite this article: Phillips, R. L. et al. Chemical formation of hybrid di-nitrogen calls fungal codenitrification into question. Sci. Rep. 6, 39077; doi: 10.1038/srep39077 (2016).

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Change history

  • Updated online 22 December 2017

    A correction has been published and is appended to both the HTML and PDF versions of this paper. The error has not been fixed in the paper.

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Acknowledgements

The authors thank the team at Landcare Research in New Zealand for supporting this idea. This work was partially funded by a USDA-NIFR grant [2014-67019-21614]; New Zealand’s Ministry of Business Innovation and Employment, Royal Society of New Zealand International Travel Programme; and New Zealand Ecosystems and Global Change Fund. The authors are grateful to Veronica Rollinson, Sujatha Senanayake, Duckchul Park, and Megan Peterson for technical assistance and Bill Schlesinger, Iris Anderson, Neha Jha, Mikki Eken, and Leah Kearns for their comments, support and editorial reviews.

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Affiliations

  1. Landcare Research, Gerald Street, Lincoln, New Zealand

    • Rebecca L. Phillips
    • , Andrew M. S. McMillan
    • , Gwen Grelet
    • , Bevan S. Weir
    •  & Thilak Palmada
  2. Dept. of Biological Sciences, Virginia Institute of Marine Science, Gloucester Point, Virginia, USA

    • Bongkeun Song
  3. Dept. of Marine Sciences, University of Connecticut, Groton, Connecticut, USA

    • Craig Tobias

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Contributions

R.L.P., B.S. and G.G. designed the experiments, T.P. performed lab experiments, A.M.S.M. analysed chromatographic data, C.T. performed isotope pairing experiments, B.S.W. and G.G. cultured, and verified fungal isolates, and R.L.P. and B.S. wrote the manuscript.

Competing interests

The authors declare no competing financial interests.

Corresponding author

Correspondence to Rebecca L. Phillips.

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https://doi.org/10.1038/srep39077

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